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RECEPTORS AND SIGNAL TRANSDUCTION
Department of Physiology, Stritch School of Medicine, Loyola University Chicago, Maywood, Illinois
Submitted 12 December 2005 ; accepted in final form 18 May 2006
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ABSTRACT |
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 TOP ABSTRACT METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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mitochondrial Ca2+; excitation-contraction coupling; cardiomyocytes
) across the inner mitochondrial membrane, and a mechanism known as rapid mode of uptake (RaM), which operates transiently during the initial phase of pulsatile elevations of extramitochondrial [Ca2+] concentration ([Ca2+]em) (6, 25). Ca2+ uptake via RaM is at least 300 times more rapid than uptake via MCU; however, the recovery of RaM after a Ca2+ pulse in isolated heart mitochondria required >60 s (6), rendering this mechanism essentially inactivated during cardiac [Ca2+]i oscillations. A recent patch-clamp study suggests that MCU is a highly selective (Kd < 2 nM) Ca2+ channel (35), with slow allosteric regulation by extramitochondrial Ca2+ (32). Generally, a biphasic time course of mitochondrial Ca2+ uptake can be anticipated from its electrogenic nature. Rapid Ca2+ entry is slowed to a level at which Ca2+ influx is balanced by H+ ejection (see, e.g., Ref. 34). Ca2+ extrusion from heart mitochondria is mediated primarily via the Na+/Ca2+ antiporter (NCXm) while the H+/Ca2+ exchanger is thought to play no or only a minor role (25). Two fundamentally different scenarios have been proposed for mitochondrial decoding of rapid cardiac [Ca2+]i transients (see Ref. 30 for review). In model I, introduced by Crompton (9), Ca2+ uptake into mitochondria is slow, followed by even slower release of accumulated Ca2+. According to this model, fast cytosolic [Ca2+]i oscillations are integrated by the Ca2+ transport machinery of the inner mitochondrial membrane. Increasing the frequency or the amplitude of [Ca2+]i transients will result in a net accumulation of Ca2+ in the matrix compartment until a new steady state is reached when Ca2+ uptake during a single cycle equals Ca2+ efflux. Consequently, beat-to-beat changes in [Ca2+]m are small, thus minimizing the energetic costs of mitochondrial Ca2+ transport.
In contrast, model II postulates a highly efficient translation of cytosolic Ca2+ signals into changes in the free Ca2+ concentration in the matrix compartment. This model necessitates the existence of both a rapid Ca2+ uptake and a Ca2+ release mechanism in mitochondria in situ. Ca2+ uptake with each contractile cycle must be large enough to outcompete matrix buffers. Another prediction of this model is that mitochondrial Ca2+ uptake would effectively buffer [Ca2+]i transients during excitation-contraction (E-C) coupling. As a consequence, SR Ca2+ release and reuptake must be large enough to compensate for this additional fast buffering. With approximately one-third of cell volume being occupied by mitochondria, the additional SR Ca2+ fluxes would be substantial. Finally, the question must be addressed as to how Ca2+-dependent matrix enzymes respond to fast oscillatory changes in [Ca2+]m.
The experimental data in support for one (slow integration, model I) or the other (beat-to-beat transmission, model II) model of response of [Ca2+]m to rapid changes of [Ca2+]i in studies on intact cardiac myocytes seem to depend on the experimental technique and species used (for reviews see Refs. 22 and 30). Even studies utilizing the same technique such as electron probe microanalysis (EPMA) provided conflicting results, either in support of (31, 52) or against (38) oscillatory changes in total mitochondrial Ca2+ during E-C coupling.
In the present study, we have developed an experimental model that allows us to monitor selectively [Ca2+]m changes in mitochondria during global [Ca2+]i transients. For this purpose fluo-3-loaded cat ventricular cardiomyocytes with dye entrapped in mitochondria were permeabilized with digitonin. Fast [Ca2+]i transients were simulated by computer-controlled pressure ejection of an internal solution containing 100 µM Ca2+ through a glass micropipette positioned upstream of the cell that was continuously superfused with bulk extramitochondrial solution. The results of this study indicate that repetitive increases of [Ca2+]em caused a net accumulation of Ca2+ into mitochondria with each [Ca2+]em pulse. Mitochondrial Ca2+ accumulation increased with an increase in pulse duration and stimulation frequency; however, no mitochondrial [Ca2+]m oscillations were observed, either under conditions facilitating maximal rate of mitochondrial calcium uptake [high [Ca2+]em, 0 extramitrochondrial Na+ concentration ([Na+]em)] or when the rate of Ca2+ removal was maximized (0 [Ca2+]em, high [Na+]em). The slow frequency-dependent increase of [Ca2+]m argues against a rapid transmission of Ca2+ signals between cytosol and mitochondria during cardiac E-C coupling.
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METHODS |
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5 min, followed by perfusion with a nominally Ca2+-free Tyrode solution. After 5 min the perfusion was switched to a low (36 µM)-Ca2+ Tyrode solution containing 0.06% collagenase (Worthington Biochemical, type II) for 25–30 min. After collagenase perfusion small pieces of ventricular muscle were sliced from the free wall of the left ventricular endocardial surface and agitated in fresh 0.06% collagenase and 0.01% protease (type XIV, Sigma, St. Louis, MO) for 5 min. Cells were stored before use in a HEPES-buffered modified Tyrode solution containing (mM) 145 NaCl, 4 KCl, 1 MgCl2, 2 CaCl2, 5 HEPES, and 11 glucose and titrated with NaOH to a pH of 7.4. Myocytes were used within 1–6 h after isolation. All experiments were carried out at room temperature (20–22°C). Ca2+ measurements and cell permeabilization. Measurements of [Ca2+]m were performed with wide-field epifluorescence microscopy (except for the data shown in Fig. 1; see below). Spatially averaged photometric [Ca2+] measurements from single ventricular myocytes were obtained with the fluorescent Ca2+ indicator fluo-3 (estimated dissociation constant for the Ca2+-fluo-3 complex in the cellular environment is 1.1 µM; Ref. 29). Cells were loaded with the membrane-permeant acetoxymethyl ester of the dye (fluo-3 AM; Invitrogen/Molecular Probes, Carlsbad, CA) at 15 µM for 40 min in standard Tyrode solution containing (mM) 135 NaCl, 5 KCl, 1 MgCl2, 2 CaCl2, 10 glucose, and 10 HEPES (pH 7.3). Cells were subsequently washed for 10 min. Fluo-3 fluorescence was excited at 485 nm, and emitted fluorescence was recorded at 535 nm. Fluorescence signals were low-pass filtered at 1 kHz and sampled at 5 Hz. [Ca2+]m is expressed as F/F0, i.e., as fluo-3 fluorescence (F) normalized to the basal fluorescence levels (F0) measured after permeabilization of the surface membrane.
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100 nM, pH 7.2. BDM was added to reduce motion artifacts resulting from exposure of permeabilized myocytes to micromolar [Ca2+]. All extramitochondrial perfusion solutions with elevated [Ca2+] (0.5–100 µM) contained 5 mM EGTA and CaCl2 in amounts calculated to yield free [Ca2+]em as indicated. In the extramitochondrial perfusion solution containing 20 or 40 mM [Na+], NaCl replaced KCl by an equimolar amount. In Na+-free extramitochondrial solution KCl was increased by an equimolar amount.
Laser scanning confocal microscopy (LSM 410, Carl Zeiss) was used for the visualization of the spatial distribution of fluo-3 fluorescence before and after permeabilization (see Fig. 1A) and for colocalization experiments with MitoTracker Red (Fig. 1B). Fluo-3 fluorescence was excited with the 488-nm line of the argon ion laser, and the emitted fluorescence signal was measured simultaneously at 
510 nm. For colocalization experiments (Fig. 1B), cells were loaded with fluo-3 and with the mitochondria-specific probe MitoTracker Red CMXRos (Invitrogen/Molecular Probes; 200 nM). Both indicators were excited at 488 nm, and the emitted fluorescence signals were measured simultaneously at 510–525 (fluo-3) and 590 (MitoTracker Red CMXRos) nm.
Rapid solution application.
To mimic cytoplasmic changes of [Ca2+] in permeabilized cells similar to those triggered by action potentials in intact myocytes during E-C coupling, [Ca2+]em was changed rapidly with a pressure ejection system. For this purpose, permeabilized cells were placed in the laminar flow of a Ca2+-free "intracellular" solution (1 mM EGTA, no CaCl2 added). A glass micropipette (tip diameter 5–10 µm) was positioned upstream of the cell with regard to the direction of the bulk bath flow (see 
Fig. 6A). A Ca2+ (100 µM)-containing solution was pressure ejected by using a computer-controlled pressure microinjection device (model PV830 Pneumatic PicoPump; WPI, Sarasota, FL) allowing the application of exactly timed steplike Ca2+ pulses to the cell. From the experiments shown in Fig. 2 and Fig. 8A, top, we estimated that [Ca2+] of the pipette solution ([Ca2+]pip) was diluted to 5 µM at the cell.
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Statistical analysis. Statistical differences of the data were determined with the Student's t-test for unpaired or paired data and considered significant at P < 0.05. Results are reported as means ± SE for the indicated number (n) of cells.
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RESULTS |
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TOPABSTRACTMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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25 µm2) as indicated in Fig. 1A, bottom. Plasma membrane permeabilization with digitonin removed cytosolic and nuclear fluo-3 (Fig. 1Ab, after digitonin) and resulted in a significant drop in fluo-3 fluorescence. The level of fluorescence after digitonin treatment (F0) represents the contribution of mitochondria and was used to normalize the fluorescence signal (F/F0). The subsequent elevation of [Ca2+]em from 0.1 to 10 µM resulted in the increase of the mitochondria-trapped fluo-3 fluorescence (Fig. 1Ac, after Ca2+ addition), suggesting Ca2+ accumulation into mitochondria. In Fig. 1B simultaneously acquired images of the mitochondrial marker MitoTracker Red (red) and fluo-3 (green) fluorescence distribution from a permeabilized ventricular myocyte are shown. These images are overlaid in Fig. 1Bc, with yellow indicating spatial overlap of the two fluorescent probes. The MitoTracker Red signal was highly colocalized with fluo-3. indicating that the measured fluo-3 signals originated from mitochondria and report [Ca2+]m.
Dependence of mitochondrial Ca2+ uptake on [Ca2+]em. To examine the [Ca2+]em dependence of mitochondrial Ca2+ uptake, permeabilized cells were exposed to various [Ca2+]em in the absence of Na+ (Fig. 2). At the end of each experiment 100 µM Ca2+ was applied to obtain a maximum fluorescence response (Fmax) and to normalize mitochondrial Ca2+ uptake. Figure 2A shows an overlay of normalized traces of mitochondrial Ca2+ uptake obtained at [Ca2+]em of 0.5, 1, and 10 µM, indicating that increasing [Ca2+]em resulted in a concentration-dependent increase in the rate and magnitude of mitochondrial Ca2+ accumulation. We used the rate of increase of the normalized [Ca2+]m transients for the quantitative characterization of the concentration dependence of mitochondrial Ca2+ uptake in cat ventricular myocytes (Fig. 2B). To minimize potential problems arising from the nonlinear relationship between [Ca2+]m and fluo-3 fluorescence, the rate of Ca2+ uptake was determined from the initial linear phase of the [Ca2+]m increase. Individual fluorescence traces were normalized to Fmax, and Ca2+ uptake rates were then calculated as percent change of F in relation to Fmax per second (thus the units %Fmax/s). Data were fitted with the Hill equation, yielding a Hill coefficient (nH) of 2.4. [Ca2+]em resulting in half-maximal rate (k0.5) of Ca2+ accumulation was 4.4 µM. There was no activation of the Ca2+ uniporter at [Ca2+]em < 0.5 µM. nH >1 suggested cooperativity of the Ca2+ uptake process.
Inhibition of mitochondrial Ca2+ uptake with RR and with protonophore FCCP.
To confirm that observed changes in [Ca2+]m were indeed due to activation of Ca2+ uniporter-mediated mitochondrial Ca2+ uptake, cells were treated with RR, an inhibitor of MCU. As shown in Fig. 3A, stepwise increases of [Ca2+]em resulted in concentration-dependent Ca2+ uptake under control conditions but failed to increase [Ca2+]m in the presence of 10 µM RR. Qualitatively similar results were obtained with the protonophore FCCP, an uncoupler of the respiratory chain that abolishes the electrical driving force for the Ca2+ uniporter through dissipation of (Fig. 3A). Figure 3B summarizes the effect of RR and FCCP on the rate of Ca2+ uptake into the mitochondrial matrix calculated as described for the previous experiment and expressed as percent Fmax per second. Elevation of [Ca2+]em from 0.1 to 1 µM increased [Ca2+]m at a rate (%Fmax/s) of 1.6 ± 0.3 (n = 16). In the presence of FCCP this rate decreased to 0.02 ± 0.01 (n = 3), whereas no measurable change in [Ca2+]m was observed in the presence of 10 µM RR (n = 5). During application of 10 µM Ca2+ the rates of [Ca2+]m increase were 0.03 ± 0.00 (n = 5) with RR, 1.0 ± 0.3 (n = 5) in the presence of FCCP, and 49.5 ± 4.5 (n = 16) in the control group (P < 0.001 for both treatments). These data indicate that the increase of the fluo-3 signal following the elevation of [Ca2+]em represents a Ca2+ uniporter-mediated, -dependent Ca2+ uptake into the mitochondria.
Role of inorganic phosphate for mitochondrial Ca2+ uptake.
Although the primary role of inorganic phosphate (Pi) in mitochondrial function is the generation of ATP from ADP and phosphate by the mitochondrial ATPase, it also plays a role in Ca2+ entry and the Ca2+ accumulation capacity of mitochondria. A portion of Ca2+ entering the mitochondria precipitates as calcium phosphate, which helps lower the levels of free mitochondrial Ca2+ and thereby maintain the chemical gradient for Ca2+ entry. Pi is the main intracellular membrane-permeant anion and enters the mitochondrial matrix together with protons. Mitochondrial Pi uptake lowers mitochondrial pH and increases and therefore the electrical driving force for Ca2+ uptake (10, 40). Thus, through these mechanisms, it is expected that higher levels of Pi would facilitate mitochondrial Ca2+ uptake. We tested the effect of low (0.5 mM) and high (5 mM) Pi concentration ([Pi]) on mitochondrial Ca2+ accumulation at two different [Ca2+]em (0.5 and 1 µM). At both [Ca2+]em tested the presence of a high [Pi] clearly enhanced mitochondrial Ca2+ uptake, whereas the relative effect was more pronounced at the lower [Ca2+]em (Fig. 4, A and B). As shown in Fig. 4C, at 0.5 µM [Ca2+]em mitochondrial Ca2+ uptake was about threefold higher with high [Pi] than with low [Pi]. However, during application of 1 µM [Ca2+]em, high [Pi] enhanced Ca2+ uptake by 30%. These data show that mitochondrial Ca2+ uptake critically depends on [Pi].
Effect of [Na+]em on mitochondrial Ca2+ uptake and extrusion. The effect of [Na+]em on both Ca2+ uptake and Ca2+ extrusion was tested in permeabilized ventricular myocytes. Cells were exposed to 1 µM Ca2+ in the absence and in the presence of 40 mM [Na+]em. In the absence of extramitochondrial Na+, [Ca2+]m increased at a faster rate and reached higher levels (Fig. 5A, left) on exposure to 1 µM Ca2+ than in the presence of 40 mM Na+ (Fig. 5A, right). [Ca2+]m did not decline after removal of Ca2+ with no Na+ present in the extramitochondrial solution; however, it decreased to basal levels as soon as 40 mM Na+ was added. The Na+ dependence suggests that Ca2+ extrusion is carried by mitochondrial Na+/Ca2+ exchange. When cells were exposed to 1 µM Ca2+ in the presence of Na+, [Ca2+]m plateaued at a lower level than in the absence of Na+, presumably because Na+-dependent Ca2+ extrusion counteracted Ca2+ uptake. Consequently, removal of Na+ in the maintained presence of 1 µM Ca2+ resulted in an increase of [Ca2+]m that eventually reached the same level as during exposure to 1 µM Ca2+ in the complete absence of Na+ (Fig. 5A, left; dashed line). Figure 5B summarizes the effect of [Na+]em on the amplitude of the mitochondrial Ca2+ accumulation. Average [Ca2+]m are expressed as percentages of the maximal amplitude achieved with exposure to 100 µM extramitochondrial Ca2+. The data indicate that the kinetics of Ca2+ uptake and extrusion as well as the magnitude of Ca2+ accumulation critically depend on [Na+]em due to Ca2+ extrusion by the mitochondrial Na+/Ca2+ exchange mechanism.
Technique used to generate rapid changes in [Ca2+]em to simulate [Ca2+]i transients. We have developed a new experimental technique that allowed us to simulate fast [Ca2+]i transients in myocytes with permeabilized cell membrane. Permeabilized cardiomyocytes were placed in the laminar flow of a Ca2+-free solution containing 1 mM EGTA (bulk flow; see Fig. 6A). For the fast application of Ca2+-containing solution ([Ca2+]pip = 100 µM), a glass micropipette with a tip diameter of 5–10 µm was positioned upstream of the cell with regard to the direction of bulk flow. The pipette solution was pressure ejected by using a computer-controlled picopump. To ensure that pressure ejection of a Ca2+-containing solution indeed induced a fast increase in [Ca2+]em, and that bulk flow was fast enough to remove the elevated [Ca2+]em to simulate a typical [Ca2+]i transient, we performed the following control experiment (Fig. 6B). In this experiment we pressure ejected a solution containing 100 µM fluo-3 free acid and 100 µM Ca2+ in a pulsatile fashion at a frequency of 2 Hz and a pulse duration of 40 ms (Fig. 6B, bottom). Each pulse resulted in the release of Ca2+ from the pipette and a rapid increase in fluo-3 fluorescence (Fig. 6B, top). The increase in fluo-3 fluorescence was transient because after every applied pulse elevated [Ca2+]em was brought back to the initial level by the fast bulk flow. The time course of the signal is clearly reminiscent of [Ca2+]i transients elicited by action potentials typically recorded in intact cardiac myocytes. Thus we created a novel technique that allowed us to imitate physiological beat-to-beat cardiac [Ca2+]i transients in a plasma membrane-permeabilized cell.
After the methodology was established, we applied short pulses (pulse duration 0.5 s; [Ca2+]pip = 100 µM) to fluo-3 AM-loaded permeabilized cardiomyocytes at a frequency of 0.5 Hz (Fig. 6C). [Na+]em in the bulk solution was 20 mM to facilitate Ca2+ extrusion. Each pulse of elevated [Ca2+]em evoked an increase in the level of [Ca2+]m with no or very little decline of [Ca2+]m at the end of each pulse. No mitochondrial [Ca2+] oscillations were observed. These data indicate that repetitive increases of [Ca2+]em caused a net accumulation of Ca2+ into mitochondria with each [Ca2+]em pulse. The absence of oscillations of [Ca2+]m could be explained by 1) the pulse duration not being long enough to raise [Ca2+]m high enough to stimulate fast uptake of Ca2+ or 2) the frequency of stimulation not allowing enough time for NCXm to remove Ca2+ efficiently.
Effects of Ca2+ pulse duration and stimulation frequency on mitochondrial Ca2+ uptake. To evaluate how Ca2+ pulse duration and/or frequency affect mitochondrial Ca2+ accumulation in cardiomyocytes, we first applied a train of Ca2+ pulses at a constant frequency (0.5 Hz) but variable pulse duration (0.2, 0.5, and 1 s). Figure 7A shows that the longer pulses (1 s) allowed mitochondrial Ca2+ to rise faster and to a higher level compared with the shorter pulses (0.2 and 0.5 s), but still no oscillatory behavior was observed. Cell stimulation with a higher frequency (1 Hz) but constant (0.5 s) pulse duration also resulted in an increased mitochondrial Ca2+ accumulation compared with lower-frequency stimulation (0.25 Hz; Fig. 7B). Depending on frequency and duration of pulses, [Ca2+]m rose to new steady-state levels where a new equilibrium between mitochondrial Ca2+ uptake and removal was reached. The net Ca2+ movements across the inner membrane during an individual Ca2+ pulse of physiologically relevant duration and frequency appear to be small, thus minimizing energetic costs of mitochondrial Ca2+ shuttling.
Mitochondrial Ca2+ uptake from continuous vs. pulsed elevations of [Ca2+]em. In the next series of experiments, we investigated whether mitochondrial Ca2+ uptake would be different in cells exposed to elevated [Ca2+]em continuously or in a repetitive pulsatile fashion. Figure 8A, top, shows a representative record of mitochondrial Ca2+ accumulation under conditions in which the cell was exposed to elevated [Ca2+]em ([Ca2+]pip = 100 µM) continuously. In another cell, Ca2+ pulses ([Ca2+]pip = 100 µM) of 0.5-s (Fig. 8A, bottom) or 1-s (not shown) duration were applied at a frequency of 0.5 Hz. Mitochondrial Ca2+ accumulation was measured for every second of total exposure time to elevated [Ca2+]em. The data are summarized in Fig. 8B and show that there was no difference in mitochondrial Ca2+ uptake whether cells were exposed to continuously elevated [Ca2+]em or repetitive Ca2+ pulses for the same cumulative amount of time.
Together, our data support the conclusion that in cardiomyocytes mitochondrial Ca2+ transport is activated during continuous or pulsatile elevations of [Ca2+]i above a threshold level; however, they are inconsistent with a beat-to-beat transmission of [Ca2+]i transients to the mitochondrial matrix, resulting in [Ca2+]m oscillations during each cycle of E-C coupling.
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DISCUSSION |
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-adrenergic stimulation of cells led to a slow rise of [Ca2+]m from 100 to 500–600 nM, but without any obvious beat-to beat changes in [Ca2+]m (15, 16, 23, 37), and that mitochondria in intact ferret and cat ventricular cells did not take up detectable amounts of Ca2+ during individual contractions (53). Evidence in support of a beat-to-beat translation of [Ca2+]i transients into [Ca2+]m oscillations (model II) was first observed in guinea pig myocytes with combined whole cell patch-clamp recording, microfluorometry, and EPMA (31, 52); however, other studies using EPMA were not able to resolve fast changes in total [Ca2+]m (see, e.g., Ref. 38). Further support for fast [Ca2+]m transients came from studies using laser scanning confocal microscopy in combination with fluorescent dyes to follow [Ca2+]m changes in rabbit ventricular myocytes (7, 39). In these experiments cells were loaded with fluo-3 AM or indo-1 AM, which accumulated in both compartments, the cytosol and mitochondria. Mitochondria were identified by costaining with voltage-sensitive probes tetramethyl rhodamine methyl ester or rhodamine 123. Recordings obtained during electrical or isoproterenol stimulation revealed [Ca2+] transients with identical time courses in both compartments. However, the lack of kinetic differences between the two signals raises the possibility that the mitochondrial signal was "contaminated" with cytosolic signal to a significant degree. Nonetheless, Mackenzie et al. (36) were able to record with the fluorescent indicator rhod-2 (an indicator that is often used as a probe to measure specifically [Ca2+]m) [Ca2+]m transients that were abolished by the mitochondrial inhibitors antimycin and oligomycin, whereas under identical experimental conditions [Ca2+]i transients could still be evoked.
To avoid the problem of signal contamination mentioned above, Robert at al. (42) applied a different experimental approach. They used genetically encoded targeted Ca2+ probes to explore beat-to-beat transmission of Ca2+ between RyR and mitochondria. Using the Ca2+-sensitive photoprotein aequorin and novel green fluorescent protein-based Ca2+ indicators termed ratiometric-pericams specifically targeted to the mitochondria, cytosol, and/or nucleus, they demonstrated that spontaneous [Ca2+]i oscillations in neonatal cultured cardiomyocytes were followed by [Ca2+]m oscillations. Elevation of extracellular [Ca2+] from 1 to 2 or 4 mM or stimulation of -adrenergic receptors with isoproterenol resulted in a substantial increase in spike amplitude in both compartments and increased basal [Ca2+] level (interspike level) in mitochondria, but no effect on diastolic cytosolic [Ca2+] was observed. Similar observations were made with the muscarinic receptor agonist carbachol. Moreover, the frequency of [Ca2+] oscillations observed in mitochondria was different from the frequency of [Ca2+]i oscillations. Together, these observations indicate that the rate of Ca2+ extrusion from mitochondria was slower than mitochondrial Ca2+ uptake, which resulted in incomplete Ca2+ extrusion and an elevation of diastolic [Ca2+]m, especially under conditions of elevated [Ca2+]i.
Szalai et al. (51) showed that activation of RyRs by Ca2+, ryanodine, and low concentrations of caffeine evoked [Ca2+]i oscillations that were synchronized with oscillations of [Ca2+]m in permeabilized cardiac H9c2 myotubes. These [Ca2+]m oscillations were due to activation of mitochondrial Ca2+ uptake through the Ca2+ uniporter and subsequent fast removal of Ca2+ by mitochondrial Ca2+ exchangers, with little contribution from the permeability transition pore. However, the frequency of [Ca2+]m oscillations observed in this study were slower than a typical physiological heart rate by nearly an order of magnitude. Furthermore, during oscillations [Ca2+]m did not always return to the prespike level after each spike. These data suggest that most probably the rate of mitochondrial Ca2+ removal through exchangers was not fast enough to extrude all Ca2+ that entered during a Ca2+ spike before the beginning of the next cytosolic spike. In our experiments we were also able to observe [Ca2+]m transients (data not shown), but only under the conditions in which the interval between two individual elevations of [Ca2+]i exceeded tens of seconds and not at stimulation frequencies between 0.25 and 1 Hz (Fig. 7). Altogether we tested a broad range of intervals between pulses (0.5–3.5 s) that covered typical cardiac beating rates. This range of stimulation intervals was tested under conditions that favored Ca2+ extrusion (Na+ present) as well as under conditions in which Ca2+ extrusion was blocked (Na+ free). Under all these conditions the interval between pulses was too short for substantial Ca2+ extrusion to occur and for setting up [Ca2+]m oscillations. As shown in Fig. 5, Ca2+ extrusion occurred on a much slower timescale.
As described above, all methodological approaches used so far had failed to reach a unequivocal conclusion as to whether mitochondria of cardiomyocytes respond to [Ca2+]i oscillation in a beat-to-beat fashion. Therefore, in the present study we sought to address this question again with a novel experimental approach.
Here we developed a novel experimental approach that allowed us to simulate fast [Ca2+]i transients in membrane-permeabilized cells. Permeabilized cells have the unique advantage that the cytosolic environment can be controlled precisely while the arrangements and interaction between intracellular membranes and organelles (SR, mitochondria) remain structurally and functionally intact (19, 45). To measure mitochondrial Ca2+ signals we took advantage of the fact that the acetoxymethyl ester form of fluorescent Ca2+ indicators sequesters into intracellular organelles, including mitochondria. For our experiments we exposed intact cat ventricular cardiomyocytes to fluo-3 AM and subsequently used digitonin to remove fluo-3 from the cytosol and nucleus (Fig. 1). Colocalization experiments with the mitochondrial probe MitoTracker Red confirmed the mitochondrial origin of the fluo-3 signal after permeabilization (Fig. 1B). Control experiments indicated that the elevation of [Ca2+]em to >0.5 µM resulted in mitochondrial Ca2+ accumulation (Fig. 2A). Mitochondrial Ca2+ accumulation was mediated by Ca2+ uptake through MCU, because the signal was sensitive to the MCU blocker RR and the protonophore FCCP (Fig. 3). Ca2+ entry via the uniporter exhibited a sigmoid dependence on [Ca2+]em and under physiological ionic conditions reached k0.5 for Ca2+ accumulation at [Ca2+]em = 4.4 µM (Fig. 2B). Mitochondrial Ca2+ uptake was not affected by the blocker of the SR Ca2+ pump thapsigargin (2 µM; n = 3; data not shown), suggesting that the fluo-3 signals recorded after permeabilization were not contaminated by contributions from dye entrapped in the SR and changes in SR Ca2+ content.
The experiments illustrated in Fig. 3A show that mitochondria of cat ventricular myocytes extrude Ca2+ exclusively via Na+/Ca2+ exchange because removal of Na+ from extramitochondrial solution completely prevented Ca2+ extrusion. This observation suggests that no significant Na+-independent Ca2+ extrusion occurred. This finding is in line with the notion that the Na+-dependent mechanism is thought to be the predominant Ca2+ extrusion pathway in cardiac mitochondria (for review see, e.g., Refs. 24 and 27); however, the coexistence of Na+-independent efflux has been suggested (11, 43). The rate of Ca2+ extrusion depended on [Na+]em, with a maximal rate observed in the range of 20–40 mM. In addition, removal of Na+ from the extramitochondrial solution led to significant increase (2-fold) in mitochondrial Ca2+ accumulation (Fig. 5A, left) compared with Ca2+ uptake in the presence of 40 mM [Na+]em (Fig. 5A, right). These data suggest that NCXm actively counteracted Ca2+ uptake; however, the rate of Ca2+ extrusion through NCXm was approximately two times slower than mitochondrial Ca2+ uptake. The cytosolic Na+ concentration ([Na+]i) dependence of NCXm is sigmoidal, with a half-maximal activity at 4–8 mM (3, 8, 21, 46). These values are close to experimentally measured resting [Na+]i observed under physiological conditions in cardiomyocytes (see, e.g., Refs. 4 and 14), making the mitochondrial Na+/Ca2+ exchange potentially sensitive to physiological fluctuations in cytosolic [Na+]i (for review, see Ref. 3). However, no significant variations in bulk [Na+]i were observed during a normal cardiac cycle, and only the substantial increase in the frequency of electrical stimulation resulted in a significant change of bulk cytoplasmic [Na+] (13). Furthermore, it has remained elusive and controversial (see, e.g., Refs. 27 and 30) whether the NCXm is an electroneutral or electrogenic antiporter. If the exchanger is electrogenic, Ca2+ extrusion would tend to depolarize the mitochondrial membrane potential, which in turn would accelerate extrusion by reducing the electrical gradient responsible for Ca2+ uptake and for retaining mitochondrial Ca2+. Together, these data suggest that mitochondrial Na+/Ca2+ exchange can actively modulate mitochondrial Ca2+ uptake and extrusion; however, significant changes in cytosolic Na+ levels are required for such a mechanism. Evidence is lacking that such fluctuations occur during a normal cardiac cycle; however, under pathological conditions large changes in [Na+]i were observed (33, 47).
We found (Fig. 4) that elevation of [Pi] significantly increased mitochondrial Ca2+ uptake. Although overall mitochondrial Ca2+ uptake was higher at higher [Ca2+]em (1 µM) compared with low [Ca2+]em (0.5 µM), the relative difference in Ca2+ uptake between low and high [Pi] was more pronounced at low [Ca2+]em (Fig. 4C). Pi forms a complex with Ca2+ in the mitochondrial matrix, thereby decreasing the free [Ca2+]m and increasing the chemical concentration gradient for Ca2+ uptake (26, 54). Furthermore, Pi uptake increases and facilitates mitochondrial Ca2+ uptake by enhancing the electrical gradient (10, 40). The regulation of mitochondrial Ca2+ uptake by Pi might play an important role in pathological conditions, such as ischemia-reperfusion, where high-energy phosphate bonds are hydrolyzed, [Pi] increases, and cytosolic [Ca2+] is high.
Mitochondrial decoding of fast [Ca2+]i transients. After we established functional mitochondrial Ca2+ uptake and extrusion in our permeabilized myocyte model, we applied specific experimental protocols that were aimed to imitate, in permeabilized cells, beat-to-beat [Ca2+]i transients as they normally occur during E-C coupling in intact cardiac myocytes (Fig. 6). The results of our study indicate that rapid switching (0.25–1 Hz) of [Ca2+]em to high levels ([Ca2+]pip = 100 µM) simulated rapid beat-to-beat changes in [Ca2+]i (with [Ca2+] transient durations of 100–500 ms) but did not lead to [Ca2+]m oscillations. There was no difference in response phenotype whether Ca2+ extrusion via NCXm was enabled (Fig. 7; [Na+]em = 20 mM) or blocked (Fig. 8; [Na+]em = 0). The slow frequency-dependent increase of [Ca2+]m disproved a rapid transmission of Ca2+ signals between cytosol and mitochondria. Comparison of [Ca2+]m changes in response to continuous exposures to elevated [Ca2+]em and to a train of brief [Ca2+]em pulses of the same cumulative duration (Fig. 8) revealed no difference in Ca2+ uptake. Our data suggest that in permeabilized cardiac myocytes fast [Ca2+]i transients are integrated by mitochondrial Ca2+ transport systems resulting in a frequency-dependent net accumulation of Ca2+ in the matrix (model I). These small, gradual changes in [Ca2+]m that accompany changes in heart rate or cellular Ca2+ load may alter matrix dehydrogenase activities and subsequently may help to regulate mitochondrial energy production. Four key mitochondrial matrix dehydrogenases are activated by low micromolar [Ca2+] (glycerol 3-phosphate dehydrogenase, pyruvate dehydrogenase, NAD-linked isocitrate dehydrogenase, and 2-oxoglutarate dehydrogenase) (28). Thus increases in mitochondrial Ca2+ via the above mechanisms could occur when [Ca2+]i is relatively high concomitant with high energy demands (i.e., when contractile activation and Ca2+ pumping are consuming ATP at high rates). With mitochondrial NADH concentration ([NADH]m)-dependent autofluorescence as an index of dehydrogenase activity in intact contracting ventricular muscle, [Ca2+]i-dependent stimulation of mitochondrial NADH production was demonstrated (5). With a sudden increase in stimulation frequency or extracellular Ca2+ there was a transient decrease in [NADH], consistent with NADH production not keeping up with the increased ATP and NADH consumption. However, this [NADH] decline was followed by a recovery toward previous levels. This recovery was entirely dependent on increased average [Ca2+]i. It was concluded that the increased average [Ca2+]i caused an increase in [Ca2+]m and stimulation of dehydrogenases and NADH production. The kinetics of the measured [NADH]m changes in response to increased pacing frequency did not require a rapid transmission of [Ca2+]i signals into the mitochondrial matrix. In fact, the time course of the delayed [NADH]m recovery was reminiscent of the slow changes in [Ca2+]m reported by Miyata and coworkers (37) under comparable conditions.
In conclusion, cat ventricular myocytes do not shuttle Ca2+ between cytosol and mitochondria on a beat-to-beat basis in ways that would lead to [Ca2+]m oscillations. However, with slower increases of basal [Ca2+]i, mitochondrial Ca2+ transport and Ca2+ accumulation can be important for stimulation energy metabolism to meet cellular metabolic demands. In addition, the relatively slow kinetics of mitochondrial Ca2+ uptake still allow mitochondria to function as a temporary storage compartment during severe Ca2+ overload and to help protect the cytoplasm from very high Ca2+ levels.
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
* M. Sedova and E. N. Dedkova contributed equally to this work. 
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