Cardiovascular Research

Cardiovascular Research 2004 62(1):63-73; doi:10.1016/j.cardiores.2003.12.031
© 2004 by European Society of Cardiology

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Copyright © 2004, European Society of Cardiology

Reduced synchrony of Ca²⁺ release with loss of T-tubules—a comparison to Ca²⁺ release in human failing cardiomyocytes

William E Louch^a, Virginie Bito^a, Frank R Heinzel^a,b, Regina Macianskiene^c, Johan Vanhaecke^a, Willem Flameng^c, Kanigula Mubagwa^c and Karin R Sipido^*,a

^aLaboratory of Experimental Cardiology, University of Leuven, Campus Gasthuisberg O/N 7th floor, Herestraat 49, B-3000 Leuven, Belgium
^bInstitute of Pathophysiology, University of Essen, Essen, Germany
^cCenter for Experimental Surgery, University of Leuven, Leuven, Belgium

^* Corresponding author. Tel.: +32-16-347153; fax: +32-16-345844. Email address: karin.sipido{at}med.kuleuven.ac.be

Received 10 September 2003; revised 10 December 2003; accepted 23 December 2003

	Abstract

Top Abstract 1. Introduction 2. Methods 3. Results 4. Discussion References

 
Objectives: During cardiac excitation–contraction coupling, Ca²⁺ release from the sarcoplasmic reticulum (SR) occurs at the junctional complex with the T-tubules, containing the L-type Ca²⁺ channels. A partial loss of T-tubules has been described in myocytes from failing canine and human hearts. We examined how graded reduction of T-tubule density would affect the synchrony of Ca²⁺ release. Methods: Adult pig ventricular myocytes were isolated and cultured for 24 and 72 h. T-tubules, visualized with di-8-ANEPPS, and [Ca²⁺]_i transients (Fluo-3) were recorded during confocal line scan imaging. Results: Cultured cardiomyocytes exhibited a progressive reduction in T-tubule density. [Ca²⁺]_i transients showed small areas of delayed Ca²⁺ release which gradually increased in number and size with loss of T-tubules. Local [Ca²⁺]_i transients in the delayed regions were reduced. Due to these changes, loss of T-tubules resulted in an overall slowing of the rise of [Ca²⁺] along the entire line scan and transient magnitude tended to be reduced, but there was no change in SR Ca²⁺ content. Human myocytes isolated from failing hearts had a T-tubule density comparable to that of freshly isolated pig myocytes. The size, but not the number, of delayed release areas tended to be larger. The overall rate of rise of [Ca²⁺]_i was significantly faster than in pig myocytes with low T-tubule density. Conclusions: Loss of T-tubules reduces the synchrony of SR Ca²⁺ release. This could contribute to reduced efficiency of excitation–contraction coupling in heart failure, though dyssynchrony in human failing cells appears to be modest.

KEYWORDS Heart failure; Calcium; Cell culture; E–C coupling; Myocytes; SR (function); Sarcolemma; T-tubules; Pig; Confocal microscopy

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This article is referred to in the Editorial by S. Hatem (pages 1–3) in this issue.

	1. Introduction

Top Abstract 1. Introduction 2. Methods 3. Results 4. Discussion References

 
In adult ventricular myocytes, contraction is initiated when Ca²⁺ entry across the sarcolemma triggers release of a much larger amount of Ca²⁺ from the sarcoplasmic reticulum (SR) (for review see Refs. [1–3]). In ventricular myocytes, this process occurs at the junction of the SR and the T-tubular system, where L-type Ca²⁺ channels face SR Ca²⁺ release channels [4]. Elementary Ca²⁺ release events can be identified as Ca²⁺ sparks during confocal line scan images when the probability of opening of the L-type Ca²⁺ channel is reduced [5,6]; these events are localized near the T-tubules [7]. In ventricular myocytes with a dense T-tubular network, SR Ca²⁺ release in near physiological conditions occurs almost simultaneously throughout the cell [5,8–10].

In contrast, in myocytes devoid of T-tubules, such as atrial cells, neonatal myocytes, and Purkinje cells, SR Ca²⁺ release is first triggered under the sarcolemma, with subsequent propagation to the center of the cell, resulting in a characteristic ‘horseshoe’ appearance of the [Ca²⁺]_i transient in confocal line scan images [11–14]. Acute detubulation of adult rat ventricular myocytes produces a similar pattern [15]. In guinea-pig ventricular myocytes which had lost T-tubules following culture, a wavelike propagation of the [Ca²⁺]_i transient was observed [8]. It can then be postulated that a more subtle reduction of T-tubule density will result in intermediate and less pronounced dyssynchrony of SR Ca²⁺ release.

Loss of T-tubule density by about 50% has been reported in the dog with tachycardia-induced heart failure [16,17], and could contribute to the reduced SR Ca²⁺ release in this model. In human heart failure, it remains at present uncertain whether there is a reduction of T-tubules. A low density of somewhat dilated T-tubules has been described in fixed tissues [18], but preliminary studies in isolated living myocytes gave conflicting data [19,20]. In the present study, we have examined more quantitatively the effect of reducing T-tubule density on the spatial characteristics of the [Ca²⁺]_i transient. Since T-tubule attenuation in culture is time-dependent, we examined different timepoints to determine dose-dependent alterations in SR Ca²⁺ release. The results were compared to the spatial characteristics of SR Ca²⁺ release in myocytes from failing human hearts.

	2. Methods

Top Abstract 1. Introduction 2. Methods 3. Results 4. Discussion References

 
2.1. Myocyte isolation and culture
Domestic pigs (aged 3 to 6 months, 40 to 50 kg, n=17) were treated in accordance with the Guide for the Care and Use of Laboratory Animals (N.I.H.), and experimental protocols were approved by the in-house ethics committee. Single ventricular myocytes were isolated as described previously [21]. Freshly isolated cells were studied within 8 h of isolation. For culture, cells were concentrated by low-speed centrifugation, washed, and suspended in supplemented Medium 199 (containing bovine albumin 2 mg/ml, ascorbic acid 10 µM, taurine 5 mM, creatine 5 mM, penicillin 50 IU, streptomycin 50 µg/ml) on 12-well culture plates (Becton Dickinson Laboratories, Franklin Lakes, NJ). Myocytes were stored in the incubator until the time of examination (5% CO₂ incubator, 37 °C).

Human ventricular myocytes were prepared from the explanted hearts at the time of transplantation as previously described [22]. The procedure was approved by the Ethical Committee of the University Hospital and conforms to the principles outlined in the Declaration of Helsinki. For the present study myocytes from six patients were examined (age 47±17 years, three patients with ischemic cardiomyopathy, two with idiopathic dilated cardiomyopathy, and one with congenital valvular disease).

2.2. Experimental protocols
In all experiments, myocytes were plated in an open-perfusion chamber mounted on the stage of an inverted microscope. For whole-cell voltage clamp the pipette solution contained (in mM) 120 K-aspartate, 20 KCl, 10 K-HEPES, 5 MgATP, 0.5 MgCl₂, 10 NaCl, and 0.05 K₅-fluo-3 (pH 7.20). The external solution was a normal Tyrode's solution containing (in mM) 130 NaCl, 5.4 KCl, 11.8 Na-HEPES, 0.5 MgCl₂, 1.8 CaCl₂, and 10 glucose (pH 7.35). Membrane potential and current recordings were made with an Axoclamp 2A amplifier (Axon Instruments, Foster City, CA) and pCLAMP software (Axon Instruments). [Ca²⁺]_i transients were studied during a 150-ms depolarizing step from –70 to 0 mV (repeated at 1 Hz, steady state). For measuring the L-type Ca²⁺ current, a prepulse of –40 mV for 450 ms was applied. To estimate SR content, we used a fast application of 10 mM caffeine (7 s) following a series of conditioning steps at 1 Hz. Cellular capacitance was measured during a voltage step from –70 to –80 mV. Action potentials were elicited by a short depolarizing current pulse.

To visualize the T-tubular network, cells were incubated with di-8-ANEPPS (10 µM, Molecular Probes, Eugene, OR) for 10 min at room temperature, followed by 10 min of washout. To visualize cell volume, myocytes were loaded with calcein-AM ester (5 µM, Molecular Probes) for 30 min, followed by 30 min of washout [23].

2.3. Confocal fluorescence imaging
Confocal fluorescence imaging was performed using the LSM 510 point scanning system (Zeiss GmbH, Jena, Germany); settings and properties were as described previously [10]. Z-stacks of XY (512 x 512 pixels) images were recorded with a spacing between adjacent slices of 1.2 µm to prevent fluorescence interference from out-of-focus regions of the cell. For recording [Ca²⁺]_i, a 512-pixel scan line was drawn along the longitudinal axis of the cell, and scanned every 1.5 ms. Sequential line scans were stacked and displayed as two-dimensional images.

2.4. Data analysis
Image analysis was performed with Scion Image (Scion NIH), Microsoft Excel, Origin (Microcal Software, Northampton, MA), and SigmaPlot (Jandel Scientific, San Rafael, CA) software. T-tubular density was calculated for three consecutive XY slices from the Z-stacks [10], and expressed as the T-index, i.e., the percentage of the surface area of the XY-plane occupied by T-tubular structures, excluding the external sarcolemma.

For calculation of cell volume and external surface area, each XY slice of the Z-stack was thresholded by the Otsu method [24] to distinguish the calcein signal from background. Cell volume was calculated by multiplying the total number of significant pixels from all slices by the pixel size and the between-slice interval (1.2 µm); perimeter measurements were summated for all slices and multiplied by the between-slice interval to estimate external surface area. Both volume and external surface area values were scaled by a correction factor (1.39), which accounted for overestimation of the cell size in the Z direction due to spherical aberration [23]. This correction factor was calculated using Z-stacks of fluorescent beads with a diameter 15 µm.

Line scan images of [Ca²⁺]_i transients were background subtracted, averaged for five consecutive steady-state transients, and divided by the fluorescence intensity at rest, F₀, to obtain a normalized image (F/F₀). The onset of depolarization was chosen as time zero, corresponding closely to the onset of the [Ca²⁺]_i transient in regions of early release. Regions for which mean fluorescence intensity during the first 200 ms did not exceed the half-maximal fluorescence of the entire line scan (F₅₀) were defined as artifacts and excluded. Regions of delayed Ca²⁺ release were defined as those regions where the local fluorescence intensity at 24 ms was still below F₅₀. Only early and delayed regions which were 2 µm were considered. Local transients in all early and delayed regions were analyzed by selecting a 2-µm section near the center of each region.

2.5. Statistics
Data are presented as mean±S.E.M. Significance of differences between population means was tested with a Student's t test or two-way ANOVA, and post-hoc Bonferroni test, using SigmaStat software (Jandel Scientific).

	3. Results

Top Abstract 1. Introduction 2. Methods 3. Results 4. Discussion References

 
3.1. Changes in the T-tubular network during cell culture
Ventricular myocytes cultured for 24 or 72 h showed slight rounding of the edges, but otherwise remained similar in appearance to freshly isolated cells (Fig. 1A–C, left panels). However, after staining with di-8-ANEPPS, images taken from the center of these myocytes (Fig. 1A–C, right panels) showed that T-tubule density was markedly and progressively reduced during culture. Quantification of the T-tubule signal showed a reduction in T-tubule density to 31% of control levels following 24-h culture, and a further significant reduction to 14% of control levels at 72-h culture (Fig. 1D). Whole-cell capacitance was also reduced following culture (Fig. 1E).

View larger version (50K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 1 T-tubule density was progressively reduced during culture. (A–C) Three different myocytes from the indicated experimental time points (control=freshly isolated). Left panels show light images; right panels are confocal images from the center of the same myocytes stained with di-8-ANEPPS (scale bar=10 µm). (D) Mean T-tubule density (control, ncells=10; 24 h, ncells=10; 72 h, ncells=10). (E) Whole-cell capacitance measurements (control, ncells=42; 24 h, ncells=53; 72 h, ncells=44; * denotes P<0.05 vs. control; # denotes P<0.05 vs. 24 h). (F) S/V ratios based on capacitance and volume measurements, (control, ncells=18; 24 h, ncells=24; 72 h, ncells=19; * denotes P<0.05 vs. control. (G) Relation between external surface area and volume.

 
The above measurements can be used to estimate the percentage of total surface area occupied by the T-tubules. As the total surface area at 24 h equals the total surface area in control minus the fraction of T-tubules lost in 24 h, we can calculate that T-tubules account for 30% of the total surface area in freshly isolated pig cells.

We also examined surface-to-volume (S/V) relationships by first measuring the volume of a calcein-loaded myocyte, and then the cell capacitance during voltage clamp. Cell volumes were unchanged from control values (41±2 pl, n_cells=39) during culture (at 24 h, 42±3 pl, n_cells=40; at 72 h, 45±3 pl, n_cells=40, P=NS); S/V ratios were reduced (Fig. 1F).

The external surface area of calcein-loaded myocytes, excluding T-tubules, was positively correlated with volume measurements in freshly isolated myocytes (Fig. 1F). We compared the value for external surface area with the value of total surface membrane derived from the capacitance in paired measurements, assuming a specific membrane capacitance of 1 µF/µm². Based on this comparison, the external surface area accounted for 75±7% (n_cells=18) of total surface area, in general agreement with our previous estimation that T-tubules account for approximately 30% of the total sarcolemma.

3.2 Is synchrony of SR Ca²⁺ release reduced with loss of T-tubules?
Small areas of delayed SR Ca²⁺ release were observed in freshly isolated cells, but these became much more prominent at 24 and at 72 h of culture (Fig. 2). In cells cultured for 72 h, maximizing phosphorylation of Ca²⁺ channels with isoproterenol (3 µM, n_cells=5) or forskolin (10 µM, n_cells=3) did not synchronize the [Ca²⁺]_i transient; a fast application of 10 mM caffeine induced Ca²⁺ release along the entire scan line, indicating the presence of functional ryanodine receptors (n_cells=6). Thus, increased dyssynchrony in Ca²⁺ release during culture likely resulted from loss of T-tubules.

View larger version (70K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 2 Dyssynchrony in SR Ca2+ release increased during culture. Longitudinal line scans show small areas of delayed Ca2+ release in freshly isolated cells (A), and larger and more numerous areas following 24 h (B) and 72 h (C) of culture, although regions of early release were still observed.

 
To quantify the degree of spatial inhomogeneity, we examined spatial profiles of line scans at several time points (Fig. 3A–C). In control cells, several small areas were below F₅₀ at 24 ms, whereas the number and width of such delayed areas progressively increased after 24 and 72 h of culture. Of the total length of the line scan, delayed areas occupied 37±7% (n_cells=15) in control, 65±8% (P<0.05, n_cells=12) at 24 h and 69±6% (P<0.05, n_cells=13) at 72 h. The mean width of delayed release regions was also increased (Fig. 3D). The larger fraction of delayed areas is expected to slow down the overall increase of [Ca²⁺]_i across the line scan. This was examined by calculating the percentage of the line scan with F>F₅₀ as a function of time. Plots of this relationship in individual cells (Fig. 3E–G) and mean data (Fig. 3H) show that this percentage increased much more gradually at 24 h and even more so at 72 h. The time for half of the line to reach F₅₀ was 19±2 ms in control, 27±3 ms at 24 h and 29±2 ms at 72 h.

View larger version (37K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 3 Cell culture increased the fraction of delayed SR Ca2+ release. (A–C) Spatial profiles of line scans at various time points. Regions of delayed release were defined as those with F<F50 at 24 ms. (D) Mean width of delayed regions at 24 ms (control, n=23; 24 h, n=31; 72 h, n=24). (E–H) Plots of the fraction of the line scan with F>F50 vs. time (E, F, G, individual cells; H, mean data). * denotes P<0.05.

 
We examined the properties of the local [Ca²⁺]_i transients in regions of early and delayed release. Transients from delayed regions were smaller and slower than those from early regions, and this became progressively more pronounced at 24 and 72 h (Fig. 4). In large regions of delayed release, we could measure the speed of propagation of the transient front. This value (98±9 µm/s, n_cells=5) was comparable to the propagation speed of spontaneous Ca²⁺ waves observed in Ca²⁺ overloaded cells (93±7 µm/s, n_cells=6), and close to previously reported values for wave propagation (e.g. [25]). Since Ca²⁺ waves are thought to result from a cascade of CICR [25], these results suggest the same mechanism underlies propagation of the Ca²⁺ transient in regions of delayed release.

View larger version (32K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 4 Local transients became slower and smaller during culture. (A–C) Representative local transients, as obtained from the images in Fig. 3. (D) Mean data for [Ca2+]i transient magnitudes (early: control, n=27; 24 h, n=40; 72 h, n=31; delayed: control, n=23; 24 h, n=31; 72 h, n=24). (E) Mean data for time-to-F50. (# denotes P<0.05 vs. control values).

 
3.3 Does increased dyssynchrony of SR Ca²⁺ release alter the kinetics and magnitude of the overall transient?
Based on our observations of local [Ca²⁺]_i transients, an increased amount of delayed Ca²⁺ release would be expected to significantly alter the overall [Ca²⁺]_i transient. Representative recordings (Fig. 5A) show that transient magnitudes tended to be reduced in cultured cells, although this effect was not statistically significant (Fig. 5B). However, total transient TTF₅₀ was increased to 122% and 130% of control values following 24 and 72 h of culture, respectively (Fig. 5C). This slower rate of rise of [Ca²⁺]_i likely resulted from an increased proportion of delayed release regions, and larger TTF₅₀ values in both early and delayed regions following culture.

View larger version (18K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 5 Overall [Ca2+]i transients were smaller and slower following culture. (A) Representative examples of spatially averaged [Ca2+]i transients. (B) Mean magnitude of [Ca2+]i transients. (C) Mean time-to-F50 (control, ncells=15; 24 h, ncells=12; 72 h, ncells=13); in the same cells time from onset to half-relaxation of the [Ca2+]i transient was 214±5 ms in control vs. 278±7ms at 24 h, and 289±20 ms at 72 h (P<0.05).

 
3.4 Are changes in SR Ca²⁺ content and Ca²⁺ currents responsible for alterations in local [Ca²⁺]_i transients?
While loss of T-tubules could explain the increased proportion and slower kinetics of delayed regions, it does not explain changes in transients from early release regions. SR Ca²⁺ content, measured as the total charge extruded by the Na/Ca exchanger during caffeine application, was comparable for control (92±23 pC, n_cells=9), 24 h (105±16 pC, n_cells=14), and 72 h of culture (83±14 pC, n_cells=11). The amplitude of the caffeine-induced [Ca²⁺] _i transient was also not significantly different (F/F₀ of control 3.26±0.21, n_cells=7; at 24 h 2.82±0.19, n_cells=13; at 72 h 2.59±0.43, n_cells=11), though relaxation was markedly slower after 24 and 72 h, consistent with loss of NCX in the T-tubules [15,26,27].

Reductions in L-type Ca²⁺ current, I_CaL, would be expected in cultured cells as Ca²⁺ channels are lost along with T-tubules. However, additional loss of L-type channels in remaining T-tubules could contribute to alterations in transients in regions of early Ca²⁺ release. Currents and associated [Ca²⁺]_i transients were examined using voltage steps from –40 to 0 mV. Elicited [Ca²⁺]_i transients were markedly reduced in magnitude following culture (at 24 h to 78±4%, n_cells=7; at 72 h to 81±3% of control, n_cells=8; P<0.05) as was peak I_CaL (at 24 h to 53±6%, at 72 h to 46±10% of control, P<0.05). The current–voltage relationship for I_CaL was markedly reduced (Fig. 6A). If L-type Ca²⁺ channels are uniformly distributed in the sarcolemma, the decrease observed in I_CaL would be eliminated when these values were normalized to capacitance. However, Fig. 6B shows that a modest decrease in current density was still apparent in cultured cells. This disproportionate loss can be consistent with preferential localization of Ca²⁺ channels in T-tubules. Based on acute detubulation experiments in rat ventricular myocytes, it has been previously reported that 75% of Ca²⁺ channels are localized in the T-tubules [27]. Using this value and our calculation that T-tubules account for 30% of the total surface area in freshly isolated pig cells, we predicted that I_CaL density should be decreased to 39% of control values following 24 h of culture. Surprisingly, we observed only a 26% reduction. This discrepancy could result from a compensatory increase in the number and/or activity of Ca²⁺ channels in the T-tubules that remain during culture. Alternatively, our estimation that 75% of L-type channels are localized in the T-tubules may not be valid in pig ventricular myocytes. Thus, it is difficult to ascertain whether alterations in [Ca²⁺]_i transients in regions of early release are related to reductions of I_CaL.

View larger version (19K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 6 The L-type Ca2+ current, ICaL, was reduced during culture. (A) Nifedipine-sensitive current, not normalized to cell capacitance (control, ncells=13; 24 h, ncells=13; 72 h, ncells=11). (B) ICaL density after correction for capacitance. * denotes P<0.05.

 
Since I_CaL is an important determinant of action potential duration (APD), we also examined the effects of culture on action potential configuration. Resting membrane potential was gradually decreased during culture (Fig. 7A). As well, the plateau phase of the action potential was progressively shortened in cultured cells, while total repolarization time was increased (Fig. 7B). Reductions in I_CaL likely explain the abbreviation of APD₅₀, but not the prolongation of APD₉₀ which could result from depression of I_K1. Indeed, the outward current at potentials between –20 and –60 mV was significantly reduced (Fig. 7C). It is likely that alterations in [Ca²⁺]_i-dependent currents, e.g. NCX current with the prolonged [Ca²⁺]_i transient (Fig. 5A), also contribute to the changes in action potential configuration.

View larger version (19K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 7 Changes in action potential configuration. (A) Representative recordings of action potentials. (B) Action potential duration at 50% repolarization (APD50) and 90% repolarization (APD90) (control, ncells=13; 24 h, ncells=10; 72 h, ncells=7). (C) Currents at the end of the 150-ms depolarizing step show the loss of outward (IK1) current at potentials below 0 mV (control, ncells=8; 24 h, ncells=5; 72 h, ncells=7, * denotes P<0.05 vs. control).

 
3.5 Spatial properties of Ca²⁺ release in ventricular myocytes from failing human hearts
Fig. 8A, left panel, shows a representative example of a confocal line scan image recorded from a human cell under the same conditions as in Fig. 2. A total of 23 cells from five hearts were studied and delayed release areas were observed in all cells. We quantified the delayed areas in nine cells studied during field stimulation; the delayed release areas occupied 39±8% of the line scan and the mean width of these areas was 7.5±1.1 µm (n=43). As illustrated in the right panel of Fig. 8A, isoproterenol (3 µM) increased the transient magnitude, but did not synchronize release (similar results in 5 cells).

View larger version (51K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 8 Presence of delayed release areas in failing human myocytes. (A) Representative line scan image showing distinct regions of early and delayed SR Ca2+ release at baseline (left panel), which persisted following treatment with 3 µM isoproterenol (right panel). (B) Time course of the fraction of the line scan for which F>F50 (ncells=9). (C) Representative example of the central plane of Z-stack images obtained with di-8-ANEPPS staining. (D) Quantification of the T-tubule density (ncells=14).

 
Local transients in regions of delayed release were smaller than those in early release regions (mean magnitude 1.52±0.04 F/F₀ in delayed (n=43) and 1.76±0.04 F/F₀ in early (n=66) (P<0.05). Using spatial profiles at different time points, we examined the rate of rise of [Ca²⁺]_i as the percentage of the line scan with F>F₅₀ (Fig. 8B). This plot shows that in the failing human cells the time for half of the line to reach F₅₀ was 21±1 ms. This value is lower than in 24-h cultured pig cells (27±3 ms, P<0.05) and not significantly different from freshly isolated pig cells. The overall [Ca²⁺]_i transient magnitude was smaller in human myocytes than in the freshly isolated pig myocytes (F/F₀ 1.53±0.08, P<0.05).

Fig. 8C shows a representative example of T-tubule staining. Applying the same algorithms as for the pig myocytes, the T index for the human myocytes was 0.26±0.04. This density of T-tubules is not significantly different from the values for the freshly isolated pig myocytes (T index of 0.32±0.03). We noticed that in some areas dilated T-tubular structures were present, which were not observed in freshly isolated or cultured pig myocytes.

	4. Discussion

Top Abstract 1. Introduction 2. Methods 3. Results 4. Discussion References

 
We used short-term culture of ventricular myocytes to model loss of T-tubules. Progressive loss of T-tubule density was paralleled by an increase in the dyssynchrony of SR Ca²⁺ release. This increased dyssynchrony resulted in an overall slowing of the [Ca²⁺]_i transients. I_CaL total current and density were reduced, but SR Ca²⁺ content was unchanged. Ventricular myocytes from human failing hearts had areas of delayed release and dyssynchrony; the rate of rise of [Ca²⁺]_i was comparable to that of freshly isolated pig myocytes and significantly faster than in pig myocytes with pronounced loss of T-tubules.

4.1 Loss of T-tubules and dyssynchrony of Ca²⁺ release: implications for heart failure
Previous work had shown that the absence of T-tubules in ventricular myocytes leads to wave-like propagation of the [Ca²⁺]_i transient from the sarcolemma to the center [8,13,15,28]. Here we show that with a reduction of the T-tubule density to 30% of control values, there is a fragmentation of the front of the [Ca²⁺]_i transient, and a slowing of the rate of rise of the [Ca²⁺]_i transient. These data suggest that the reduction of T-tubule density to 50% of control values reported for the dog with tachycardia-induced heart failure [16,17] will significantly affect the rate of rise of [Ca²⁺]_i. Although there are currently no data on the spatial characteristics of the [Ca²⁺]_i transient in this model, data on the overall characteristics of the [Ca²⁺]_i transient are available [36]. These indeed show a reduction in the rate of rise. However, several other defects contribute to the altered excitation–contraction coupling in this model of heart failure, such as the down-regulation of SERCA and reduction in SR content [29]. It is noteworthy that in our study there was no reduction of the SR Ca²⁺ content, which could help to explain why the amplitude of the [Ca²⁺]_i transient was only modestly affected. The maintained SR content could be due to ‘autoregulation’ in the presence of a reduced trigger for Ca²⁺ release [3].

In human ventricular myocytes, the issue of whether the T-tubular network is altered is still debated. Because of the preferential location of Ca²⁺ channels in T-tubules, a loss of T-tubules would disproportionately decrease I_CaL density [27]. Therefore, the absence of evidence that the global density of the L-type Ca²⁺ current is reduced in human heart failure (see review in Ref. [30]) argues against the idea that there is a significant reduction of T-tubules. Loss of channels in T-tubules could, however, also be compensated by a higher activity of the channels [27,31,32], perhaps specifically in the external sarcolemma. Histologic examination of explanted hearts shows a less dense but more dilated T-tubular network [18]. We also observed some dilated structures, and, as our index measures the area occupied by di-8-ANEPPS staining, we may underestimate the decrease in T-tubule number. Preliminary data in isolated myocytes suggest that the density of T-tubules is decreased [20] or unaltered [19], based on analysis of the stained T-tubule network. More recently, it was reported that the S/V ratio is comparable for myocytes from failing and non-failing hearts (online data supplement of Ref. [33]). If changes in T-tubule density are present they are likely to be small, and will require a detailed analysis of the T-tubular network. S/V measurements may not be the best index. In the complex geometry of the ventricular cell, increases in volume are not necessarily linearly related to surface area, particularly with the pronounced hypertrophy observed in myocytes from end-stage failing hearts. In addition, if the T-tubule fraction of the total surface area is low, a small reduction in this fraction will have less impact on the S/V ratio. This is illustrated by the data of Fig. 1D and E, showing that the further reduction of T-tubule density by 50% between 24 and 72 h of culture did not significantly affect the S/V ratio.

Our observations in myocytes from failing human hearts are inconsistent with a pronounced loss of T-tubules. The values for T-tubule density, for the number and size of delayed release regions, and for the overall rate of rise of [Ca²⁺]_i are not significantly different for failing human myocytes and freshly isolated myocytes from pig hearts. Thus, our findings suggest that loss of T-tubules and reduced synchrony of SR Ca²⁺ release may not be an important mediator of the reduced efficiency of EC coupling in end-stage human heart failure. However, experiments such as in Fig. 8 clearly need to be performed in myocytes from undiseased human hearts as well, but in our institution it is very difficult to obtain such tissue for experimental studies. Based on a comparison between pig and mouse ventricular myocytes, we have previously proposed that the properties of pig myocytes could be more representative for mammals with a slower heart rate [10], though this needs to be confirmed for humans. Despite this limitation in the reference to control tissues, our observations indicate that the T-tubular network is still prominent in the failing human heart.

In the rabbit with heart failure following myocardial infarction, myocytes from the infarct border zone showed large inhomogeneities in the [Ca²⁺]_i transient front [34]. In this model, β-adrenergic stimulation improved the synchrony of Ca²⁺ release, suggesting that dephosphorylation of Ca²⁺ cycling proteins was partly responsible for reduced homogeneity of transients. However, it is possible that decreased T-tubule density may have contributed to delayed Ca²⁺ release in those regions not synchronized by β-adrenergic stimulation. In failing human cells, Ca²⁺ release was not synchronized by β-adrenergic stimulation (Fig. 8A), suggesting that inhomogeneous Ca²⁺ transients in these cells were solely related to the T-tubule density.

4.2. Alterations of excitation–contraction coupling with culture: implications for the use of cultured cells in long-term studies
Our data show a profound loss of T-tubules, as well as alterations in ion channels, with short-term culture of pig ventricular myocytes. Removing the serum from the culture medium did not prevent T-tubule loss (data not shown). These alterations are in line with decreased T-tubule density [8,35], I_K [35,36] and I_CaL [37] following culture of adult ventricular myocytes from other species. These findings have important implications for employing viral gene transfer in cultured cells to study excitation–contraction coupling, and stress the need for adequate control experiments. Although the difficulties of culturing adult myocytes are recognized, this approach has proven invaluable for the study of growth and its regulation (e.g. Refs. [37–39]). In addition, the morphological and electrophysiological alterations which occur during cell culture can also be used to the advantage as experimental models. In the present study, we have used myocyte culture to model loss of T-tubules. Nuss et al. [40] have employed cell culture to model cardiac hyper-excitability due to loss of I_K. In such modeling studies, some systems may prove more practical than others, as cultured-induced alterations in myocytes may vary depending on species and the specific culture conditions.

4.3. Conclusions
T-tubule loss is associated with a dose-dependent increase in dyssynchrony of SR Ca²⁺ release. When T-tubules are lost during cardiac remodeling, this will contribute to slower and smaller [Ca²⁺]_i transients. It remains to be established whether this mechanism is of major importance in end-stage human heart failure.

	Acknowledgements

 
This study was supported by the Fonds voor Wetenschappelijk Onderzoek-Vlaanderen, the Flanders Fund for Sciefsntific Research (K.R.S). The authors wish to thank Dr. Lothar Blatter for helpful comments and discussions. We thank P. Holemans, D. Decoux and T. Stassen for technical assistance.

	Notes

 
Time for primary review 20 days

	References

Top Abstract 1. Introduction 2. Methods 3. Results 4. Discussion References

 

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