Control of mitochondrial motility in the physiological range of global [Ca²⁺]_c

When EGTA and Iono were added together to nonstimulated cells to lower the [Ca²⁺]_c no change in mitochondrial movement activity was observed (n = 4; unpublished data), suggesting that the motility was maximal at the resting level of [Ca²⁺]_c (50–100 nM). To quantitate the [Ca²⁺]_c dependence of the inhibition of mitochondrial motility, [Ca²⁺]_c and motility were measured in cells that were incubated in a Ca²⁺ free buffer supplemented with EGTA, thapsigargin (Tg), an inhibitor of the sarco-endoplasmic reticulum Ca²⁺ pump and Iono to ensure rapid equilibration of the cytosol with the extracellular [Ca²⁺], and then varying amounts of CaCl₂ were added (Fig. 2, A and B). As shown in Fig. 2 A, stepwise increases in [Ca²⁺]_c were accompanied with stepwise decreases in mitochondrial motility. The inhibition of motility was plotted against the [Ca²⁺]_c (Fig. 2 B, filled circles). The majority of the Ca²⁺-induced attenuation of mitochondrial motility was obtained in the sub-micromolar range of [Ca²⁺]_c (IC₅₀ ≈ 400 nM), indicating that mitochondrial motility is controlled in the physiological range of [Ca²⁺]_c.

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Figure 2.

Relationship between [Ca ²⁺ ] _c and mitochondrial motility. (A) Stepwise increases in [Ca²⁺]_c induce stepped decreases in mitochondrial motility. MitoYFP-expressing and fura2-loaded cells were incubated in Ca²⁺-free ECM supplemented with 2 mM EGTA, 2 μM Tg, and 10 μM of Iono for 12 min. Simultaneous measurements of [Ca²⁺]_c and mitochondrial motility were performed in single cells exposed to stepped increases in extracellular CaCl₂ from 0 to 15 mM. [Ca²⁺]_c was calibrated in terms of nanomoles using in vitro calibration of fura2. (B) Dose–response relationship between [Ca²⁺]_c and mitochondrial motility. Simultaneous measurements of [Ca²⁺]_c and mitochondrial motility were performed in single cells as described for A. Mean mitochondrial motility inhibition and mean [Ca²⁺]_c were calculated for each [CaCl₂]. Mean ± SEM are marked by filled circles (n = 68 cells from 40 experiments). The IC₅₀ ≈ 400 nM, indicating that mitochondrial motility is controlled in the physiological range of [Ca²⁺]_c. (C) Mitochondrial motility (red line) and [Ca²⁺]_c (black line) were recorded in single cells stimulated by varying doses of VP (0.1–100 nM). In addition, the mean mitochondrial motility inhibition and mean [Ca²⁺]_c were calculated for each VP concentration and are plotted in B (empty circles and dashed line; n = 44 cells from 28 experiments).

In VP-stimulated cells, the inhibition of mitochondrial motility was also in proportion to the agonist dose and the size of the [Ca²⁺]_c elevation (Fig. 2 C). Low doses of VP (e.g., 0.1 nM) were sufficient to cause considerable inhibition of the movement (Fig. 2 C). When the VP-induced inhibition of motility was plotted against the VP-induced [Ca²⁺]_c rise, the data points were close to the curve obtained by the in situ [Ca²⁺]_c titration (Fig. 2 B, filled circles). In other words, the competence of VP to attenuate mitochondrial movement was in proportion to the [Ca²⁺]_c signal. Thus, inhibition of mitochondrial movement during IP₃-mediated Ca²⁺ mobilization can be attributed to the [Ca²⁺]_c signal.

No role for mitochondrial Ca²⁺ uptake and mitochondrial membrane potential (ΔΨ_m) in the arrest of mitochondrial motility by Ca²⁺

The IP₃-induced [Ca²⁺]_c signal is propagated to the mitochondria, giving rise to a mitochondrial matrix [Ca²⁺] ([Ca²⁺]_m) rise that controls the activity of Ca²⁺-dependent enzymes and ion channels (for review see Duchen, 2000). The primary driving force of the mitochondrial Ca²⁺ uptake is the ΔΨ_m that also drives mitochondrial ATP synthesis. To determine whether the [Ca²⁺]_m signal or ΔΨ_m is necessary for the Ca²⁺-dependent inhibition of mitochondrial motility, we treated cells with an uncoupler (FCCP added with the ATPase inhibitor, oligomycin to preserve cytosolic ATP; Fig. 3 A). This treatment has been shown to cause rapid dissipation of the ΔΨ_m (Pacher and Hajnóczky, 2001) and to reduce mitochondrial Ca²⁺ uptake in H9c2 cells (Szalai et al., 2000). Uncoupler induced a slow attenuation in mitochondrial motility (Fig. 3 A), which may reflect a fall in perimitochondrial ATP that is likely to be needed for movement. However, the VP-induced arrest of mitochondrial movement was preserved in uncoupler-pretreated cells (Fig. 3 A). Thus the [Ca²⁺]_m signal and ΔΨ_m appears to be dispensable for the Ca²⁺-mediated inhibition of mitochondrial movement.

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Figure 3.

Spatio-temporal pattern of the calcium signal and inhibition of mitochondrial motility. (A) Calcium signal propagation to the mitochondria is not required for the inhibition of motility. Uncoupler (5 μM FCCP and 5 μg/ml Oligo; solid line) or solvent (dashed line) was added to the cells 5 min before stimulation with VP. The uncoupler causes dissipation of the ΔΨ_m in <1 min and in turn decreases the driving force of the mitochondrial Ca²⁺ uptake in H9c2 myoblasts (Szalai et al., 2000). Mitochondrial motility showed a slowly developing decrease (red solid line) compared with the control (red dash line). However uncoupler did not prevent the effect of VP on mitochondrial motility, indicating that mitochondrial Ca²⁺ uptake and ΔΨ_m were not necessary for the control of motility by the calcium signal. In another set of experiments, the cells were pretreated with uncoupler for 10 min before the recording was started. The inset shows that the residual motility was effectively inhibited by 100 nM VP. Data are the means of nine experiments with uncoupler and four experiments for the control. (B) Reversible and reproducible inhibition of mitochondrial motility by Ca²⁺. To evaluate the effect of pulsatile increases of [Ca²⁺]_c (black traces) on mitochondrial motility (red traces), the 10 μM of Iono-induced [Ca²⁺]_c elevation was recorded for 10 min (solid lines) or was reversed by addition of 5 mM EGTA and, subsequently, was reestablished by addition of 10 mM CaCl₂ (dashed lines). Data are the means of 36 cells from 22 experiments.

Spatio-temporal control of the mitochondrial movement by [Ca²⁺]_c oscillations and waves

Our data have shown that the control of mitochondrial motility can respond to repetitive stimulation by an IP₃-linked agonist (Fig. 2 C). Along this line, Fig. 3 B shows that the [Ca²⁺]_c rise-induced inhibition of mitochondrial movement can be reversed by termination of the [Ca²⁺]_c elevation and subsequently, reproduced by a second step of [Ca²⁺]_c elevation. This suggests that no desensitization of the Ca²⁺ regulation of mitochondrial motility occurred. If there is no desensitization and the reversal of the movement inhibition is slower than the decay of the [Ca²⁺]_c spike, [Ca²⁺]_c oscillations may be able to cause a prolonged depression of mitochondrial motility. In differentiated H9c2 myotubes, RyR-mediated [Ca²⁺]_c oscillations have been demonstrated (Szalai et al., 2000). As shown in Fig. 4, [Ca²⁺]_c oscillations were also observed in H9c2 myoblasts transfected with RyR1 (Bhat et al., 1997) and stimulated with caffeine (Caff). Similar to the IP₃-linked [Ca²⁺]_c spikes, the RyR-mediated [Ca²⁺]_c spikes also triggered inhibition of mitochondrial motility and the oscillations of [Ca²⁺]_c were often associated with oscillations in movement activity (Fig. 4 A, cell marked by an arrow). Each burst of RyR1-mediated Ca²⁺ release could result in maximal inhibition of mitochondrial motility (Fig. 4, A and B). Isolated spikes of mitochondrial movement inhibition were observed during low frequency [Ca²⁺]_c oscillations (unpublished data). However, if the frequency of [Ca²⁺]_c spiking was higher and the recovery of motility was slow, [Ca²⁺]_c oscillations could produce an essentially sustained maximal inhibition in movement activity (Fig. 4 B). Thus the frequency-modulated [Ca²⁺]_c signals are translated into a time-averaged motility response. Notably, at supramaximal stimulation the [Ca²⁺]_c oscillations run together and fuse into a large and slowly decaying single [Ca²⁺]_c spike (Szalai et al., 2000), but this [Ca²⁺]_c signal could not provide for sustained inhibition of the mitochondrial motility (e.g., Fig. 1 B). In this way, the control of mitochondrial motility may serve as a model whereby [Ca²⁺]_c oscillations are an effective signal for long-term modulation, but a nonoscillatory [Ca²⁺]_c signal is unable to maintain inhibition.

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Figure 4.

Control of mitochondrial motility by [Ca ²⁺ ] _c oscillations and waves. Mitochondrial motility was evaluated simultaneously with [Ca²⁺]_c in cells cotransfected with constructs encoding mitoYFP and RyR1 and loaded with fura2. To promote RyR-mediated Ca²⁺ mobilization, the cells were exposed to 10 mM of caffeine (Caff). (A) In the image series, mitochondrial movements are visualized in a cell that showed [Ca²⁺]_c oscillations in response to stimulation by Caff (arrow) and in a cell that did not show a Caff-induced [Ca²⁺]_c rise (top left corner) as described for Fig. 1. Also shown is the effect of 100 nM VP that elicited a [Ca²⁺]_c signal in both cells. The time course of [Ca²⁺]_c (black trace) and mitochondrial motility (red trace) for the Caff-sensitive cell is plotted. (B) Sustained inhibition of mitochondrial motility in a cell that displayed a relatively high frequency [Ca²⁺]_c oscillation in response to stimulation by Caff. (C) Calcium waves were induced in mitoYFP-expressing cells by treatment with 75 μM thimerosal (TM) and 0.1 nM VP. The image series shows the mitoYFP fluorescence (i) and the first two calcium waves after addition of VP and TM (t = 60 s; ii–ix). The first wave begins simultaneously at the ends of the cell and converges in the center as indicated by the arrows (iv). The second wave begins at the lower end of the cell and weakens as it propagates to the top (vi–viii, arrow marks the direction of the wave propagation in vii). Measurement of [Ca²⁺]_c (x) and mitochondrial movement (ix) in three distinct regions of the cell, labeled 1–3 in panel ii, reveals spatio-temporal heterogeneity in the inhibition of movement corresponding to the local calcium concentration.

The [Ca²⁺]_c signal often displays regional heterogeneity that may also result in heterogeneity in the mitochondrial motility. To test this possibility, we studied mitochondrial motility in cells that showed spatial heterogeneity in the [Ca²⁺]_c rise. In the cell shown in Fig. 4 C, the first [Ca²⁺]_c wave started simultaneously at the ends of the cell (regions 1 and 3) and converged in the center (region 2) as indicated by the arrows (iv). The [Ca²⁺]_c rise was relatively large in the center (Fig. 4 C, x). Reproducing the spatial pattern of the [Ca²⁺]_c signal, inhibition of the mitochondrial motility was substantially larger in the center than at the ends of the cell (Fig. 4 C, ix). The second [Ca²⁺]_c wave began at the lower end of the cell and weakened as it propagated to the top (Fig. 4 C, vi–viii). Similarly, the inhibition of mitochondrial motility was substantial at the lower end, smaller in the center, and hardly noticeable at the upper end (Fig. 4 B, ix). This experiment revealed intracellular spatio-temporal heterogeneity in the inhibition of movement corresponding to the local [Ca²⁺]_c concentration. Such differences in the movement may affect the spatial distribution of mitochondria. Mitochondria would be stopped in the vicinity of the active Ca²⁺ release sites. If mitochondria stay in high [Ca²⁺]_c areas for a relatively long time and cross quickly through low [Ca²⁺]_c areas, this also provides a mechanism for the redistribution of mitochondria to the regions of high [Ca²⁺]_c. Notably, one or two localized [Ca²⁺]_c spikes did not result in an obvious increase in the mitochondrial density (unpublished data). Acquisition of mitochondria from low [Ca²⁺]_c zones may require prolonged or repetitive Ca²⁺ release because it would not be driven by an attractive force but would use immobilization of mitochondria that move into the region of the [Ca²⁺]_c rise on their own. However, stabilization of the position of the mitochondria in the areas of Ca²⁺ release seems to be sufficient to promote the delivery of [Ca²⁺]_c spikes to the mitochondria (see Arrest of the mitochondria supports rapid Ca²⁺ delivery to the mitochondria). Thus, through the control of motility, the spatial distribution of the mitochondrial Ca²⁺ buffering and ATP production may be dynamically regulated by the local [Ca²⁺]_c.

Organization of mitochondrial motility by the MTs

Long-range mitochondrial movements are facilitated by the MTs and certain forms of local movement may use the MFs (Couchman and Rees, 1982; Nangaku et al., 1994; Morris and Hollenbeck, 1995; Varadi et al., 2004). In H9c2 myoblasts expressing tubulinGFP and DsRed targeted to the mitochondria, green fluorescence both marked a complex network of fibers and appeared as a homogeneous signal throughout the cells (Fig. 5 A, i). Labeling of the fibers became more intense, and the homogeneous signal effectively disappeared when the cells were pretreated with taxol, a drug that promotes the polymerization of tubulin (Fig. 5 A, ii). The fibers were retained and the homogeneous signal was promptly eliminated if the cells were permeabilized (Fig. 5 A, x). Based on these data, the fibers represent the MTs and the homogeneous fluorescence is likely to be accounted by monomeric tubulinGFP.

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Figure 5.

Spatial relationship between MTs, MFs, and mitochondria. (A) H9c2 cells expressing tubulinGFP and mitoDsRed and incubated in the absence (i) or presence (ii) of 10 μM of an MT-stabilizing agent, taxol. (iii–v) Magnified time-lapse images of a peripheral region of the taxol-pretreated cell. Arrowheads mark the mitochondria that move substantially from the previous image. (vi–ix) Further magnified region showing a single mitochondrion sliding along an MT. (x and xi) A naive and a nocodazole-pretreated cell after permeabilization with digitonin. (B) Inhibition of mitochondrial motility and enhancement of the VP-induced mitochondrial [Ca²⁺] signal in nocodazole-pretreated cells. (top left) Mitochondrial motility and [Ca²⁺]_c in nocodazole-treated cells (solid lines, 23 cells in 10 measurements) as compared with control cells (dashed lines, 22 cells in 11 measurements). The effect of 100 nM VP is also shown. (top right) Resting [Ca²⁺]_c and the peak value of the VP-induced [Ca²⁺]_c signal in control and nocodazole-pretreated (10 μM for 25–30 min) cells. (bottom) Nuclear matrix and mitochondrial matrix [Ca²⁺] measured in cells expressing both nuclear and mitochondrial pericam using fast, ratiometric imaging (2 ratio/s). Cells were pretreated with nocodazole (10 μM for 25–30 min) or solvent (control) and stimulated by 100 nM VP. The VP-induced initial [Ca²⁺]_c signal was measured (change in nuclear pericam fluorescent ratio at the first point of the [Ca²⁺] rise, <25% of the maximal change) and the corresponding change in [Ca²⁺]_m (change in mitochondrial pericam fluorescent ratio) was calculated for each cell (mean ± SEM; n = 16). (C) MFs and mitochondria in H9c2 cells expressing actinGFP and mitoDsRed. Cells were incubated in the absence or presence of 10 μM of nocodazole.

Distribution of the red fluorescence showed colinearity of the long axis of tubular mitochondria and MTs (Fig. 5 A, i–iii and x; Video 2, available at http://www.jcb.org/cgi/content/full/jcb.200406038/DC1). Furthermore, the time series of images shows that mitochondria move along MTs (Fig. 5 A, iii–v, arrows point to the sites of movement). Similar to the mitochondrial movement shown in a taxol-pretreated cell (Fig. 5 A, vi–ix), in control cells, the average rate was 0.31 ± 0.05 μm/s (n = 19) for mitochondria that showed continuous movement along a straight track for ≥10 s. The dynamics of mitochondrial motility along the MTs is further illustrated by a movie sequence (Video 2). To quantify the MT-associated movement, for each mitochondrion that moved ≥3.5 μm, it was evaluated whether or not the movement was along an MT. In four cells, 49 out of 68 mitochondria (72%) followed the track of MTs and no mitochondrion (0%) moved clearly independent of the MTs. For the remaining 17 mitochondria (28%), it was not possible to discern if the movement was along an MT, due to the homogeneous tubulinGFP fluorescence in some regions of the cells. Exposure of the cells to nocodazole (10 μM), a drug disrupting the tubulin polymers resulted in loss of the MTs without inducing a major change in mitochondrial morphology (Fig. 5 A, x and xi). However, mitochondrial motility displayed a progressive decrease in nocodazole-treated cells (Fig. 5 B). Strikingly, the nocodazole-induced inhibition of mitochondrial motility occurred in the absence of a [Ca²⁺]_c elevation (Fig. 5 B). However, nocodazole evoked a decrease in ER Ca²⁺ storage because the VP-induced Ca²⁺ mobilization was smaller in nocodazole-pretreated cells (Fig. 5 B, histograms).

In cells expressing actinGFP and mitoDsRed, the green fluorescence showed fibers (Fig. 5 C, left) that were also labeled with rhodamine-phalloidin, an MF-specific tracer (n = 4; not depicted). Although the red fluorescent mitochondria often appeared close to the MFs, usually mitochondria were not aligned to the MFs (Fig. 5 C, left). In the time series of images, the movement of the mitochondria along the MFs was quantified as described for the movement along the MTs above. In four cells, only 10 out of 66 mitochondria (15%) followed the track of an MF, whereas 37 mitochondria (56%) appeared to move independent of the MFs. In the remaining 19 cases (29%), it was not possible to discern whether or not the movement was along an MF. In regard to the role of MFs in mitochondrial motility it is also relevant that the MFs were undamaged in nocodazole-treated cells (Fig. 5 B, right), which did not display considerable mitochondrial movement activity (Fig. 5 B). Collectively, the aforementioned data suggest that mitochondrial movement depends on the integrity of the MTs and closely follows the tracks provided by the MTs. Furthermore, MFs alone do not appear to provide the primary track for mitochondrial movements.

In the movement of mitochondria along MTs, cytoplasmic dynein and kinesin (Kif1b)-based motors are likely to play a role. Mitochondrion-specific kinesin heavy and light chains have been identified in mammalian cells (Nangaku et al., 1994; Khodjakov et al., 1998). However, recent resolution of the molecular structure of the mammalian cytoplasmic dynein and kinesin did not reveal any Ca²⁺ or CaM binding site (for review see Vale, 2003). Furthermore, we have observed that neither inhibitors of the Ca²⁺/CaM-dependent kinases (KN-62, 10 μM, n = 5; KN-93, 5 μM, 30 min pretreatment, n = 5; myristoylated-autocamtide-2–related inhibitory peptide, 10 μM, 1 h pretreatment, n = 5) nor inhibitors of calcineurin, the Ca²⁺-dependent protein phosphatase (cyclosporine A, 5 μM, n = 3; FK506 10 μM, 30 min, n = 5; deltamethrin 10 μM, 30 min pretreatment, n = 5) prevented the VP-induced inhibition of mitochondrial motility. Based on these data, the Ca²⁺ signal is not likely to control mitochondrial movement through phosphorylation or dephosphorylation of dynein or kinesin. Thus, it seems that a distinct Ca²⁺ sensor molecule is required to translate the Ca²⁺ signal for the microtubular motor proteins.

Transient expression

When the cell culture reached 70% confluency (2 d after plating), transfection was performed using Lipofectamine 2000 (Invitrogen) according to the manufacturer's instructions with 2 μg/ml of each vector. One or two of the vectors that encode pEYFP-mito, pDsRed-mito, pEGFP-tubulin, pEGFP-actin (BD Biosciences), or ratiometric-pericams targeted to the cytoplasm, nucleus, or mitochondria (provided by A. Miyawaki, RIKEN, Wako City, Japan) or RyR1 (a gift from J. Ma, University of Medicine and Dentistry of New Jersey, Piscataway, NJ) were used. Transfected cells were further incubated in the culture for 24–36 h before the imaging experiments.

Live cell imaging

Fluorescence imaging was performed using an inverted microscope (model IX70; Olympus; 40×, UApo340, NA 1.35) fitted with a cooled CCD camera (Pluto; Pixelvision) and a high speed wavelength switcher (model Lambda DG; Sutter Instruments) controlled by Spectralyzer (custom) software. Simultaneous detection of fura2 (340 and 380 nm excitation) and mitoYFP (495 nm excitation) fluorescence was achieved using a multiwavelength beamsplitter/emission filter combination (Chroma Technology Corp.). Pericam was excited at 415 and 495 nm. The data acquisition rate was 0.4 triplet per second in the fura2 and mitoYFP imaging and 2 duplet per second in the pericam imaging experiments. To correct for the camera noise, the dark (excitation path closed) image was obtained and was subtracted from each image.

Confocal imaging of the fluorescent proteins targeted to the mitochondria, MFs, and MTs was performed using an imaging system (model Radiance 2100; Bio-Rad Laboratories) equipped with a Kr/Ar-ion laser source (488 and 568 nm excitation) fitted to an inverted microscope (40×, UApo340, NA 1.35). Mitotracker green, GFP, and YFP were excited at 488 nm, and DsRed at 568 nm. Images were obtained every 3.1–3.3 s using LaserSharp software (Bio-Rad Laboratories).

To simultaneously measure [Ca²⁺]_c with mitochondrial motility, mitoYFP-transfected cell cultures were preincubated for 20 min in an extracellular medium (ECM; 2% BSA, 121 mM NaCl, 5 mM NaHCO3, 10 mM Na-Hepes, 4.7 mM KCl, 1.2 mM KH₂PO4, 1.2 mM MgSO₄, 2 mM CaCl₂, and 10 mM glucose, pH 7.4), and then loaded with 5 μM fura2/AM for 25–30 min at RT in the presence of 0.003% (wt/vol) pluronic acid and 200 μM sulfinpyrazone. After washing of the dye-loaded cells, imaging measurements were performed in ECM containing 0.25% BSA at 35°C. Imaging of pericam, mitoYFP, actinGFP+mitoDsRed, and tubulin-GFP+mitoDsRed was also performed in 0.25% BSA-ECM at 35°C.

To calibrate the changes in [Ca²⁺]_c, the ratio of 340/380 nm fluorescence values was calculated for the fura2-loaded cells after subtraction of background fluorescence. The ratio was converted to nanomole values using in vitro calibration of fura2-free acid (K_d = 224 nM). To quantitate mitochondrial motility a difference image protocol was used. By subtraction of sequential images, the fluorescence change for each pixel was calculated, and then pixels that exhibited a change after 3 × 3 median filtering (positive or negative) greater than an empirically determined threshold (2.5% of the mean fluorescence intensity/pixel) were counted for each time point. Changes in the pixel number were normalized to the initial value calculated for cells before stimulation.

All image analysis was done in Spectralyzer imaging software. Experiments were performed with ≥3 different cell preparations. Traces represent single cell responses unless indicated otherwise. Data are presented as means ± SEM. Significance of differences from the relevant controls was calculated by t test.

The authors thank Drs. Jianjie Ma and Atsushi Miyawaki for plasmid DNA and Dr. Gabor Szalai for his help with preliminary experiments.

This work was supported by grant RO1 DK51526 from the National Institutes of Health to G. Hajnóczky.