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1 Department of Physiology and 2 Department of Medicine, and the 3 Cardiovascular Research Laboratories, David Geffen School of Medicine at UCLA, Los Angeles, CA 90095-1679, USA
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Abstract |
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[Ca2+]i= 466 ± 48 nM in transgenics versus 892 ± 104 nM in wild-type, P < 0.0005) and substantially reduced gain of excitation–contraction coupling. These alterations in excitation–contraction coupling may underlie the tendency for these animals to develop heart failure following haemodynamic stress.
(Received 23 September 2003;
accepted after revision 27 November 2003;
first published online 28 November 2003)
Corresponding author J. I. Goldhaber: David Geffen School of Medicine at UCLA, Division of Cardiology, 47–123 CHS 10833 LeConte Avenue, Los Angeles, CA 90095-1679, USA. Email: jgoldhaber{at}mednet.ucla.edu
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Introduction |
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The purpose of the present study was to determine whether homozygous overexpression of Na+–Ca2+ exchange leads to abnormalities in intracellular Ca2+ ([Ca2+]i) regulation and excitation–contraction (E-C) coupling in non-failing mice that might account for their tendency to develop CHF during haemodynamic stress. We find that in addition to the expected marked increase in Na+–Ca2+ exchange activity, these animals display abnormal E-C coupling, characterized by large slowly inactivating L-type Ca2+ currents, preserved sarcoplasmic reticulum (SR) Ca2+ stores, and reduced gain of E-C coupling. Exchanger overexpression has a strong modulatory effect on the amplitude and kinetics of the Ca2+ current, which has not been reported previously.
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Methods |
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We generated Na+–Ca2+ exchanger overexpressing mice as described in detail previously (Adachi-Akahane et al. 1997) and subsequently bred them to homozygosity (Roos et al. 2000). The animals used in this study did not display any gross pathology, had good cardiac function and showed no clinical evidence of heart failure.
Isolation of ventricular myocytes from adult mouse hearts
After I.P. injection of sodium heparin (1000 U kg-1), we anaesthetized adult mice with I.P. sodium pentobarbital and quickly removed hearts via thoracotomy in accordance with the guidelines of the UCLA Office for Protection of Research Subjects. We then isolated single ventricular myocytes enzymatically using collagenase (2 mg ml-1, Type II collagenase; Gibco BRL, Life Technologies, Gaithersburg, MD, USA) and protease (0.166 mg ml-1, Type XIV protease; Sigma-Aldrich, St Louis, MO, USA) digestion according to the method of Mitra & Morad (1985). Following isolation, we stored the dissociated cells for up to 6 h at room temperature in modified Tyrode solution, containing (mM): 136 NaCl, 5.4 KCl, 10 Hepes, 1.0 MgCl2, 0.33 NaH2PO4, 1.0 CaCl2, 10 glucose, pH 7.4 with NaOH. This solution was also used, with modifications described below, as the standard bath for electrophysiological recordings.
Electrophysiology
To record whole cell membrane currents, we placed the cells in an experimental chamber (0.5 ml) mounted on the stage of a Nikon Diaphot inverted microscope. A heated bath solution (26°C) continuously perfused the chamber. Patch electrodes were pulled from borosilicate glass (World Precision Instruments, Sarasota, FL, USA, TW150F-3) on a Sutter P-97 horizontal puller (Sutter Instruments, Novato, CA, USA). The fire-polished electrodes had a tip diameter of 2–3 µm and a resistance of 1–2 M
when filled with patch electrode solutions (described below). We recorded whole cell membrane current using an Axopatch 200 patch clamp amplifier (Axon Instruments, Union City, CA, USA) and a Digidata 1200 (Axon Instruments) data acquisition system controlled by pCLAMP 6 software (Axon Instruments). We applied series resistance compensation to all recordings.
Fluorescence measurements
To simultaneously record [Ca2+]i during voltage clamping, we employed a custom designed photometric epifluorescence detection system, described in detail previously (Goldhaber et al. 1991). Cells were loaded with the [Ca2+]i indicator fura-2 via the pipette solution, which contained (mM): 110 CsCl, 30 TEA-Cl, 10 NaCl, 10 Hepes, 5 MgATP, 0.1 cAMP, 0.1 K5fura-2, pH 7.3 with CsOH. [Ca2+]i concentration was calculated from the ratio (R) of the fluorescence intensities at the two excitation wavelengths (ratios at 600 Hz) using the method of Grynkiewicz et al. (1985), according to the equation:
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Solution exchange
Miniature solenoid valves (The Lee Co., Westbrook, CT, USA) controlled by pCLAMP's digital outputs controlled the bath solution flow through a micromanifold (ALA Scientific Instruments, Westbury, NY, USA). This enabled precise timing of solution exchanges in relation to the voltage clamp protocol. The solution surrounding the cell exchanges in less than 100 ms.
Dihydropyridine binding
For analysis of the L-type Ca2+ channel or dihydropyridine receptor (DHPR), we performed dihydropyridine binding studies with ventricular homogenates using methods similar to those previously described (Maan & Hosey, 1987). We washed freshly dissected hearts in ice-cold Hepes buffer containing (mM): 10 Hepes, 1.0 Na4EDTA, 2 MgSO4, 10 NaCl, pH 7.4 with NaOH. After trimming away atrial tissue, we homogenized the remaining myocardium and quantified protein (Lowry et al. 1951). For binding experiments, we used Hepes buffer containing 100 µg heart homogenate and varying concentrations of [3H]PN 200-110 (0.02–2.0 nM) in a total volume of 500 µl. The sample was incubated at 30°C for 60 min to allow equilibration of the receptor with the radioligand. Non-specific binding in the presence of 83 µM nifedipine (in the dark) was subtracted from the total bound radioactive ligand. We terminated all reactions by rapid filtration through Whatman GF/C filters (Whatman, Clifton, NJ, USA) using a vacuum manifold (Millipore Corp., Bedford, MA, USA) and washing the filters three times with 6 ml of ice-cold Hepes buffer. Radioactivity retained on the filters was determined in a scintillation counter. All experiments were performed in duplicate. (+)-[Methyl-3H]PN 200-110 was obtained from Amersham Pharmacia Biotech, Buckinghamshire, England and 45CaCl2 from ICN Radiochemicals, Irvine, CA, USA. Nifedipine was purchased from Calbiochem Corp., La Jolla, CA, USA.
Statistical analysis
Data are expressed as means ±S.E.M. Student's unpaired t test was used for direct comparisons of WT versus HOM. In experiments where a range of voltages was tested in each group, we used two-way ANOVA with Tukey–Fisher LSD post hoc testing (JMP 5.01a, SAS Institute, Cary, NC, USA). To test the effect of bath Na+ on the directional change of the Ca2+ transient, we used the Wilcoxon rank sum test (JMP 5.01a, SAS Institute). A P value < 0.05 was considered statistically significant.
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Results |
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We measured outward Na+–Ca2+ exchange current (Ca2+ influx mode) and [Ca2+]i in whole cell patched myocytes during 1 s step depolarizations from –80 to +80 mV. For these experiments, we used the fura-2 pipette solution described in Methods, and we replaced K+ in the standard bath Tyrode solution with Cs+ to block K+ currents. We also included 10 µM tetrodotoxin (TTX, Calbiochem) in the bath to block Na+ currents during depolarization. Representative tracings are shown in Fig. 1. We repeated the depolarization 100 ms after applying 5 mM Ni2+ to the bath using a rapid exchanger. In WT myocytes, the Ni2+-subtracted outward current during depolarization averaged 434 ± 92 pA (n= 12), whereas in HOM myocytes the current averaged 1016 ± 90 pA (n= 17, P < 0.005). This difference remained significant after correction for cell capacitance, which was increased in HOM cells (WT: 208 ± 16 pF; HOM: 260 ± 15 pF, P < 0.05).
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SR Ca2+ overload leads to regenerative Ca2+ release in isolated myocytes (Orchard et al. 1983). We measured SR Ca2+ content and inward Na+–Ca2+ exchange current (Ca2+ efflux mode) by recording [Ca2+]i and membrane current simultaneously during rapid application of 5 mM caffeine to the bath using a rapid solution exchanger. We administered a train of five 100 ms prepulses from –80 to 0 mV at 1 Hz immediately prior to caffeine application to ensure steady-state SR load, and maintained a constant holding potential of –80 mV during exposure to caffeine. Figure 3 shows [Ca2+]i during caffeine application and the resulting inward membrane current for representative WT and HOM myocytes. The peak inward current in HOM myocytes was about 3-fold larger than WT when normalized to capacitance (HOM: 5.5 ± 0.36 pA pF-1, n= 17; WT: 1.9 ± 0.16 pA pF-1, n= 12, P < 0.05).
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Next we measured the peak of the caffeine-induced [Ca2+]i transient to determine whether the releasable pool of SR Ca2+ was sufficient to raise cytoplasmic calcium to comparable levels in WT and HOM mycoytes (Fig. 3). We found no difference in the caffeine-induced [Ca2+]i transient between WT and HOM myocytes (WT: 815 ± 192 nM, n= 12; HOM: 727 ± 99 nM, n= 17, n.s.). In the presence of Ni2+ to block the Na+–Ca2+ exchanger during caffeine administration, there was still no significant difference in the peak [Ca2+]i transient between WT and HOM myocytes (data not shown). These results indicate that homozygous overexpression of the exchanger does not deplete cellular Ca2+ stores.
Caffeine-induced [Ca2+]i transients decayed faster in HOM myocytes than WT, consistent with increased effectiveness of Ca2+ removal by Na+–Ca2+ exchange (Fig. 3). When the declining phase of the Ca2+ transient triggered by 5 mM caffeine was fitted to a single exponential, WT transients decayed with a time constant of 134 ± 18 ms (n= 5). The rate of decay in HOM myocytes was increased twofold (64 ± 5 ms, n= 16, P < 0.01, HOM versus WT). In WT and HOM myocytes, 5 mM Ni2+ (which blocks Na+–Ca2+ exchange) blocked the caffeine-induced transient inward current and slowed the rate of decline of the [Ca2+]i transients 37-fold and 100-fold, respectively (WT: 5239 ± 1093 ms, n= 3, P < 0.05 versus Ni2+-free; HOM: 7056 ± 1039 ms, n= 5, P < 0.05versus Ni2+-free). The more dramatic reduction in the rate of decline of the [Ca2+]i transient in HOM myocytes is consistent with their increased dependence upon Na+–Ca2+ exchange for relaxation of the caffeine-induced [Ca2+]i transient.
Reduced E-C coupling gain in HOM myocytes
E-C coupling is altered in several animal models of cardiac hypertrophy and failure (Gómez et al. 1997; Dipla et al. 1999; Pogwizd et al. 1999; Wang et al. 2000; Hobai & O'Rourke, 2001). We therefore compared E-C coupling in HOM myocytes to WT. Cells were whole cell patch clamped using the fura-2 internal solution and TTX-containing bath solution described above. We first applied a train of six conditioning pulses from –80 to 0 mV at 1 Hz to equilibrate the SR, and then prepulsed to –40 mV for 1 s to inactivate current through Na+ channels. We did not observe any Ca2+ release during the prepulse. We then gave a test pulse from –40 to 0 mV for 200 ms while measuring both membrane current and [Ca2+]i. As shown in Fig. 4, the peak amplitude of ICa in HOM myocytes (8.2 ± 0.7 pA pF-1, n= 12) was significantly larger than WT (5.8 ± 0.9 pA pF-1, n= 12, P < 0.02). However [Ca2+]i release was significantly decreased in HOM compared to WT ([Ca2+]i= 466 ± 48 nM in HOM versus 892 ± 104 nM in WT, P < 0.0005). This decrease in the ratio of [Ca2+]i release to ICa indicates reduced E-C coupling gain in HOM myocytes (Fig. 4E). Since SR Ca2+ content is not decreased in HOM myocytes (see above), a reduction in releasable Ca2+ cannot account for the reduced Ca2+-induced [Ca2+]i release (CICR) and E-C coupling gain observed in these experiments.
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To test whether the Na+–Ca2+ exchanger in the HOM myocytes could actively modulate the critical components of E-C coupling, we measured ICa and [Ca2+]i release before and after transiently blocking the exchanger by removal of bath Na+. For these experiments, we again used the Na+-free pipette solution and a rapid switcher to briefly replace Na+ in the K+-free bath with Li+. The bath also contained TTX (10 µM). These interventions rapidly and transiently blocked the exchanger, but still permitted us to record ICa and the [Ca2+]i transient. Six 100 ms conditioning pulses to 0 mV were applied at 1 Hz as above to maintain steady-state SR Ca2+ stores. A 1 s prepulse to –40 mV immediately preceded the 200 ms depolarization to test potentials ranging from –30 to +40 mV.
In HOM myocytes, blocking Na+–Ca2+ exchange by abrupt removal of bath Na+ significantly reduced the amplitude of ICa at –20 mV (P < 0.005 by two-way ANOVApost hoc testing; Fig. 6A and B). ICa was reduced at –10 mV as well, but with borderline statistical significance (P = 0.053). However blocking Na+–Ca2+ exchange had no effect on the amplitude of ICa at positive potentials. The difference plot in Fig. 6C illustrates the voltage dependence of the exchanger's activating influence on the amplitude of ICa. The difference current is unlikely to be Na+–Ca2+ exchange current itself, since exchanger current would be present at all potentials tested because of its very positive reversal potential in the absence of pipette Na+ (even assuming residual subsarcolemmal Na+ as high as 1 mM).
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Although exchanger activity did not alter peak ICa at positive potentials, Na+–Ca2+ exchange activity significantly slowed inactivation of ICa at all potentials tested (Fig. 6F). The slowed inactivation may in part be secondary to the slight decrease in Ca2+ release flux we observed at those potentials, consistent with previous studies suggesting that Ca2+-induced inactivation of ICa is influenced predominantly by Ca2+ released by the SR (Adachi-Akahane et al. 1996). In WT cells, blocking the exchanger had no effect on ICa amplitude, Ca2+ release or kinetics. In Na+-free conditions, we still observed increased ICa, slowed ICa inactivation, and reduced Ca2+ release in HOM myocytes compared to WT (Fig. 7).
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To investigate whether increased expression of DHPR contributes to the enhanced whole cell Ca2+ current in HOM myocytes, we measured expression directly using a DHPR binding assay with the radiolabelled antagonist [3H]PN200-110. As shown in Fig. 8, there was no difference in the maximal binding capacity (Bmax) between WT, heterozygous, or homozygous transgenic hearts. Thus, the increased ICa we observed in HOM myocytes cannot be explained by an increase in the number of channels.
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Discussion |
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We have previously produced transgenic animals heterozygous for Na+–Ca2+ exchange overexpression, and though enhanced exchanger activity is clearly evident in these animals, they do not display a heart failure phenotype and substantial abnormalities in E-C coupling have not been noted. It is also noteworthy that heterozygous overexpressors do not exhibit significant increases in cell capacitance (Adachi-Akahane et al. 1997; Terracciano et al. 1998). Homozygous transgenic Na+–Ca2+ exchanger overexpressing animals have substantially greater exchanger activity than the heterozygotes studied previously. Hearts from HOM mice are hypertrophic but otherwise do not exhibit any gross pathology or functional abnormalities; postpartum females and mice of either sex subjected to transaortic banding are more susceptible to developing heart failure than wild-type counterparts (Roos et al. 2000). Thus, the increase in exchanger activity in HOM animals provokes important changes in cardiac growth, which are likely to be related to alterations in Ca2+ signalling.
Calcium handling during reverse exchange in HOM myocytes
Our electrophysiological data confirm that Na+–Ca2+ exchange activity (both forward and reverse) in HOM myocytes is markedly increased compared to WT. These results are consistent with 45Ca2+ uptake measurements in isolated membranes (Roos et al. 2000). Exchanger activity is sufficiently high in the HOM myocytes that reverse exchange activated by step depolarization to +80 mV always caused a rise in [Ca2+]i that the SR could not effectively buffer. The extent of Ca2+ influx was sufficient to cause oscillatory [Ca2+]i release in 13 out of 28 HOM myocytes, probably secondary to [Ca2+]i overload. Baseline elevations in SR Ca2+ content probably do not account for the response of HOM myocytes to reverse exchange, since our experiments indicate that SR Ca2+ content is unchanged compared to control. Instead, our findings are more consistent with Ca2+ entry during reverse exchange leading to Ca2+ overfilling of the SR, which then causes regenerative SR Ca2+ release (Orchard et al. 1983; Han et al. 1994; Bassani et al. 1995; Sipido et al. 2000). In heterozygous overexpressors, which reportedly have increased SR Ca2+ content (Terracciano et al. 1998), a similar voltage clamp protocol with 10 mM Na+ in the pipette did not consistently cause oscillatory Ca2+ release (Adachi-Akahane et al. 1997). Thus, even if we have underestimated SR Ca2+ content in HOM myocytes, we can conclude that Ca2+ influx via the overexpressed exchanger overfills the SR during depolarization to +80 mV and leads to oscillatory behaviour.
Increased L-type Ca2+ current and slowed inactivation kinetics in HOM myocytes: influence of Na+–Ca2+ exchange
The increased peak ICa, slowed inactivation, and left-shifted I–V relationship in HOM myocytes were unanticipated (Figs 5 and 7). These striking findings have not been reported in heterozygous overexpressors of Na+–Ca2+ exchange (Adachi-Akahane et al. 1997; Yao et al. 1998). The increased peak ICa in HOM myocytes cannot be explained by an increased number of Ca2+ channels, since we found no increase in dihydropyridine binding compared with WT. Since ICa was still larger in HOM compared with WT, even in Na+-free conditions (Fig. 7), it is unlikely that Na+–Ca2+ exchange activity is directly responsible for the difference in ICa amplitude between HOM and WT.
However, our results clearly demonstrate that overexpression of the Na+–Ca2+ exchanger significantly increases its ability to influence ICa regulation. Acutely blocking the exchanger by removing bath Na+ accelerated ICa inactivation kinetics (Fig. 6). Previous studies in normal guinea-pig and rat ventricular myocytes have found that SR Ca2+ release, not Na+–Ca2+ exchange, is the major source of the Ca2+ that regulates Ca2+-dependent inactivation of ICa (Sipido et al. 1995; Adachi-Akahane et al. 1996). When compared to WT, [Ca2+]i transients in HOM myocytes are significantly reduced, which could contribute to slowed inactivation of ICa. However, in HOM myocytes, the effect of blocking the exchanger on ICa inactivation kinetics occurred in the absence of significant increases in the [Ca2+]i transient. Overall, the results suggest that the packing of additional exchangers into close proximity of L-type Ca2+ channels in HOM myocytes facilitates greater interaction between the proteins. Robust exchanger activity in the dyadic clefts of HOM myocytes appears to rapidly remove Ca2+ from the microenvironment of L-type Ca2+ channels during depolarization. This reduces Ca2+-induced inactivation of ICa, which may also contribute to the increase in the peak amplitude of ICa that we observe at negative potentials.
We took several precautions to avoid contamination of ICa records by Na+-Ca2+ exchange current and sodium current. Prior to recording ICa, we prepulsed to -40 mV and applied TTX to the bath solution to eliminate Na+ current. Artifacts due to poor voltage control are improbable, as we were careful to apply series resistance compensation in all experiments. By substituting Li+ for Na+ we minimized any voltage errors due to changes in surface charge. The risk of ICa contamination by Na+-Ca2+ exchange is highest at negative potentials, where we found the most dramatic effect of the exchanger on ICa amplitude. However, the consistent decrease in [Ca2+]i release and inward current at -20 mV upon blockade of the exchanger is also inconsistent with contamination of the ICa record by Na+-Ca2+ exchange. A reduction in forward exchange (Ca2+ efflux) caused by Na+ removal would instead tend to enhance the Ca2+ transient.
In summary, ICa is increased in HOM myocytes, even when exchanger current is acutely blocked. In addition, the increased Na+-Ca2+ exchange activity in these myocytes appears to directly influence ICa, increasing peak ICa at negative potentials and slowing ICa inactivation, especially at positive potentials. There is a logic as to how exchange activity may influence inactivation, but the reason for the increased peak ICa is unclear.
Defective E-C coupling
We (Goldhaber et al. 1999) and others (Litwin et al. 1998; Viatchenko-Karpinski & Gyorke, 2001) have demonstrated that Na+–Ca2+ exchange is capable of modulating SR Ca2+ release, presumably by altering the concentration of Ca2+ in the dyadic cleft. In the setting of massive exchanger overexpression, we hypothesized that Na+–Ca2+ exchange would have a significant effect on CICR. Indeed, we found that in HOM myocytes ICa was significantly less effective at releasing SR Ca2+ than in WT myocytes (Figs 4, 5 and 7). The reduction in E-C coupling gain was not due to SR Ca2+ depletion as it clearly occurred in the presence of pipette Na+ and intact [Ca2+]i stores (Fig. 4E). The small increase in Ca2+ release flux associated with blockade of Na+–Ca2+ exchange at +10 mV (Fig. 6E) is consistent with the hypothesis that the overexpressed exchanger population acts as a sink for Ca2+, reducing trigger Ca2+ flux before ryanodine receptors sense that Ca2+. As discussed above, this removal of Ca2+ from the dyadic cleft may also contribute to the slowed inactivation of ICa that we observed in HOM myocytes. Such a situation arises because additional exchangers are packed into the sarcolemmal membrane, resulting in possible closer proximity to L-type Ca2+ channels. Nevertheless, inhibition of the exchanger by Na+ removal did not substantially increase gain in HOM myocytes. Furthermore, the primary differences in ICa amplitude, ICa kinetics and [Ca2+]i release between HOM and WT were evident even when Na+–Ca2+ exchange was blocked (Fig. 7). Thus, while exchange activity in HOM myocytes can acutely modulate ICa and [Ca2+]i release, the reduced gain in these transgenic animals is independent of exchange activity and appears to be an inherent property of this model. We hypothesize that the reduced gain is a maladaptive hypertrophic response secondary to abnormal Ca2+ signalling.
An alternative explanation for the reduced gain in HOM myocytes is a change in the spatial arrangement of SR Ca2+-release channels and L-type Ca2+ channels. If the average distance between the two channels is increased, the SR release channels will be less effectively activated by the local Ca2+ influx produced by the opening of a DHPR (Stern, 1992; Gómez et al. 1997) thus leading to the reduction in gain. This speculative explanation has been invoked in other cases of reduced gain (Gómez et al. 1997). Such a situation might be aggravated by the presence of additional exchangers close to L-type Ca2+ channels. A reduction in ryanodine receptor (RyR) levels cannot explain the reduced gain since RyR levels, as indicated by microarray analysis, are unchanged in the hearts from HOM mice (Roos et al. 2000). We cannot exclude the possibility that RyR gating is modified in HOM myocytes, but we have no direct evidence to support this.
Role of overexpressed Na+-Ca2+ exchange in hypertrophy and failure
This study was undertaken in part to determine what E-C coupling abnormalities underlie the tendency for homozygous overexpressors to develop heart failure. We found that non-failing myocytes from these transgenic animals have increased capacitance and depressed E-C coupling gain. The reduced gain is similar to that seen in some animal models of hypertrophy leading to failure, such as the hypertensive Dahl salt-sensitive rat and aortic-banded rat hearts, though hypertrophy and failure models do not typically exhibit enhanced L-type Ca2+ current density (Gómez et al. 1997; Ahmmed et al. 2000; Volk & Ehmke, 2002). The increased whole cell L-type Ca2+ current density, and slowed inactivation kinetics observed in HOM myocytes may help support near normal [Ca2+]i release from the SR during depolarization and compensate for the decreased gain in this model. In conclusion, we have found that high level overexpression of Na+-Ca2+ exchange in transgenic animals leads to significant changes in E-C coupling that may contribute to the susceptibility of these animals to heart failure.
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References |
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Adachi-Akahane S, Lu L, Li Z, Frank JS, Philipson KD & Morad M (1997). Calcium signaling in transgenic mice overexpressing cardiac Na+-Ca2+ exchanger. J General Physiol 109, 717–729.
Ahmmed GU, Dong PH, Song G, Ball NA, Xu Y, Walsh RA & Chiamvimonvat N (2000). Changes in Ca2+ cycling proteins underlie cardiac action potential prolongation in a pressure-overloaded guinea pig model with cardiac hypertrophy and failure. Circ Res 86, 558–570.
Bassani JW, Yuan W & Bers DM (1995). Fractional SR Ca release is regulated by trigger Ca and SR Ca content in cardiac myocytes. Am J Physiol 268, C1313–C1319.[Medline]
Dipla K, Mattiello JA, Margulies KB, Jeevanandam V & Houser SR (1999). The sarcoplasmic reticulum and the Na+/Ca2+ exchanger both contribute to the Ca2+ transient of failing human ventricular myocytes. Circ Res 84, 435–444.
Goldhaber JI, Lamp ST, Walter DO, Garfinkel A, Fukumoto GH & Weiss JN (1999). Local regulation of the threshold for calcium sparks in rat ventricular myocytes: role of sodium-calcium exchange. J Physiol 520, 431–438.
Goldhaber JI, Parker JM & Weiss JN (1991). Mechanisms of excitation-contraction coupling failure during metabolic inhibition in guinea-pig ventricular myocytes. J Physiol 443, 371–386.
Gómez AM, Valdivia HH, Cheng H, Lederer MR, Santana LF, Cannell MB, McCune SA, Altschuld RA & Lederer WJ (1997). Defective excitation-contraction coupling in experimental cardiac hypertrophy and heart failure. Science 276, 800–806.
Grynkiewicz G, Poenie M & Tsien RY (1985). A new generation of Ca2+ indicators with greatly improved fluorescence properties. J Biol Chem 260, 3440–3450.
Han S, Schiefer A & Isenberg G (1994). Ca2+ load of guinea-pig ventricular myocytes determines efficacy of brief Ca2+ currents as trigger for Ca2+ release. J Physiol 480, 411–421.
Hobai IA & O'Rourke B (2000). Enhanced Ca2+-activated Na+-Ca2+ exchange activity in canine pacing-induced heart failure. Circ Res 87, 690–698.
Hobai IA & O'Rourke B (2001). Decreased sarcoplasmic reticulum calcium content is responsible for defective excitation-contraction coupling in canine heart failure. Circulation 103, 1577–1584.
Litwin SE, Li J & Bridge JH (1998). Na-Ca exchange and the trigger for sarcoplasmic reticulum Ca release: studies in adult rabbit ventricular myocytes. Biophys J 75, 359–371.
Lowry OH, Rosebrough NJ, Farr AL & Randall RJ (1951). Protein measurement with the folin phenol reagent. J Biol Chem 193, 265–275.
Maan AC & Hosey MM (1987). Analysis of the properties of binding of calcium-channel activators and inhibitors to dihydropyridine receptors in chick heart membranes. Circ Res 61, 379–388.
Mitra R & Morad M (1985). A uniform enzymatic method for dissociation of myocytes from hearts and stomachs of vertebrates. Am J Physiol 249, H1056–H1060.[Medline]
Orchard CH, Eisner DA & Allen DG (1983). Oscillations of intracellular Ca2+ in mammalian cardiac muscle. Nature 304, 735–738.[CrossRef][Medline]
Pogwizd SM, Qi M, Yuan W, Samarel AM & Bers DM (1999). Upregulation of Na+/Ca2+ exchanger expression and function in an arrhythmogenic rabbit model of heart failure. Circ Res 85, 1009–1019.
Roos KP, Jordan MC, Lu L, Lu Y, Philipson KD & Ross RS (2000). Transgenic overexpression of the Na+-Ca2+ exchanger results in postpartum hypertrophy and heart failure. FASEB J 14, A696.
Sipido KR, Callewaert G & Carmeliet E (1995). Inhibition and rapid recovery of Ca2+ current during Ca2+ release from sarcoplasmic reticulum in guinea pig ventricular myocytes. Circ Res 76, 102–109.
Sipido KR, Volders PG, de Groot SH, Verdonck F, Van de Werf F, Wellens HJ & Vos MA (2000). Enhanced Ca2+ release and Na/Ca exchange activity in hypertrophied canine ventricular myocytes: potential link between contractile adaptation and arrhythmogenesis. Circulation 102, 2137–2144.
Stern MD (1992). Theory of excitation-contraction coupling in cardiac muscle. Biophys J 63, 497–517.
Studer R, Reinecke H, Bilger J, Eschenhagen T, Bohm M, Hasenfuss G, Just H, Holtz J & Drexler H (1994). Gene expression of the cardiac Na+-Ca2+ exchanger in end-stage human heart failure. Circ Res 75, 443–453.
Terracciano CM, Souza AI, Philipson KD & MacLeod KT (1998). Na+-Ca2+ exchange and sarcoplasmic reticular Ca2+ regulation in ventricular myocytes from transgenic mice overexpressing the Na+-Ca2+ exchanger. J Physiol 512, 651–667.
Viatchenko-Karpinski S & Gyorke S (2001). Modulation of the Ca2+-induced Ca2+ release cascade by ß-adrenergic stimulation in rat ventricular myocytes. J Physiol 533, 837–848.
Volk T & Ehmke H (2002). Conservation of L-type Ca2+ current characteristics in endo- and epicardial myocytes from rat left ventricle with pressure-induced hypertrophy. Pflugers Arch 443, 399–404.[CrossRef][Medline]
Wang W, Cleemann L, Jones LR & Morad M (2000). Modulation of focal and global Ca2+ release in calsequestrin-overexpressing mouse cardiomyocytes. J Physiol 524, 399–414.
Wang Z, Nolan B, Kutschke W & Hill JA (2001). Na+-Ca2+ exchanger remodeling in pressure overload cardiac hypertrophy. J Biol Chem 276, 17706–17711.
Yao A, Su Z, Nonaka A, Zubair I, Lu L, Philipson KD, Bridge JH & Barry WH (1998). Effects of overexpression of the Na+-Ca2+ exchanger on transients in murine ventricular myocytes. Circ Res 82, 657–665.
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Acknowledgements |
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Author's present address
H. Reuter: Laboratory for Muscle Research and Molecular Cardiology, Klinik III für Innere Medizin, Universität zu Köln, Joseph-Stelzmann-Str. 9, 50924 Köln, Germany.
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K. Imahashi, C. Pott, J. I. Goldhaber, C. Steenbergen, K. D. Philipson, and E. Murphy Cardiac-Specific Ablation of the Na+-Ca2+ Exchanger Confers Protection Against Ischemia/Reperfusion Injury Circ. Res., October 28, 2005; 97(9): 916 - 921. [Abstract] [Full Text] [PDF]
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 H. Reuter, C. Pott, J. I. Goldhaber, S. A. Henderson, K. D. Philipson, and R. H.G. Schwinger Na+-Ca2+exchange in the regulation of cardiac excitation-contraction coupling Cardiovasc Res, August 1, 2005; 67(2): 198 - 207. [Abstract] [Full Text] [PDF]
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S. A. Henderson, J. I. Goldhaber, J. M. So, T. Han, C. Motter, A. Ngo, C. Chantawansri, M. R. Ritter, M. Friedlander, D. A. Nicoll, et al. Functional Adult Myocardium in the Absence of Na+-Ca2+ Exchange: Cardiac-Specific Knockout of NCX1 Circ. Res., September 17, 2004; 95(6): 604 - 611. [Abstract] [Full Text] [PDF]
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) for inactivation of ICa (D) were obtained in a subset of cells using Na+-free fura-2 pipette solution. Peak ICa and its inactivation kinetics were increased in HOM (n= 6) compared to WT (n= 3), and the voltage dependence of ICa was shifted to the left. [Ca2+]i release and gain were decreased in HOM. See text for details. 
P < 0.05 for HOM versus WT by two-way ANOVA. *P < 0.05 at indicated voltages by Fisher–Tukey post hoc testing.
