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Article |
Correspondence to Gyorgy Hajnóczky: gyorgy.hajnoczky{at}jefferson.edu
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Abstract |
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Introduction |
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Distribution of mitochondria to strategic sites is likely to be established by a cytoskeleton-based transportation system. Mitochondria have been visualized in association with the microfilaments (MFs; Drubin et al., 1993; Morris and Hollenbeck, 1995), microtubules (MTs; Ball and Singer, 1982), and intermediate filaments (Summerhayes et al., 1983; Stromer and Bendayan, 1990) in various cell types. For the binding of cytoskeletal elements, docking proteins have been identified on the mitochondria (e.g., dynactin for the microtubular motor protein, cytoplasmic dynein [Habermann et al., 2001; Varadi et al., 2004], or for both dynein and kinesin [Deacon et al., 2003]). Mitochondria-bound molecular motors provide a means for the organelles to move along the cytoskeletal fibers. Mice lacking Kif1B, the kinesin motor that binds to the mitochondria, are embryonic lethal and in their cells mitochondria are clustered around the nucleus (Tanaka et al., 1998). However, mitochondrial motility is not restricted to the delivery of organelles from the site of biogenesis to their destinations. In fact, mitochondria exist as highly dynamic structures in the cells (for review see Yaffe, 1999). Mitochondrial motility appears in the form of both long-distance travel and complex local movements, mostly wiggling. Movements may result in a change in the distribution of mitochondria in the cell and, in turn, rearrange the spatial pattern of ATP production and Ca2+ buffering. Mitochondrial movements may also increase the chance of dynamic interactions between discrete organelles and may aid the transport of molecules between the cytoplasm and mitochondria. However, movements may interfere with the rapid transport of molecules from an organelle to a neighboring one if the transport depends on activation of the acceptor transport site by a short-lasting concentration surge that is confined to the vicinity of the donor site. A bidirectional coupling has also been proposed between mitochondrial motility and morphology, whereby the dyneindynactin complex contributes to the mitochondrial targeting of Drp1 that promotes mitochondrial fission to make possible anterograde transport of the mitochondria by kinesin family motors (Varadi et al., 2004). Thus, mitochondrial motility and dynamic changes in motility may affect in many ways the signaling pathways and cell function.
Recently, inhibition of mitochondrial motility was described as an early and Ca2+-dependent event during the NMDA receptor-mediated excitotoxic injury in primary forebrain neurons (Rintoul et al., 2003). Furthermore, evidence has been presented that the direction of movement of mitochondria along the MTs is affected by the level of phosphatidyl inositol 4,5-bisphosphate (De Vos et al., 2003). Cleavage of phosphatidyl inositol 4,5-bisphosphate to produce inositol 1,4,5-trisphosphate (IP3) and IP3-induced mobilization of Ca2+ to establish [Ca2+]c oscillations are fundamental steps in the signaling cascade induced by a variety of hormones, neurotransmitters, and growth factors. These observations raise the possibility that mitochondrial motility may be controlled in a dynamic manner by the messenger molecules that mediate the effect of agonists on cell function.
The aims of the present work were to determine how physiological calcium signals affect mitochondrial motility and to study the mechanisms that may relay the effect of calcium to mitochondrial motor proteins. We have established a fluorescence imaging approach to monitor mitochondrial movement activity simultaneously with calcium spikes and oscillations. Using this approach, we show that mitochondrial movement is effectively stopped during both IP3 receptor and ryanodine receptor (RyR)mediated [Ca2+]c spikes and oscillations in H9c2 myoblasts. Inhibition of motility followed the spatial and temporal pattern of the [Ca2+]c signal. Although the decay of each [Ca2+]c spike was followed by recovery of mitochondrial motility, during high frequency [Ca2+]c oscillations sustained inhibition of mitochondrial motility occurred. Our results also indicate that depression of mitochondrial movement promotes the [Ca2+]c signal propagation into the mitochondria. Thus, diminished mitochondrial motility in the region of the [Ca2+]c rise would support recruitment of the mitochondria to enhance local Ca2+ buffering and ATP production. The increase in local mitochondrial Ca2+ uptake and in the energy supply of Ca2+ pumps facilitate the decay of the [Ca2+]c rise, serving as a feedback mechanism in calcium signaling.
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Results |
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As a first approach to evaluate the mitochondrial motility, two images obtained 10 s apart from each other were colored green and red, respectively, and were subsequently overlaid (Fig. 1 A, i). In the overlay, the yellow (green+red) pixels represent the mitochondria that maintained their position, whereas the green and red pixels indicate the sites of movement. One way to show only the sites of movement is to calculate the difference of the two images (F13.3s F23.2s; Fig. 1 A, ii, negative values shown in green, positive values in red, respectively). The amount and distribution of green and red pixels in the difference image corresponds with that in the overlay image (Fig. 1 A, i and ii). Green and red pixels are mostly side-by-side, indicating lateral movement of the organelles, whereas the single green or red pixels are likely to reflect movement into or out of the focus plane. Similar analysis was performed with two images recorded after addition of VP. In this case, very few green and red pixels were obtained, confirming a decrease in mitochondrial mobility (Fig. 1 A, iii and iv).
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A fundamental second messenger mobilized by VP is Ca2+ that controls many forms of motility. To test the possibility that the Ca2+ signal was important for the VP effect on mitochondrial motility, cells were also treated with ionomycin (Iono), a Ca2+ ionophore that allows both Ca2+ release from intracellular stores and Ca2+ entry from the bathing medium to elevate [Ca2+]c. Iono triggered a rapid and almost complete loss of motility (Fig. 1 A, vx). Thus, a rise in [Ca2+] is likely to account for the arrest of mitochondrial movement.
To determine the relation between the [Ca2+]c signal and mitochondrial motility, cells expressing mitoYFP were loaded with fura2/AM, a Ca2+-sensitive fluorescent probe, and [Ca2+]c was monitored simultaneously with YFP fluorescence. As shown in Fig. 1 B, the VP-induced decrease in mitochondrial motility (Fig. 1 B, top row) was closely coupled to the rising phase of the [Ca2+]c spike (Fig. 1 B, bottom row) and the recovery of mitochondrial motility lagged slightly behind the decay of the [Ca2+]c signal. During the [Ca2+]c signal, removal of extracellular Ca2+ by EGTA caused [Ca2+]c to return to the resting level and at the same time, mitochondrial motility to return to the original activity (Fig. 1 B). When Ca2+ was added back to the bathing medium, Ca2+ entry provided for a substantial [Ca2+]c increase. This [Ca2+]c signal was also closely followed by inhibition of mitochondrial movement. Again, the [Ca2+]c rise and the inhibition could be reversed by addition of EGTA (Fig. 1 B). To ascertain the role of Ca2+ in the effect of VP, VP was also added to Ca2+-depleted cells. In the absence of a [Ca2+]c rise, no VP-induced change in mitochondrial movement was recorded (Fig. 1 C). Together, these results show that the mitochondrial motility is dynamically controlled by the VP-induced, IP3-linked [Ca2+]c signal.
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 (50100 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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No role for mitochondrial Ca2+ uptake and mitochondrial membrane potential (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 m that also drives mitochondrial ATP synthesis. To determine whether the [Ca2+]m signal or
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
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
m appears to be dispensable for the Ca2+-mediated inhibition of mitochondrial movement.
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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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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-2related 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.
Arrest of the mitochondria supports rapid Ca2+ delivery to the mitochondria
Movement is important for the delivery of mitochondria from the site of biogenesis to the sites where energy is needed. Furthermore, the present data on the spatio-temporal control of the movement by the [Ca2+]c signal shows immobilization of mitochondria in the regions that display a [Ca2+]c rise (Fig. 4 C). Because mitochondria have the capacity to accumulate Ca2+, another role of the movement and Ca2+-induced inhibition of the movement may be to dynamically control the distribution of the mitochondrial Ca2+ buffer to the spatial pattern of the ER/SR Ca2+ release or Ca2+ entry. In many paradigms, Ca2+ signal propagation to the mitochondria depends on exposure of the mitochondrial Ca2+ uptake sites to the high [Ca2+] microdomains generated in the vicinity of the ER/SR Ca2+ release sites (Rizzuto et al., 1993; 1998; Szalai et al., 2000; Pacher et al., 2002; for comparison see Collins et al., 2001). Movement may affect the distance between ER/SR and mitochondria and may affect the alignment between the Ca2+ release and uptake sites if there is physical coupling between the two organelles.
We postulated that rapid inhibition of the mitochondrial movement may promote the mitochondrial Ca2+ uptake during the brief Ca2+ release events. We first sought for a means to prevent the inhibition of mitochondrial movement by the calcium signal and to study [Ca2+]m in mitochondria that are not arrested by the [Ca2+]c signal. However, no approach has been identified to uncouple the inhibition of mitochondrial movement from the [Ca2+]c rise. Therefore, we tried to use the [Ca2+]c-independent decrease in mitochondrial movement induced by nocodazole (Fig. 5 B). Because the arrest of mitochondrial movement appears after the onset of the [Ca2+]c signal, we speculated that inhibition of mitochondrial motility by nocodazole-pretreatment may enhance the initial phase of the [Ca2+]m signal. To test this idea, cells were transfected with two constructs encoding ratiometric pericam, a Ca2+-sensitive fluorescent protein targeted to the nucleus and mitochondria, respectively (Nagai et al., 2001). Before stimulation, the cells were pretreated with nocodazole (10 µM for 2030 min) or with solvent (DMSO) and were stimulated with VP, while ratiometric imaging of pericam fluorescence was performed. By measuring separately the extranuclear and nuclear areas, [Ca2+]m was determined simultaneously with nuclear matrix [Ca2+]. Nuclear matrix [Ca2+] was used as a surrogate of [Ca2+]c that was not feasible to monitor simultaneously with [Ca2+]m. In separate experiments, no difference was noticed between the kinetics of the [Ca2+] signals recorded in VP-stimulated cells by cytosolic and nuclear pericam, respectively (n = 4; unpublished data).
The initial nuclear [Ca2+] rise was determined for both control and nocodazole-pretreated cells and the simultaneously measured [Ca2+]m rise response was also evaluated for the same cells (Fig. 5 B, bottom). The nuclear [Ca2+] rise was smaller in the nocodazole-pretreated cells than in the control cells. In contrast, the [Ca2+]m elevation was >60% greater in the nocodazole-pretreated cells than in the control cells (P < 0.01; n = 16). Thus, at the beginning of Ca2+ mobilization, the efficacy of the [Ca2+]c signal delivery to the mitochondria was increased in nocodazole-pretreated cells. Although it remains to find a more specific inhibitor of the mitochondrial motility, our result seems to support the idea that the arrest of mitochondrial motility during the [Ca2+]c signal promotes the [Ca2+]c signal propagation to the mitochondria. During repetitive stimulation or during [Ca2+]c oscillations (Fig. 5), the movement inhibition evoked by the first [Ca2+]c spike may last until the second [Ca2+]c spike rises and may facilitate the effect of the second [Ca2+]c spike on the mitochondria. Thus, inhibition of motility may provide a mechanism underlying the enhancement of mitochondrial calcium signaling during [Ca2+]c oscillations, a phenomenon that has been documented in several paradigms (for review see Csordás and Hajnóczky, 2003). The effective propagation to the mitochondria enables frequency-modulated [Ca2+]c oscillations to control the Ca2+-sensitive enzymes of ATP production over the full range of potential activities (Hajnóczky et al., 1995; Robb-Gaspers et al., 1998).
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Discussion |
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The scheme in Fig. 6 and the animation in Video 3 (available at http://www.jcb.org/cgi/content/full/jcb.200406038/DC1) show the mechanism of the Ca2+-dependent control of mitochondrial movement determined in the present work. At resting [Ca2+]c (Fig. 6, left), mitochondria display maximal movement activity and the majority of the movements occur along the MTs (Fig. 6 left, arrows). Movements of the mitochondria toward the plus end are promoted by kinesin motors, whereas movements to the minus end are facilitated by dynein motor proteins (Tanaka et al., 1998; Habermann et al., 2001; Deacon et al., 2003; Varadi et al., 2004). When a [Ca2+]c rise occurs due to either Ca2+ mobilization or Ca2+ entry, the mitochondrial movement decreases. Even a modest increase in [Ca2+]c is sufficient to attenuate mitochondrial motility and the elevation of global [Ca2+]c to 1 µM, a level that is commonly attained during [Ca2+]c spikes and oscillations results in almost complete loss of mitochondrial movement (Video 3, right panel). Ca2+ does not seem to activate Ca2+/CaM-dependent kinases or the Ca2+-dependent protein phosphatase to establish control over mitochondrial motility because several inhibitors of these enzymes failed to interfere with the movement inhibition by Ca2+. Furthermore, Ca2+ does not seem to target directly the microtubular motors because the molecular structure of mammalian cytoplasmic dynein and kinesin does not indicate the presence of a site for binding of Ca2+ or CaM (Vale, 2003). Thus, we propose that a distinct Ca2+ sensor molecule is required to translate the Ca2+ signal for the microtubular motor proteins. Binding of Ca2+ to the Ca2+ sensor would induce the MT-bound motors to lock in a stationary position or to detach from the MTs (Video 3, red symbol). One potential candidate for the Ca2+ sensor is myosin Va, a motor protein that binds CaM and is controlled by Ca2+ (for reviews see Reck-Peterson et al., 2000; Vale, 2003). Myosin Va displays a Ca2+-dependent interaction with actin-filaments (Tauhata et al., 2001; Krementsov et al., 2004) and MTs (Cao et al., 2004). Interaction of the head domain of myosin Va with actin provides a motor for movement along the MFs but the tail domain-based interaction with MTs does not present by itself a microtubular motor. However, recent papers have raised the possibility that myosin V can interact directly with dynein and kinesin and through this interaction may affect motor function at the MTs (Benashski et al., 1997; Huang et al., 1999; Stafford et al., 2000; Lalli et al., 2003). Immunocytochemistry studies indicate that myosin Va is present on the mitochondria in H9c2 cells, and myosin Va is also retained on isolated mitochondria (unpublished data). However, to clarify the role of myosin V in the Ca2+-dependent control of mitochondrial motility and to explore alternative mechanisms, further studies are needed.
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Recruitment of mitochondria to the sites of Ca2+ release by Ca2+-induced immobilization may form the basis for a novel homeostatic mechanism in calcium signaling. Targeting of mitochondria to the sites of Ca2+ release results in an increase in the local Ca2+ buffering capacity. Also, mitochondrial Ca2+ uptake serves as a means for stimulation of mitochondrial ATP production that provides a localized energy source for Ca2+ reuptake by the ER/SR (Landolfi et al., 1998). The increase in local Ca2+ buffering and in ATP production represents a feedback mechanism that contributes to the control of the [Ca2+]c rise and strengthening of the Ca2+ scavenger mechanisms may also be important to avoid Ca2+-dependent cell injury. Once Ca2+ release stops, motility is recovered and the mitochondria may be recruited by Ca2+ release in other regions of the cell. Thus, the Ca2+-dependent control of mitochondrial motility offers a means to adjust the subcellular spatial arrangements of the Ca2+ buffering and energy production as needed.
In conclusion, mitochondrial movement along the MTs ensures that mitochondria are available throughout the cell, whereas regulation by Ca2+ targets mitochondria to the sites where mitochondrial Ca2+ uptake and ATP production may be required to ensure the handling of Ca2+. Dynamic control of mitochondrial motility and distribution presents a powerful, novel mechanism to optimize the use of the mitochondrial pool in cellular Ca2+ transport and signaling.
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Materials and methods |
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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 2436 h before the imaging experiments.
Live cell imaging
Fluorescence imaging was performed using an inverted microscope (model IX70; Olympus; 40x, 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 (40x, UApo340, NA 1.35). Mitotracker green, GFP, and YFP were excited at 488 nm, and DsRed at 568 nm. Images were obtained every 3.13.3 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 2530 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 (Kd = 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 x 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.
Online supplemental material
Video 1 shows the inhibition of mitochondrial motility in an H9c2 cell stimulated with VP; Video 2 demonstrates the movement of mitochondria along MTs; and Video 3 is an animated representation of the proposed mechanism for calcium control of mitochondrial motility. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200406038/DC1.
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Acknowledgments |
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This work was supported by grant RO1 DK51526 from the National Institutes of Health to G. Hajnóczky.
Submitted: 7 June 2004
Accepted: 6 October 2004
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References |
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---|
Ball, E.H., and S.J. Singer. 1982. Mitochondria are associated with microtubules and not with intermediate filaments in cultured fibroblasts. Proc. Natl. Acad. Sci. USA. 79:123126.[Abstract]
Bhat, M.B., J. Zhao, W. Zang, C.W. Balke, H. Takeshima, W.G. Wier, and J. Ma. 1997. Caffeine-induced release of intracellular Ca2+ from Chinese hamster ovary cells expressing skeletal muscle ryanodine receptor. Effects on full-length and carboxyl-terminal portion of Ca2+ release channels. J. Gen. Physiol. 110:749762.
Benashski, S.E., A. Harrison, R.S. Patel-King, and S.M. King. 1997. Dimerization of the highly conserved light chain shared by dynein and myosin V. J. Biol. Chem. 272:2092920935.
Bernardi, P., V. Petronilli, F. Di Lisa, and M. Forte. 2001. A mitochondrial perspective on cell death. Trends Biochem. Sci. 26:112117.[CrossRef][Medline]
Cao, T.T., W. Chang, S.E. Masters, and M.S. Mooseker. 2004. Myosin-Va binds to and mechanochemically couples microtubules to actin filaments. Mol. Biol. Cell. 15:151161.
Collins, T.J., P. Lipp, M.J. Berridge, and M.D. Bootman. 2001. Mitochondrial Ca2+ uptake depends on the spatial and temporal profile of cytosolic Ca2+ signals. J. Biol. Chem. 276:2641126420.
Couchman, J.R., and D.A. Rees. 1982. Organelle-cytoskeleton relationships in fibroblasts: mitochondria, Golgi apparatus, and endoplasmic reticulum in phases of movement and growth. Eur. J. Cell Biol. 27:4754.[Medline]
Csordás, G., and G. Hajnóczky. 2003. Plasticity of mitochondrial calcium signaling. J. Biol. Chem. 278:4227342282.
Csordás, G., A.P. Thomas, and G. Hajnóczky. 1999. Quasi-synaptic calcium signal transmission between endoplasmic reticulum and mitochondria. EMBO J. 18:96108.
Deacon, S.W., A.S. Serpinskaya, P.S. Vaughan, M. Lopez Fanarraga, I. Vernos, K.T. Vaughan, and V.I. Gelfand. 2003. Dynactin is required for bidirectional organelle transport. J. Cell Biol. 160:297301.
Demaurex, N., and C. Distelhorst. 2003. Cell biology. Apoptosisthe calcium connection. Science. 300:6567.
De Vos, K.J., J. Sable, K.E. Miller, and M.P. Sheetz. 2003. Expression of phosphatidylinositol (4,5) bisphosphate-specific pleckstrin homology domains alters direction but not the level of axonal transport of mitochondria. Mol. Biol. Cell. 14:36363649.
Drubin, D.G., H.D. Jones, and K.F. Wertman. 1993. Actin structure and function: roles in mitochondrial organization and morphogenesis in budding yeast and identification of the phalloidin-binding site. Mol. Biol. Cell. 4:12771294.[Abstract]
Duchen, M.R. 2000. Mitochondria and calcium: from cell signalling to cell death. J. Physiol. 529:5768.
Filippin, L., P.J. Magalhaes, G. Di Benedetto, M. Colella, and T. Pozzan. 2003. Stable interactions between mitochondria and endoplasmic reticulum allow rapid accumulation of calcium in a subpopulation of mitochondria. J. Biol. Chem. 278:3922439234.
Habermann, A., T.A. Schroer, G. Griffiths, and J.K. Burkhardt. 2001. Immunolocalization of cytoplasmic dynein and dynactin subunits in cultured macrophages: enrichment on early endocytic organelles. J. Cell Sci. 114:229240.
Hajnóczky, G., L.D. Robb-Gaspers, M. Seitz, and A.P. Thomas. 1995. Decoding of cytosolic calcium oscillations in the mitochondria. Cell. 82:415424.[Medline]
Hajnóczky, G., G. Csordás, and M. Yi. 2002. Old players in a new role: mitochondria-associated membranes, VDAC, and ryanodine receptors as contributors to calcium signal propagation from endoplasmic reticulum to the mitochondria. Cell Calcium. 32:363377.[CrossRef][Medline]
Hoth, M., C.M. Fanger, and R.S. Lewis. 1997. Mitochondrial regulation of store-operated calcium signaling in T lymphocytes. J. Cell Biol. 137:633648.
Huang, J.D., S.T. Brady, B.W. Richards, D. Stenolen, J.H. Resau, N.G. Copeland, and N.A. Jenkins. 1999. Direct interaction of microtubule- and actin-based transport motors. Nature. 397:267270.[CrossRef][Medline]
Jacobson, J., and M.R. Duchen. 2001. What nourishes me, destroys me: towards a new mitochondrial biology. Cell Death Differ. 8:963966.[CrossRef][Medline]
Khodjakov, A., E.M. Lizunova, A.A. Minin, M.P. Koonce, and F.K. Gyoeva. 1998. A specific light chain of kinesin associates with mitochondria in cultured cells. Mol. Biol. Cell. 9:333343.
Krementsov, D.N., E.B. Krementsova, and K.M. Trybus. 2004. Myosin V: regulation by calcium, calmodulin, and the tail domain. J. Cell Biol. 164:877886.
Kroemer, G., and J.C. Reed. 2000. Mitochondrial control of cell death. Nat. Med. 6:513519.[CrossRef][Medline]
Lalli, G., S. Gschmeissner, and G. Schiavo. 2003. Myosin Va and microtubule-based motors are required for fast axonal retrograde transport of tetanus toxin in motor neurons. J. Cell Sci. 116:46394650.
Landolfi, B., S. Curci, L. Debellis, T. Pozzan, and A.M. Hofer. 1998. Ca2+ homeostasis in the agonist-sensitive internal store: functional interactions between mitochondria and the ER measured in situ in intact cells. J. Cell Biol. 142:12351243.
Lawrie, A.M., R. Rizzuto, T. Pozzan, and A.W. Simpson. 1996. A role for calcium influx in the regulation of mitochondrial calcium in endothelial cells. J. Biol. Chem. 271:1075310759.
Marchant, J.S., V. Ramos, and I. Parker. 2002. Structural and functional relationships between Ca2+ puffs and mitochondria in Xenopus oocytes. Am. J. Physiol. Cell Physiol. 282:C1374C1386.
Martinou, J.C., and D.R. Green. 2001. Breaking the mitochondrial barrier. Nat. Rev. Mol. Cell Biol. 2:6367.[CrossRef][Medline]
Montero, M., M.T. Alonso, E. Carnicero, I. Cuchillo-Ibanez, A. Albillos, A.G. Garcia, J. Garcia-Sancho, and J. Alvarez. 2000. Chromaffin-cell stimulation triggers fast millimolar mitochondrial Ca2+ transients that modulate secretion. Nat. Cell Biol. 2:5761.[CrossRef][Medline]
Morris, R.L., and P.J. Hollenbeck. 1995. Axonal transport of mitochondria along microtubules and F-actin in living vertebrate neurons. J. Cell Biol. 131:13151326.[Abstract]
Nagai, T., A. Sawano, E.S. Park, and A. Miyawaki. 2001. Circularly permuted green fluorescent proteins engineered to sense Ca2+. Proc. Natl. Acad. Sci. USA. 98:31973202.
Nangaku, M., R. Sato-Yoshitake, Y. Okada, Y. Noda, R. Takemura, H. Yamazaki, and N. Hirokawa. 1994. KIF1B, a novel microtubule plus end-directed monomeric motor protein for transport of mitochondria. Cell. 79:12091220.[Medline]
Pacher, P., and G. Hajnóczky. 2001. Propagation of the apoptotic signal by mitochondrial waves. EMBO J. 20:41074121.
Pacher, P., A.P. Thomas, and G. Hajnóczky. 2002. Ca2+ marks: miniature calcium signals in single mitochondria driven by ryanodine receptors. Proc. Natl. Acad. Sci. USA. 99:23802385.
Parekh, A.B. 2003. Store-operated Ca2+ entry: dynamic interplay between endoplasmic reticulum, mitochondria and plasma membrane. J. Physiol. 547:333348.
Park, M.K., M.C. Ashby, G. Erdemli, O.H. Petersen, and A.V. Tepikin. 2001. Perinuclear, perigranular and sub-plasmalemmal mitochondria have distinct functions in the regulation of cellular calcium transport. EMBO J. 20:18631874.
Petersen, O.H. 2002. Calcium signal compartmentalization. Biol. Res. 35:177182.[Medline]
Reck-Peterson, S.L., D.W. Provance Jr., M.S., Mooseker, and J.A. Mercer. 2000. Class V myosins. Biochim. Biophys. Acta. 1496:3651.[Medline]
Rintoul, G.L., A.J. Filiano, J.B. Brocard, G.J. Kress, and I.J. Reynolds. 2003. Glutamate decreases mitochondrial size and movement in primary forebrain neurons. J. Neurosci. 23:78817888.
Rizzuto, R., M. Brini, M. Murgia, and T. Pozzan. 1993. Microdomains with high Ca2+ close to IP3-sensitive channels that are sensed by neighboring mitochondria. Science. 262:744747.[Medline]
Rizzuto, R., P. Pinton, W. Carrington, F.S. Fay, K.E. Fogarty, L.M. Lifshitz, R.A. Tuft, and T. Pozzan. 1998. Close contacts with the endoplasmic reticulum as determinants of mitochondrial Ca2+ responses. Science. 280:17631766.
Rizzuto, R., P. Bernardi, and T. Pozzan. 2000. Mitochondria as all-round players of the calcium game. J. Physiol. 529:3747.
Rizzuto, R., M.R. Duchen, and T. Pozzan. 2004. Flirting in little space: the ER/mitochondria Ca2+ liaison. Sci. STKE. 2004:re1.
Robb-Gaspers, L.D., P. Burnett, G.A. Rutter, R.M. Denton, R. Rizzuto, and A.P. Thomas. 1998. Integrating cytosolic calcium signals into mitochondrial metabolic responses. EMBO J. 17:49875000.
Scorrano, L., and S.J. Korsmeyer. 2003. Mechanisms of cytochrome c release by proapoptotic BCL-2 family members. Biochem. Biophys. Res. Commun. 304:437444.[CrossRef][Medline]
Stafford, P., J. Brown, and G.M. Langford. 2000. Interaction of actin- and microtubule-based motors in squid axoplasm probed with antibodies to myosin V and kinesin. Biol. Bull. 199:203205.[Medline]
Stromer, M.H., and M. Bendayan. 1990. Immunocytochemical identification of cytoskeletal linkages to smooth muscle cell nuclei and mitochondria. Cell Motil. Cytoskeleton. 17:1118.[Medline]
Szalai, G., G. Csordás, B. Hantash, A.P. Thomas, and G. Hajnóczky. 2000. Calcium signal transmission between ryanodine receptors and mitochondria. J. Biol. Chem. 275:1530515313.
Summerhayes, I.C., D. Wong, and L.B. Chen. 1983. Effect of microtubules and intermediate filaments on mitochondrial distribution. J. Cell Sci. 61:87105.[Abstract]
Tanaka, Y., Y. Kanai, Y. Okada, S. Nonaka, S. Takeda, A. Harada, and N. Hirokawa. 1998. Targeted disruption of mouse conventional kinesin heavy chain, kif5B, results in abnormal perinuclear clustering of mitochondria. Cell. 93:11471158.[Medline]
Tauhata, S.B., D.V. dos Santos, E.W. Taylor, M.S. Mooseker, and R.E. Larson. 2001. High affinity binding of brain myosin-Va to F-actin induced by calcium in the presence of ATP. J. Biol. Chem. 276:3981239818.
Vale, R.D. 2003. The molecular motor toolbox for intracellular transport. Cell. 112:467480.[Medline]
Varadi, A., L.I. Johnson-Cadwell, V. Cirulli, Y. Yoon, V.J. Allan, and G.A. Rutter. 2004. Cytoplasmic dynein regulates the subcellular distribution of mitochondria by controlling the recruitment of the fission factor dynamin-related protein-1. J. Cell Sci. 117:43894400.
Wang, H.J., G. Guay, L. Pogan, R. Sauve, and I.R. Nabi. 2000. Calcium regulates the association between mitochondria and a smooth subdomain of the endoplasmic reticulum. J. Cell Biol. 150:14891498.
Yaffe, M.P. 1999. Dynamic mitochondria. Nat. Cell Biol. 1:E149E150.[CrossRef][Medline]
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