1Department of Physiology and 2Biomedical Imaging Group, University of Massachusetts Medical School, Worcester, Massachusetts 01655
Submitted 2 September 2003 ; accepted in final form 31 December 2003
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ABSTRACT |
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mitochondria; mitochondrial membrane potential; intracellular calcium; permeability transition pore; sarcoplasmic reticulum
The mitochondrial membrane potential (m) provides the electrical gradient necessary for mitochondrial Ca2+ uptake via the Ca2+ uniporter, a low-affinity, high-capacity electrogenic Ca2+ transporter. Due to the low affinity of the Ca2+ uniporter, it was difficult to understand how mitochondria could be sensitive to the small amplitude of cytosolic Ca2+ signals in many cells (5). To account for Ca2+ buffering by mitochondria at low global [Ca2+]c after release from endoplasmic reticulum (ER) via inositol 1,4,5-trisphosphate (IP3) receptors (IP3Rs), Rizzuto et al. proposed an explanation based on microdomains (39, 40). ER surfaces containing IP3Rs were hypothesized to be in close contact with mitochondria. Thus, after Ca2+ release from IP3Rs, the [Ca2+]c in the vicinity of mitochondrial Ca2+ uptake sites is much higher than the mean global submicromolar [Ca2+]c. This hypothesis has been generalized to include Ca2+ release via ryanodine receptors (RyRs) on sarcoplasmic reticulum (SR) although Rizzuto et al. did not deal with these receptors (5). Transient microdomains of high Ca2+ have been imaged and called "sparks" in cardiomyocytes (9) and "puffs" in oocytes (46). Sparks and puffs are caused by spontaneous, transient release of Ca2+ via RyRs or IP3Rs, respectively.
m is not stable but subject to spontaneous fluctuations in many cell types. These transient depolarizations of
m were first observed in neuroblastoma cells loaded with tetramethylrhodamine ethyl ester perchlorate (TMRE), a potential sensitive fluorescent indicator (32). TMRE accumulates rapidly and reversibly in the mitochondria of living cells, a higher concentration of dye localizing to the matrix of energized (i.e. polarized) mitochondria than to the matrix of depolarized mitochondria. Changes in
m cause redistribution of TMRE across the inner mitochondrial membrane; this is detected as variations in the measured fluorescence intensity (FI) of the dye. In cardiomyocytes, the term "flickers" was coined to describe these changes in
m (17).
It has been suggested that flickers result from mitochondrial uptake of Ca2+. Duchen and coworkers (17) demonstrated that flickers in rat cardiomyocytes are almost abolished by inhibition of Ca2+ release from SR, by the intracellular Ca2+ chelator BAPTA-AM, and by inhibition of the mitochondrial Ca2+ uniporter. They concluded that flickers are transient depolarizations caused by mitochondrial Ca2+ uptake after Ca2+ sparks in cardiomyocytes (17). There are several mechanisms by which mitochondrial uptake of Ca2+ from a microdomain may result in loss of m. As described above, the Ca2+ uniporter is an electrogenic transporter, and thus Ca2+ uptake into mitochondria may be expected to cause depolarization of
m (31). Moreover, should the mitochondrial inner membrane have a high electrical resistance, then very little Ca2+ uptake is required to cause significant mitochondrial depolarization (17). Thus Ca2+ uptake alone may be sufficient to cause flickers. Alternatively, an increase in [Ca2+]m may activate Ca2+-sensitive ion channels in the mitochondrial membrane. One such transporter is the mitochondrial permeability transition pore (PTP), a nonselective ion channel in the inner mitochondrial membrane. Flickers have frequently been attributed to PTP stimulation after phototoxicity- or oxidative stress-induced increases in reactive oxygen species (ROS) within mitochondria (12, 13, 18, 25, 28, 49). PTP activation may also be achieved in the absence of an inducer (such as ROS) by an increase in [Ca2+]m (21). In some studies where flickers were attributed to PTP activation, the transient mitochondrial depolarizations were attenuated by inhibition of Ca2+ signaling (18, 28).
Our purpose here was to examine the relationship between focal Ca2+ release from SR and mitochondrial flickers. We used a preparation of freshly dissociated smooth muscle cells where both Ca2+ sparks and mitochondrial flickers have been examined in some detail using new quantitative methods that we have devised (36, 47). In a previous study (36), we have shown that in smooth muscle cells, nearly all mitochondria flicker and do so independently of one another and not as part of a network. The quantitative method used in that study and employed again here allowed us to demonstrate that mean flicker amplitude is <20 mV although magnitudes range from <10 mV to >100 mV. Moreover, the data indicated that an ion channel or electrogenic exchanger in the mitochondrial inner membrane caused these transient depolarizations in m (36). In the current study we wished to investigate the mechanism by which flickers occur. In particular, we sought to determine whether flickers can be elicited by uptake of Ca2+ released focally by RyRs during a spark and, if so, are they caused by dissipation of
m simply due to Ca2+ influx. PTP activation was thought to be unlikely under the experimental conditions employed (see RESULTS). Nonetheless, we also assessed the possibility that either Ca2+-dependent or Ca2+-independent PTP activation caused flickers.
High-speed three-dimensional imaging of TMRE-loaded cells revealed spontaneous oscillations in the fluorescence from individual mitochondria that signal transient mitochondrial depolarizations. Flickers were apparent both in cells that exhibited spontaneous transient outward currents (STOCs), which are elicited by sparks, and in cells without STOCs. Mitochondrial flickers were also observed under conditions that are known to inhibit Ca2+ sparks. Flickers were not affected by either ryanodine (100 µM) or xestospongin C (0.5 µM), inhibitors of Ca2+ release from ryanodine- and IP3-sensitive intracellular Ca2+ stores, respectively. Store depletion induced by caffeine and thapsigargin failed to inhibit flickers; in fact, both flicker size and frequency increased after emptying of the stores. Neither cyclosporin A (a PTP inhibitor) nor trolox (a ROS scavenger) abolished flickers. This study provides evidence that transient changes in m occur in the absence of Ca2+ release from intracellular stores or PTP activation in smooth muscle cells. Some of these findings have been reported previously in abstract form (34, 35).
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MATERIALS AND METHODS |
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TMRE loading protocol. Freshly isolated myocytes were incubated with TMRE (25 nM) for 10 min at room temperature. Cells were transferred to a tissue bath, where the final concentration of TMRE was 2.5 nM. Cells were allowed to equilibrate for 10 min before whole cell patch-clamp recording was established.
Electrophysiology. All cells were patch clamped in the whole cell configuration at room temperature using thin-walled borosilicate patch electrodes (5 M
) (World Precision Instruments). The membrane potential was voltage clamped to 0 mV, and cells were allowed to equilibrate for 510 min before data acquisition. Thus cytosolic [TMRE] ([TMRE]c) was constant and equal to that in the bath solution, i.e., 2.5 nM.
Imaging. Three-dimensional images of TMRE-labeled mitochondria were obtained using a high-speed, wide-field digital imaging microscope and CCD camera (Lincoln Laboratory, MIT), which have been described in detail elsewhere (48). TMRE was excited at 514 nm using an argon-krypton laser (widefield illumination flux 4 x 1019 photons·cm-2·s-1). A shutter was used to limit the total light exposure of the cells. TMRE emission was detected with a 550-nm long-pass filter. The Nikon 60x, NA 1.4, oil immersion objective was rapidly focused using a piezoelectric translator. A 10-plane three-dimensional image stack was acquired in 140 ms using 0.5-µm Z spacing with 5-ms exposures at 10-ms intervals to allow movement of the objective. Twenty image stacks were acquired at 5-s intervals to generate image sequences of 95-s duration. The pixel size was 333 x 333 nm, and the total area imaged was 42.6 x 42.6 µm. Three-dimensional imaging made it possible to resolve the fluorescence of individual mitochondria reliably without contamination of the signal by fluorescence from mitochondria in other planes. It also ensured that movement into and out of the plane of focus (due to movement of the entire cell or contraction in a portion of the cell) was not mistaken for changes in mitochondrial FI.
Image processing and data analysis. Image processing and data analysis were performed using custom software on a SiliconGraphics work station. Due to the wide-field illumination, blurred fluorescence from out of focus objects contributes to the sum of the fluorescence at any given point within the acquired images. The data were processed using an iterative constrained deconvolution algorithm to restore out of focus light to its point of origin and enable accurate data analysis (6, 7). Mitochondria were easily identified in the restored three-dimensional images (see RESULTS). Voxels were selected over the entire length of the mitochondrion, the number of voxels used depending on the length of the mitochondrion. The central z-plane of each mitochondrion was identified in each three-dimensional image stack; each voxel was centered in the midpoint of mitochondrial depth and width. The average fluorescence of these voxels was designated as the FI. Mitochondria that drifted out of the field of view due to small movements of the cell were not included in the analysis, avoiding false identification of changes in m. The percent noise in the signal was calculated to be 5.9%. A change in FI of >20% (3.4-fold the mean %noise) in a 5- to 10-s period was considered a change in mitochondrial potential and designated a "flicker."
As we have previously reported, significant levels of TMRE bind to the inner mitochondrial membrane; thus absolute m cannot be calculated using the Nernst equation (36). However, because [TMRE]c is constant, changes in
m can be calculated as follows
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The percentage of mitochondria that flickered in each cell was calculated as follows: 100 x number of flickering mitochondria/total number of mitochondria in the field. Flicker frequency in flickering mitochondria was calculated as follows: total number of flickers observed during an image sequence/number of flickering mitochondria.
Image preparation. Custom software was used to prepare the images shown throughout the paper. Figure legends indicate whether a single plane from the three-dimensional image stack or a composite image (formed by projecting the 10 planes of a three-dimensional image stack onto a single plane) is displayed. Polarized mitochondria appear brighter than depolarized mitochondria.
Solutions and reagents. Experiments were carried out using the following solutions (in mM): pipette, 137 KCl, 3 MgCl2, 20 HEPES, 3 Na2ATP, pH 7.2 with KOH; bath, 130 NaCl, 3 KCl, 1.8 CaCl2, 1 MgCl2, 10 HEPES, pH 7.4 with NaOH. TMRE was purchased from Molecular Probes; ryanodine, thapsigargin, and xestospongin C were from Calbiochem. All other reagents were from Sigma. Stock solutions of TMRE (10 mM) and xestospongin C (500 µM) were made up in DMSO. Thapsigargin stock (1 mM) was prepared in ethanol. Ryanodine stock (50 mM) was made up in ultrapure water. Stocks were stored at -20°C. Caffeine (20 mM) was made up in bath solution and applied directly to the cells using a picospritzer II (General Valve).
Statistical analysis. The data result from an examination of 243 individual mitochondria and 1,862 flickers. Data are presented as means ± SE. Statistical significance was assessed using Graphpad Instat (3.05) software. Kruskal-Wallis nonparametric ANOVA was utilized to assess the data, which did not follow a normal distribution. P 0.05 was considered statistically significant.
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RESULTS |
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Flickers, transient fluctuations of m, can be seen in Fig. 1B. The arrows indicate two mitochondria that flicker during the 95-s image sequence. The traces in Fig. 1C show differences in the magnitude and duration of the mitochondrial flickers. The graph displays changes in
m during flickers in these mitochondria rather than absolute
m. In theory
m can be calculated easily using the Nernst equation, but this approach is complicated by two important factors. First, in previous studies high concentrations of TMRE have often been used so that this probe becomes self-quenching within the mitochondrial matrix, disrupting the simple exponential relationship between mitochondrial [TMRE] ([TMRE]m) and
m (4, 10, 17, 28). We overcame this by employing a much lower cytosolic concentration than used heretofore (2.5 nM), which was made possible with a high-sensitivity, low-noise, digital imaging system. Such a low concentration also minimizes phototoxicity (see below). Second, there appears to be partitioning of TMRE into the inner mitochondrial membrane (36, 41), but as we have demonstrated, such partitioning is not an impediment to the calculation, in millivolts, of changes in
m, i.e., flickers (36). Results are presented on a log scale as the change in
m is proportional to the log of the ratio of the free [TMRE]m before and during depolarization. As previously observed, mitochondria flicker independently of their neighbors, indicating that mitochondria operate individually rather than as part of a network (36).
Two successive image sequences were obtained from each cell before experimental manipulations and used as controls. There was no significant difference in flicker amplitude, percentage of flickering mitochondria, or frequency of flickers within flickering mitochondria between these two consecutive image sequences (Table 1).
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Mitochondrial flickers are independent of ryanodine-sensitive Ca2+ release from SR. The cells used in this study generate Ca2+ sparks via release of Ca2+ from SR through RyRs. The sparks in turn activate large-conductance Ca2+-activated K+ channels, causing STOCs that occur at a frequency of 2.5/s (48). Ryanodine inhibits Ca2+ efflux via RyRs at concentrations in the micromolar range (8). Cells were treated with 100 µM ryanodine to inhibit Ca2+ release from SR. At the start of the experiment, STOCS were apparent in two of the three cells studied, but 15 min after ryanodine addition STOCs were almost completely abolished, indicating that sparks had been inhibited. Image sequences were taken at 0.5, 5, and 15 min after addition of ryanodine to the bath. Flickers are evident 15 min after the application of ryanodine (Fig. 2A). Ryanodine was without effect on the percentage of mitochondria that were flickering, the number of flickers per flickering mitochondrion, and the flicker amplitude (Fig. 2B).
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Mitochondrial flickers are not affected by xestospongin C, a blocker of IP3Rs. IP3R-mediated SR Ca2+ release has been linked to elevation of global cytosolic Ca2+ in several smooth muscle cell types. Mitochondria take up Ca2+ after release through IP3Rs in rat pulmonary artery myocytes (15). In colonic myocytes, localized Ca2+ transients and STOCs were inhibited by xestospongin C [a membrane-permeable blocker of IP3-induced Ca2+ release (20)] but not by ryanodine (1). To investigate IP3R involvement, 0.5 µM xestospongin C was added to the bath, and cells were imaged at 10 s, 5 min, and 10 min after its application. Flickers can be seen 10 min after the addition of xestospongin C to the bath (Fig. 3A). A small inward shift in the holding current was seen after application of xestospongin C. Xestospongin C did not inhibit the percentage of mitochondria that flickered, the flicker frequency, or amplitude (Fig. 3B).
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Mitochondrial flickers are increased by SR store depletion. Chelation of cytosolic Ca2+ should block Ca2+-mediated cellular events regardless of whether the Ca2+ has an intra- or extracellular origin. Flickers were still observed in the presence of the membrane-permeable Ca2+ chelator BAPTA-AM (10 µM). Flickers were not expected to be dependent on Ca2+ influx. Had they been, they would have been observed primarily in those mitochondria in close proximity to the plasma membrane, whereas they were detected in almost all mitochondria. Nevertheless, lack of inhibition by BAPTA indicates that uptake of Ca2+ after influx across the cell membrane does not cause flickers. However, it has been suggested that mitochondrial Ca2+ uptake after intracellular Ca2+ release in microdomains is resistant to inhibition by BAPTA (28). Thus another approach was required to investigate release of store Ca2+ as the cause of the flickers. Cells were incubated with 1 µM thapsigargin and 7 min later exposed to 20 mM caffeine for 5 s. This procedure has been shown to deplete SR stores in the cells used here (48). STOCs were abolished after exposure to caffeine, indicating that sparks were inhibited. In contrast, flickers were observed in all three cells throughout the experiment (Fig. 4). Store depletion slightly but significantly increased the percentage of mitochondria that flickered (P = 0.01), the frequency of flickers (P = 0.03), and the flicker amplitude (P = 0.009) (Fig. 4B).
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Mitochondrial flickers are independent of phototoxicity and PTP activity. Flickers were not inhibited by any measures taken to prevent Ca2+ release from intracellular stores. Thus neither Ca2+-dependent dissipation of m nor Ca2+-dependent activation of the PTP causes flickers. However, in many previous studies, flickers were attributed to repetitive transient openings of the PTP, caused by phototoxicity-induced accumulation of ROS in the mitochondria (10, 12, 13, 25, 26, 49). In each of those studies, flickers culminated in complete dissipation of
m in the entire mitochondrial population. In cardiac myocytes, PTP inhibitors and ROS scavengers prevented loss of
m, but neither Ca2+ chelation nor inhibition of mitochondrial Ca2+ uptake had any effect (49). Thus Ca2+-independent PTP activation, and consequently mitochondrial depolarization, can occur as a result of phototoxicity. Phototoxic effects are dependent on the concentration of TMRE and the strength and duration of light exposure (10, 12, 13).
The high-sensitivity CCD camera used in the present study made it possible to limit [TMRE]c and photon exposure to low levels. Thus the likelihood of ROS generation and PTP activation was minimized. Moreover, the pipette solution included Mg2+ and ATP, known to inhibit or reverse PTP activation (2, 22, 24, 33).
The data presented in Table 1 indicate that phototoxic activation of PTP was not the cause of the flickers, as there was no significant difference in amplitude or frequency of flickering with additional light exposure over the two consecutive control sequences. Indeed, with a single exception (Fig. 4), the size and frequency of flickers did not change over the entire course of the experimental manipulations that followed these initial control sequences. Furthermore, the degree of progressive mitochondrial depolarization over the course of the experiments detailed above (Figs. 2, 3, 4) was evaluated. The maximum FI of each individual mitochondrion during control 1 was compared with the maximum FI of each mitochondrion in the final image sequence taken. The change in mitochondrial FI represents a 15.2 ± 0.9 mV decrease in m. This does not correspond to the complete loss of
m ultimately observed in all other studies after PTP activation. These findings are contrary to those expected if flickers were caused by phototoxic activation of the PTP.
Qualitative examination of the images also indicates that phototoxicity does not account for the flickers. Mitochondria of similar resting FI (i.e. similar [TMRE]m) imaged simultaneously do not depolarize in concert (Fig. 5A). Strikingly, in several instances mitochondria depolarized between imaging sequences in the absence of illumination and repolarized subsequently on renewed exposure to light, as imaging resumed (Fig. 5B).
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To further assess PTP activity, we applied cyclosporin A, a PTP inhibitor, to the cells. In astrocytes it has been shown that the rate of progressive global depolarization is significantly delayed in the presence of cyclosporin A (28). Caffeine (20 mM) was applied to the cells for 5 s to elicit a rise in global [Ca2+]c, and FI was monitored for 10 min after the caffeine pulse. Caffeine alone had no effect on the flickers (Fig. 6B). Cyclosporin A (10 µM) was then added to the bath, and mitochondria were imaged at 0.5, 5, and 10 min after its application. Flickers were still apparent 10 min after the addition of cyclosporin A to the bath (Fig. 6A). Cyclosporin A did not change the percentage of flickering mitochondria, the rate of flickering, or flicker amplitude (Fig. 6B). In addition, cells incubated in 10 µM cyclosporin A alone for periods ranging from 45 to 90 min continued to flicker. Finally, the ROS scavenger trolox (1 mM for up to 10 min) did not abolish mitochondrial flickering. The data indicate that PTP activation is not the mechanism underlying spontaneous transient mitochondrial flickers.
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DISCUSSION |
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Mitochondrial flickers and the PTP. In all studies in which flickers were attributed to ROS generation or PTP activation, flickers preceded an irreversible global mitochondrial depolarization. Inhibition of the PTP by cyclosporin A prevents mitochondrial depolarization in astrocytes (16, 28) and cardiomyocytes (3). In the present study cyclosporin A had no effect on mitochondrial flickers. It has been reported that in some cell preparations cyclosporin A fails to inhibit the PTP (25, 49). However, the patch pipette solution contained Mg2+ and ATP, which inhibit Ca2+-mediated activation of the PTP (22, 24). Mg2+ also facilitates PTP inhibition by cyclosporin A (2, 33). The ROS scavenger trolox inhibits the PTP in cardiomyocytes and Cos-7 cells (12, 49). In contrast, trolox did not inhibit mitochondrial flickering in the cells studied here (data not shown). We conclude that flickers do not reflect progressive failure of mitochondrial function due to phototoxic activation of the PTP. In neurons, irreversible depolarization of the mitochondria does not occur during imaging, and consequently mitochondrial flickering is not attributed to phototoxicity (4). Additionally, Jacobson and Duchen (28) report that with low illumination, flickers are smaller, more infrequent, and do not result in cell death. Therefore, it appears that flickers that are independent of phototoxic events can occur in several cell types.
The PTP can operate in a low-conductance state, which does not impair mitochondrial function. It has been proposed that rapid uptake of Ca2+ into the mitochondria after localized release from stores activates the low-conductance PTP (27). However, in light of the lack of inhibition of flickers by manipulations designed to prevent Ca2+ signaling, it is unlikely that Ca2+-dependent low-conductance PTP activity underlies flickers. Furthermore, reversal of PTP activation occurs rapidly after chelation of free Ca2+ (11, 21, 24). Here, buffering of cytosolic Ca2+ with BAPTA-AM failed to abolish the flickers. It is known that the PTP can switch from low to high conductance after an increase in [Ca2+]m or a slight mitochondrial depolarization (2, 27). Such an increase in [Ca2+]m is expected after caffeine application in the presence of thapsigargin, i.e., in the immediate aftermath of store depletion. If flickers were caused by low-conductance PTP activity, we might expect that store depletion would result in global mitochondrial depolarization due to Ca2+-dependent activation of high-conductance PTP. This was not observed after store depletion. The accumulated evidence indicates that flickering is not caused by induction of PTP in these cells.
Molecular basis of mitochondrial flickers. Many ion transporters have been identified in the inner mitochondrial membrane, several of which might cause flickers (37). Entry of H+ into the matrix via the F1F0ATPase during ATP generation (4) or through uncoupling proteins, recently identified in intestinal smooth muscle (42), may result in mitochondrial depolarization. Opening of mitochondrial K+ channels may also result in depolarization. In isolated mitochondria, activation of ATP-sensitive K+ (KATP) channels in the inner mitochondrial membrane has been shown to result in m depolarization, although the extent of depolarization is currently debated (23, 30). KATP channel opening is unlikely in the present study as the patch pipette included ATP. However, Ca2+-activated K+ channels were recently identified in mitochondria of glioma and cardiac cells (43, 45), and K+ entry into the mitochondrial matrix via these channels may also elicit
m depolarization. Finally, anion channels in the inner mitochondrial membrane have been linked to PTP-independent oscillations in
m in substrate-deprived myocytes (37).
Their reversibility and independence from spontaneous intracellular Ca2+ release events and PTP activity indicate that flickers represent a previously unknown process in myocyte mitochondria. The quantitative analysis of flickers used here and in our earlier study (36) should be useful in the further elucidation of the mechanism and role of mitochondrial flickers in smooth muscle and other cell types.
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ACKNOWLEDGMENTS |
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Present address for R. M. Drummond: Dept. of Physiology and Pharmacology, Univ. of Strathclyde, Glasgow, UK.
GRANTS
This research was funded by National Heart, Lung, and Blood Institute Grant HL-61297 to J. V. Walsh, Jr.
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FOOTNOTES |
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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.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
2. Bernardi P. Mitochondrial transport of cations: channels, exchangers, and permeability transition. Physiol Rev 79: 1127-1155, 1999.
3. Bowser DN, Minamikawa T, Nagley P, and Williams DA. Role of mitochondria in calcium regulation of spontaneously contracting cardiac muscle cells. Biophys J 75: 2004-2014, 1998.
4. Buckman JF and Reynolds IJ. Spontaneous changes in mitochondrial membrane potential in cultured neurons. J Neurosci 21: 5054-5065, 2001.
5. Carafoli E. Historical review. Mitochondria and calcium: ups and downs of an unusual relationship. Trends Biochem Sci 28: 175-181, 2003.[CrossRef][ISI][Medline]
6. Carrington WA, Fogarty KE, and Fay FS. 3D fluorescence imaging of single cells using image restoration. In: Noninvasive Techniques in Cell Biology, edited by Foster K. New York: Wiley-Liss, 1990, p. 53-72.
7. Carrington WA, Lynch RM, Moore ED, Isenberg G, Fogarty KE, and Fay FS. Superresolution three-dimensional images of fluorescence in cells with minimal light exposure. Science 268: 1483-1487, 1995.[ISI][Medline]
8. Cheng H, Lederer MR, Xiao RP, Gomez AM, Zhou YY, Ziman B, Spurgeon H, Lakatta EG, and Lederer WJ. Excitation-contraction coupling in heart: new insights from Ca2+ sparks. Cell Calcium 20: 129-140, 1996.[ISI][Medline]
9. Cheng H, Lederer WJ, and Cannell MB. Calcium sparks: elementary events underlying excitation-contraction coupling in heart muscle. Science 262: 740-744, 1993.[ISI][Medline]
10. Collins TJ, Berridge MJ, Lipp P, and Bootman MD. Mitochondria are morphologically and functionally heterogeneous within cells. EMBO J 21: 1616-1627, 2002.
11. Crompton M and Costi A. Kinetic evidence for a heart mitochondrial pore activated by Ca2+, inorganic phosphate and oxidative stress. A potential mechanism for mitochondrial dysfunction during cellular Ca2+ overload. Eur J Biochem 178: 489-501, 1988.[Abstract]
12. De Giorgi F, Lartigue L, and Ichas F. Electrical coupling and plasticity of the mitochondrial network. Cell Calcium 28: 365-370, 2000.[CrossRef][ISI][Medline]
13. Diaz G, Falchi AM, Gremo F, Isola R, and Diana A. Homogeneous longitudinal profiles and synchronous fluctuations of mitochondrial transmembrane potential. FEBS Lett 475: 218-224, 2000.[CrossRef][ISI][Medline]
14. Drummond RM, Mix TCH, Tuft RA, Walsh JV Jr., and Fay FS. Mitochondrial Ca2+ homeostasis during Ca2+ influx and Ca2+ release in gastric myocytes from Bufo marinus. J Physiol 522: 375-390, 2000.
15. Drummond RM and Tuft RA. Release of Ca2+ from the sarcoplasmic reticulum increases mitochondrial [Ca2+] in rat pulmonary artery smooth muscle cells. J Physiol 516: 139-147, 1999.
16. Duchen MR. Mitochondria and Ca2+ in cell physiology and pathophysiology. Cell Calcium 28: 339-348, 2000.[CrossRef][ISI][Medline]
17. Duchen MR, Leyssens A, and Crompton M. Transient mitochondrial depolarizations reflect focal sarcoplasmic reticular calcium release in single rat cardiomyocytes. J Cell Biol 142: 975-988, 1998.
18. Fall CP and Bennett JP Jr. Visualization of cyclosporin A and Ca2+-sensitive cyclical mitochondrial depolarizations in cell culture. Biochim Biophys Acta 1410: 77-84, 1999.[ISI][Medline]
19. Fay FS, Hoffmann R, Leclair S, and Merriam P. Preparation of individual smooth muscle cells from the stomach of Bufo marinus. Methods Enzymol 85: 284-292, 1982.[ISI][Medline]
20. Gafni J, Munsch JA, Lam TH, Catlin MC, Costa LG, Molinski TF, and Pessah IN. Xestospongins: potent membrane permeable blockers of the inositol 1,4,5-trisphosphate receptor. Neuron 19: 723-733, 1997.[ISI][Medline]
21. Gunter TE, Gunter KK, Sheu SS, and Gavin CE. Mitochondrial calcium transport: physiological and pathological relevance. Am J Physiol Cell Physiol 267: C313-C339, 1994.
22. Haworth RA and Hunter DR. The Ca2+-induced membrane transition in mitochondria. II. Nature of the Ca2+ trigger site. Arch Biochem Biophys 195: 460-467, 1979.[ISI][Medline]
23. Holmuhamedov EL, Wang L, and Terzic A. ATP-sensitive K+ channel openers prevent Ca2+ overload in rat cardiac mitochondria. J Physiol 519: 347-360, 1999.
24. Hunter DR, Haworth RA, and Southard JH. Relationship between configuration, function, and permeability in calcium-treated mitochondria. J Biol Chem 251: 5069-5077, 1976.[Abstract]
25. Hüser J and Blatter LA. Fluctuations in mitochondrial membrane potential caused by repetitive gating of the permeability transition pore. Biochem J 343: 311-317, 1999.[CrossRef][ISI][Medline]
26. Hüser J, Rechenmacher CE, and Blatter LA. Imaging the permeability pore transition in single mitochondria. Biophys J 74: 2129-2137, 1998.
27. Ichas F and Mazat JP. From calcium signaling to cell death: two conformations for the mitochondrial permeability transition pore. Switching from low- to high-conductance state. Biochim Biophys Acta 1366: 33-50, 1998.[ISI][Medline]
28. Jacobson J and Duchen MR. Mitochondrial oxidative stress and cell death in astrocytesrequirement for stored Ca2+ and sustained opening of the permeability transition pore. J Cell Sci 115: 1175-1188, 2002.
29. Jouaville LS, Pinton P, Bastianutto C, Rutter GA, and Rizzuto R. Regulation of mitochondrial ATP synthesis by calcium: evidence for a long-term metabolic priming. Proc Natl Acad Sci USA 96: 13807-13812, 1999.
30. Kowaltowski AJ, Seetharaman S, Paucek P, and Garlid KD. Bioenergetic consequences of opening the ATP-sensitive K+ channel of heart mitochondria. Am J Physiol Heart Circ Physiol 280: H649-H657, 2001.
31. Loew LM, Carrington W, Tuft RA, and Fay FS. Physiological cytosolic Ca2+ transients evoke concurrent mitochondrial depolarizations. Proc Natl Acad Sci USA 91: 12579-12583, 1994.
32. Loew LM, Tuft RA, Carrington W, and Fay FS. Imaging in five dimensions: time-dependent membrane potentials in individual mitochondria. Biophys J 65: 2396-2407, 1993.[Abstract]
33. Novgorodov SA, Gudz TI, Brierley GP, and Pfeiffer DR. Magnesium ion modulates the sensitivity of the mitochondrial permeability transition pore to cyclosporin A and ADP. Arch Biochem Biophys 311: 219-228, 1994.[CrossRef][ISI][Medline]
34. O'Reilly CM, Drummond RM, Fogarty KE, Tuft RA, and Walsh JV Jr. Mitochondrial flickers do not require intracellular Ca2+ release in smooth muscle cells. Biophys J 80: 236a, 2001.
35. O'Reilly CM, Drummond RM, Fogarty KE, Tuft RA, and Walsh JV Jr. Mitochondrial flickers occur in the absence of Ca2+ sparks in smooth muscle cells. J Gen Physiol 116: 50A, 2000.
36. O'Reilly CM, Fogarty KE, Drummond RM, Tuft RA, and Walsh JV Jr. Quantitative analysis of spontaneous mitochondrial depolarizations. Biophys J 85: 3350-3357, 2003.
37. O'Rourke B. Pathophysiological and protective roles of mitochondrial ion channels. J Physiol 529: 23-36, 2000.
38. Pacher P, Thomas AP, and Hajnoczky G. Ca2+ marks: miniature calcium signals in single mitochondria driven by ryanodine receptors. Proc Natl Acad Sci USA 99: 2380-2385, 2002.
39. Rizzuto R, Brini M, Murgia M, and Pozzan T. Microdomains with high Ca2+ close to IP3-sensitive channels that are sensed by neighboring mitochondria. Science 262: 744-747, 1993.[ISI][Medline]
40. Rizzuto R, Pinton P, Carrington W, Fay FS, Fogarty KE, Lifshitz LM, Tuft RA, and Pozzan T. Close contacts with the endoplasmic reticulum as determinants of mitochondrial Ca2+ responses. Science 280: 1763-1766, 1998.
41. Scaduto RC Jr. and Grotyohann LW. Measurement of mitochondrial membrane potential using fluorescent rhodamine derivatives. Biophys J 76: 469-477, 1999.
42. Shabalina I, Wiklund C, Bengtsson T, Jacobsson A, Cannon B, and Nedergaard J. Uncoupling protein-1: involvement in a novel pathway for -adrenergic, cAMP-mediated intestinal relaxation. Am J Physiol Gastrointest Liver Physiol 283: G1107-G1116, 2002.
43. Siemen D, Loupatatzis C, Borecky J, Gulbins E, and Lang F. Ca2+-activated K channel of the BK-type in the inner mitochondrial membrane of a human glioma cell line. Biochem Biophys Res Commun 257: 549-554, 1999.[CrossRef][ISI][Medline]
44. Tinel H, Cancela JM, Mogami H, Gerasimenko JV, Gerasimenko OV, Tepikin AV, and Petersen OH. Active mitochondria surrounding the pancreatic acinar granule region prevent spreading of inositol trisphosphate-evoked local cytosolic Ca2+ signals. EMBO J 18: 4999-5008, 1999.
45. Xu W, Liu Y, Wang S, McDonald T, Van Eyk JE, Sidor A, and O'Rourke B. Cytoprotective role of Ca2+-activated K+ channels in the cardiac inner mitochondrial membrane. Science 298: 1029-1033, 2002.
46. Yao Y, Choi J, and Parker I. Quantal puffs of intracellular Ca2+ evoked by inositol trisphosphate in Xenopus oocytes. J Physiol 482: 533-553, 1995.[Abstract]
47. ZhuGe R, Fogarty KE, Tuft RA, Lifshitz LM, Sayar K, and Walsh JV Jr. Dynamics of signaling between Ca2+ sparks and Ca2+-activated K+ channels studied with a novel image-based method for direct intracellular measurement of ryanodine receptor Ca2+ current. J Gen Physiol 116: 845-864, 2000.
48. ZhuGe R, Tuft RA, Fogarty KE, Bellve K, Fay FS, and Walsh JV Jr. The influence of sarcoplasmic reticulum Ca2+ concentration on Ca2+ sparks and spontaneous transient outward currents in single smooth muscle cells. J Gen Physiol 113: 215-228, 1999.
49. Zorov DB, Filburn CR, Klotz LO, Zweier JL, and Sollott SJ. Reactive oxygen species (ROS)-induced ROS release: a new phenomenon accompanying induction of the mitochondrial permeability transition in cardiac myocytes. J Exp Med 192: 1001-1014, 2000.
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