Department of Physiology, University of Maryland School of Medicine, Baltimore, Maryland 21201
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ABSTRACT |
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The role of mitochondria in Ca2+ homeostasis is controversial. We employed the Ca2+-sensitive dye rhod 2 with novel, high temporal and spatial resolution imaging to evaluate changes in the matrix free Ca2+ concentration of individual mitochondria ([Ca2+]m) in agonist-stimulated, primary cultured aortic myocytes. Stimulation with 10 µM serotonin (5-HT) evoked modest cytosolic Ca2+ transients [cytosolic free Ca2+ concentration ([Ca2+]cyt) <500 nM; measured with fura 2] and triggered contractions in short-term cultured myocytes. However, 5-HT triggered a large mitochondrial rhod 2 signal (indicating pronounced elevation of [Ca2+]m) in only 4% of cells. This revealed heterogeneity in the responses of individual mitochondria, all of which stained with MitoTracker Green FM. In contrast, stimulation with 100 µM ATP evoked large cytosolic Ca2+ transients (>1,000 nM) and induced pronounced, reversible elevation of [Ca2+]m (measured as rhod 2 fluorescence) in 60% of cells. This mitochondrial Ca2+ uptake usually lagged behind the cytosolic Ca2+ transient peak by 3-5 s, and [Ca2+]m declined more slowly than did bulk [Ca2+]cyt. The uptake delay may prevent mitochondria from interfering with rapid signaling events while enhancing the mitochondrial response to large, long-duration elevations of [Ca2+]cyt. The responses of arterial myocytes to modest physiological stimulation do not, however, depend on such marked changes in [Ca2+]m.
mitochondria; rhod 2; vascular smooth muscle; fluorescence digital imaging
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INTRODUCTION |
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MOBILIZATION OF STORED Ca2+ and elevation of cytosolic free Ca2+ concentration ([Ca2+]cyt) are important in cellular signaling (4). The central role of the sarcoplasmic reticulum/endoplasmic reticulum (SR/ER) in the modulation of [Ca2+]cyt through Ca2+ sequestration and mobilization is well documented, but the role of mitochondria is still controversial. One view is that mitochondria sequester large amounts of Ca2+ only at the high [Ca2+]cyt levels associated with cytotoxicity (3, 37). Indeed, the slow kinetics of the Ca2+ uptake pathway indicated that little Ca2+ was accumulated by mitochondria during physiological stimulation (Ref. 39, and see Ref. 38). Nevertheless, mitochondrial uptake of Ca2+ during physiological stimulation might effectively regulate the Krebs cycle (11, 23).
Development of selective Ca2+-sensitive indicators and application of photometry and video microscopy have led to a reassessment of the role of mitochondria in Ca2+ homeostasis. This has fostered the alternative view that mitochondria buffer and/or modulate [Ca2+]cyt during and after activation in many types of cells (12-14, 16, 18, 36), including amphibian smooth muscle (5). Furthermore, studies on isolated mitochondria led to the identification of a rapid Ca2+ uptake mode that should be modulated by [Ca2+]cyt during agonist stimulation (38). Mitochondria may regulate [Ca2+]cyt in cytosolic microdomains and thereby modulate ion channels (14, 36).
In many of the aforementioned studies, aequorin or rhod 2 was used to assess the intramitochondrial free Ca2+ concentration ([Ca2+]m) in single cells or cell populations, often with a photomultiplier tube (PMT); in some cases, very high extracellular Ca2+ concentrations were employed (1, 5). A PMT offers high temporal resolution but no spatial resolution; moreover, to avoid confusion between cytosolic and mitochondrial signals, it requires almost complete elimination of cytosolic rhod 2, which is difficult to achieve (14). Most published imaging studies provide limited temporal resolution (<1 image/s) (16, 29, 33) and do not resolve individual mitochondria. Indo 1 also has been employed for PMT and imaging studies, but Mn2+ has been needed to quench cytosolic dye (26, 40).
The time to reach peak [Ca2+]m reportedly lags behind peak [Ca2+]cyt by ~1-6 s (1, 16, 40). Spatial resolution in these studies was, however, inadequate to resolve [Ca2+]m in individual mitochondria at rates fast enough to measure the lag between the time taken to reach peak [Ca2+]cyt and peak [Ca2+]m (e.g., Ref. 16) or to detect heterogeneity in the mitochondrial responses to cell activation (14, 33). Recently, however, Nitschke et al. (29) measured, with good spatial resolution, a very long delay (45-60 s) between increases in [Ca2+]cyt and [Ca2+]m in carbachol-stimulated HT-29 (colon carcinoma) cells. Good spatial resolution is essential because indirect evidence indicates that the position of mitochondria, relative to SR/ER Ca2+ release sites and/or Ca2+ influx sites, may be crucial in determining whether mitochondria sequester Ca2+ (20, 31, 32).
Novel wide-field digital imaging methods now enable us to assess, for the first time, dynamic changes in [Ca2+]m in individual mitochondria with high spatial resolution (8) and high temporal resolution (>4 images/s). We used this technology to examine the temporal and spatial relationships between [Ca2+]cyt elevation and mitochondrial Ca2+ accumulation in individual mitochondria during activation of aortic myocytes.
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MATERIALS AND METHODS |
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Cell culture. Aortic smooth muscle cells from Sprague-Dawley rats were isolated and cultured in media containing 10% fetal bovine serum (FBS) as described (35). Cultures 3 to 6 days old were serum deprived for 24-32 h before loading with Ca2+ indicator. More than 95% of the cells stained positively for smooth muscle actin. Unless otherwise stated, subconfluent cultures were used because cell flattening and lack of overcrowding improved spatial resolution of intracellular organelles.
Some studies were performed on myocytes that were maintained in the contractile state. For these studies, the isolated myocytes were cultured in FBS-free DMEM containing 0.3% BSA; cells were used <72 h after plating.Measurement of
[Ca2+]cyt.
Cultured myocytes, on coverslips, were incubated in FBS-free DMEM
containing 0.3% BSA and fura 2-AM (4 µM, 45 min, 22-24°C, 5% CO2-95%
O2). Coverslips were then
mounted in a chamber on a microscope stage and were superfused for
20-25 min with physiological salt solution (PSS; in mM: 140 NaCl,
5.9 KCl, 1.2 NaH2PO4,
1.8 MgCl2, 1.8 CaCl2, 11.1 glucose, 10 HEPES, and
4.17 NaHCO3, pH 7.4) at
32-34°C. Fura 2 fluorescence was observed using a 40× oil immersion (1.3 numerical aperture) fluorescence objective mounted
on an Axiovert 100 microscope (Carl Zeiss, Thornwood, NY). Images from
a Gen III ultrablue intensified charge-coupled device camera (Stanford
Photonics, Palo Alto, CA) were digitized (512 × 480 pixels at 8 bits) and collected using MetaFluor software (Universal Imaging, West
Chester, PA). Cells were excited at 350 and 380 nm using a Polychrome
II illumination system (Applied Scientific Instruments, Eugene, OR).
Light (12-nm bandwidth) was passed through a UG1 filter; all excitation
light was passed through a KG1 filter. Excitation intensity was
aperture controlled to minimize photodamage. Four image frames (510-nm
emission; 40-nm bandpass filter) were averaged at each excitation
wavelength. During periods of rapid changes in
[Ca2+]cyt,
ratio pairs were acquired at fast rates (1 pair/s); at other times,
the rate was reduced (
0.2 pairs/s) to minimize photodamage and dye
bleaching. Chamber contents (volume ~0.25 ml) were exchanged at a
flow rate of 2.5 ml/min.
Measurement of [Ca2+]m. To assess changes in [Ca2+]m with rhod 2, we modified the method of Mix and colleagues (28). Cells were loaded with 8 µM rhod 2-AM for 90 min (37°C, 5% CO2-95% air) in FBS-free DMEM containing 0.3% BSA. Coverslips were rinsed with FBS-free DMEM containing 0.3% BSA and kept at 37°C (5% CO2-95% air) for a further 30 min. The delocalized positive charge of rhod 2 (25) and incubation at 37°C (8) promote dye sequestration into mitochondria; the dye is insensitive to mitochondrial membrane potential (1). Some dye was retained in the cytosol; nevertheless, with high spatial resolution imaging (8), cytosolic rhod 2 could be resolved from dye in other compartments by its diffuse staining pattern (14) and different response to stimulation (see RESULTS). A less sensitive intensified camera would not have detected the weak fluorescence signal from the residual cytosolic rhod 2 and would have detected only the large increases in mitochondrial rhod 2 fluorescence during stimulation. The loading protocol employed here was optimal for loading mitochondria with rhod 2 for these high temporal and spatial resolution experiments. Our arterial myocytes did not tolerate overnight incubation after rhod 2 loading (12). Optimization of rhod 2 loading protocols appears to be cell type specific; for example, we found that the protocol optimized for arterial myocytes was unsuitable for hippocampal neurons, which required a different loading protocol (unpublished data).
Coverslips were mounted on the microscope stage and superfused with PSS as described above. Rhod 2 was excited at 548 nm (10-nm bandwidth); images were acquired at 605 nm (55-nm bandpass). Unless otherwise stated, two to eight video frames were averaged during image acquisition. Quantitative mitochondrial data (e.g., see Fig. 1E) are based on 3 × 3-pixel areas (~1 µm2; see Fig. 1D). Images were acquired at rates ranging from 5 toSimultaneous determination of [Ca2+]cyt and changes in [Ca2+]m. Aortic smooth muscle cells, on coverslips, were incubated with 8 µM rhod 2-AM for 90 min (37°C, 5% CO2-air) in FBS-free DMEM containing 0.3% BSA. The coverslips were then rinsed with FBS-free DMEM-0.3% BSA as above. The cells were then incubated in FBS-free DMEM containing 0.3% BSA and fura 2-AM (4 µM, 45 min, 22-24°C, 5% CO2-95% O2). The coverslips were rinsed with FBS-free DMEM-0.3% BSA, mounted on the microscope stage, and superfused with PSS (20-25 min, 32-34°C). The cells were imaged as described above. Fura 2 was excited at 350 and 380 nm, and rhod 2 was excited at 550 nm; a Chroma Technologies fura 2-rhodamine beam splitter and a dual-band emission filter were used for these experiments.
Identification of mitochondria by staining with MTG. Either before or after experiments (see RESULTS), coverslips were superfused with 100 nM MTG in PSS (20-30 min, 32-34°C). MTG was excited at 488 nm; fluorescent emission was detected at 535 nm (40-nm bandpass). At the equivalent intensifier gain, no mitochondrial fluorescence signal was detected at 535 nm (488-nm excitation) in cells loaded with rhod 2 before loading with MTG. During MTG image acquisition, 16-32 frames were averaged.
Many structures with morphology typical of mitochondria were stained by MTG (also see Ref. 14); this was clearly evident in high-magnification images (not shown here, but see Ref. 8). Moreover, all of the structures that were clearly stained by MTG had the morphology of mitochondria.Materials. Fura 2-AM, MTG, and rhod 2-AM were purchased from Molecular Probes (Eugene, OR). ATP, DMSO, 5-HT, oligomycin, FCCP, rotenone, and carbonyl cyanide m-chlorophenylhydrazone were obtained from Sigma (St. Louis, MO). All other reagents were analytical grade. Agonist solutions were prepared fresh daily. All optical filters were from Chroma Technologies (Brattleboro, VT).
Movies in Quicktime format. Movies in Quicktime format can be downloaded in binary mode via file transfer protocol (FTP) from 134.192.128.114 (login: AJP; password: AJP; the login and password are case sensitive). The movies are stored as smaller size windows (file names end in "A"), and as larger size windows, which show more detail (file names end in "B"). The movies are sequential fluorescence images; frame-by-frame analysis enables temporal differences in [Ca2+]cyt and [Ca2+]m to be visualized. Resolution of images was reduced for Quicktime format. Quicktime movie viewers can be downloaded from http://quicktime.apple.com.
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RESULTS |
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Effects of 5-HT on [Ca2+]cyt and dynamic changes in [Ca2+]m. A critical question is whether [Ca2+]m increases during stimulation with physiological agonists. To address this question, we employed the Ca2+-sensitive dye rhod 2 in primary cultured rat aortic myocytes. This dye accumulates in mitochondria, but a small amount remains in the cytosol; high-resolution wide-field imaging enabled us to resolve the mitochondrial signal from that of the surrounding cytosol.
Initially, we tested the effects of 10 µM 5-HT. This evoked modest increases in [Ca2+]cyt as measured with fura 2 (basal [Ca2+]cyt 82 ± 30 nM, n = 1,022; 10 µM 5-HT peak [Ca2+]cyt 350 ± 248 nM, n = 843 cells; means ± SD). Data from two representative rhod 2 experiments are presented in Figs. 1 and 2. Quiescent cells (Fig. 1A) typically exhibited perinuclear SR fluorescence (the ground glass appearance around the nucleus) and small, bright spherical areas on the nucleus (probably nucleoli; Refs. 1, 14). Rhod 2 [dissociation constant (Kd) ~1 µM] sequestered in the SR (Ca2+ concentration in the SR >60 µM; Refs. 8, 27) should be saturated with Ca2+ and should not contribute to dynamic changes in rhod 2 fluorescence. Also, the signal from dye within the cytosol of resting cells was low (Fig. 1A). Therefore, in many later experiments (e.g., Fig. 2), the initial (control) image was subtracted from all subsequent images.
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Effects of ionomycin on
[Ca2+]m.
Heterogeneity among cells and among individual mitochondria could be
the result of a number of physiological factors (see Effects of large elevations of
[Ca2+]cyt
on
[Ca2+]m).
Alternatively, some mitochondria may not load with rhod 2 or may be
incapable of accumulating Ca2+. To
examine this possibility, cells were treated with ionomycin to induce a
high level of
[Ca2+]cyt,
to eliminate Ca2+ gradients
between the mitochondrial matrix and the cytosol, and to saturate rhod
2 with Ca2+ (extracellular
Ca2+ is 1.8 mM). In contrast to
5-HT, 5-10 µM ionomycin (45 s), which had no effect on
fluorescence in the absence of rhod 2, consistently increased
mitochondrial rhod 2 fluorescence (see Fig.
3B) and eventually caused the mitochondria to round up (not shown). This demonstrates that heterogeneity in mitochondrial rhod 2 fluorescence could not simply be attributed to heterogeneity in rhod 2 loading of
individual mitochondria. The brighter rhod 2 fluorescence signal from
mitochondria is consistent with greater accumulation of rhod 2 into
mitochondria (1, 14) than in the cytosol. This is predictable because
of the delocalized positive charges on rhod 2 (25) and the expected
sequestration of dyes during long incubations (>1 h) at 37°C (8).
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Effects of ATP on
[Ca2+]cyt
and changes in
[Ca2+]m.
ATP evokes large increases in
[Ca2+]cyt
in smooth muscle cells (17, 21). In rat aortic myocytes, ATP (100 µM,
30 s) consistently induced higher peak
[Ca2+]cyt
levels (1,029 ± 436 nM, n = 179;
mean ± SD) than did 5-HT (P < 0.001). The ATP was also much more effective than 10 or 100 µM 5-HT
(45 s) in elevating
[Ca2+]m;
indeed, ATP induced large increases in mitochondrial fluorescence in a
majority of cells (Fig. 5). ATP, like 5-HT,
evoked an initial, diffuse increase in cytosolic rhod 2 fluorescence
(Fig. 5A; also see movie CMOV3).
Subsequently, numerous small areas of intense rhod 2 fluorescence
appeared (Fig. 5B). These areas
continued to fluoresce after
[Ca2+]cyt
(i.e., cytosolic fluorescence) returned to near resting levels (Fig.
5C), but the intensity gradually
declined to background levels during continued superfusion with PSS
(Fig. 5D; also see movie CMOV3). The
small areas that accumulated Ca2+
(Fig. 5B) also stained with MTG
(Fig. 5E). Even when 10 µM 5-HT did not elevate
[Ca2+]m
substantially (Fig. 5G), subsequent
application of 100 µM ATP did (Fig.
5H). Thus the absence of such a
response to 5-HT did not mean that the mitochondria did not load with
rhod 2 or that they were incapable of accumulating
Ca2+.
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ATP induced
[Ca2+]m
waves in cultured aortic myocytes.
Figure 7 shows a group of myocytes during
stimulation with 100 µM ATP. Images captured sequentially during and
immediately after the addition of ATP revealed both intracellular and
intercellular [Ca2+]cyt
waves, as well as the expected
[Ca2+]m
elevations that follow the localized increases in
[Ca2+]cyt.
For example, in cell
A (see Fig. 7,
bottom
right), the rise in
[Ca2+]cyt
(and
[Ca2+]m)
spread from the lower portion to the upper portion of the cell. Also,
the increases in
[Ca2+]cyt
and
[Ca2+]m
were first observed in cell
A, then in the other cells at the right side of the field, next in the cells in the middle (including cell
B), and finally in the cells at the
left side of the field. In all cases,
[Ca2+]cyt
rose before
[Ca2+]m.
A movie from this experiment can be downloaded (movie CMOV4; see
Movies in Quicktime format).
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High temporal resolution studies in ATP-stimulated cells.
Figures 1, 2, 5, and 7 show that, after stimulation by either 5-HT or
ATP,
[Ca2+]m
declined much more slowly than did
[Ca2+]cyt.
The data also reveal that mitochondrial uptake of
Ca2+ was often delayed relative to
the rise of
[Ca2+]cyt.
To characterize this effect further, we used high temporal and spatial
resolution methods to image individual mitochondria and to measure
changes in
[Ca2+]m
directly and simultaneously with
[Ca2+]cyt
(Fig. 8). The delay between peak
[Ca2+]cyt
and peak
[Ca2+]m
was determined during the initial portion of the response to ATP in
small areas without frame averaging. This reduced image quality, but
spatial and temporal differences in rhod 2 fluorescence were still
clear (Fig. 8). Figure 8Aa is the
initial F/F image (monochrome scale) before the introduction of ATP.
Figure 8Ab shows the diffuse increase
in cytosolic fluorescence (i.e., the rise in
[Ca2+]cyt)
evoked by 100 µM ATP. About 2.5-3 s later, small, intensely fluorescent spots, indicative of large increases in
[Ca2+]m,
appeared (Fig. 8Ac); many of these
spots continued to fluoresce when
[Ca2+]cyt
declined (Fig. 8Ad) to resting
levels (Fig. 8Ae). Thus the increased image acquisition rate confirmed the delay in the rise of
[Ca2+]m.
The
x-z
projections (line scans in which z is
time) of the
F/F values along the dashed lines
(x) in images in Fig.
8A labeled Mi and Mj
(which traverse 2 mitochondria; see boxes labeled Ma and Mb in Fig.
8Ae) clearly illustrate this delay.
This is also depicted in Fig. 8B, in
which the period of ATP stimulation is shown on an expanded time scale:
the fluorescence in boxes Ma and Mb initially rises rapidly and then
slows for a few seconds before rising rapidly again. Figure
8B,
inset, shows the mitochondrial and
cytosolic
F/F values scaled so that the maxima are superimposed; this better illustrates the much slower decline of fluorescence in the
mitochondria than in the cytosol.
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5-HT- and ATP-evoked responses in contracting aortic myocytes.
The preceding studies show that mitochondria can sequester
Ca2+ after stimulation with
physiological agonists; the amount of Ca2+ accumulated appears to be
linked to the degree of stimulation and elevation of
[Ca2+]cyt.
The question remains, however, as to where this mitochondrial Ca2+ sequestration fits into the
spectrum of physiological and pathophysiological responses. To address
this question, aortic myocytes were cultured in serum-free medium;
these myocytes retained their contractile properties (see
Cell culture). Like
cells cultured in serum-containing medium (Fig. 1), these cells
responded to 10 µM 5-HT with transient increases in bulk
[Ca2+]cyt
(Fig.
9B),
followed by large increases in
[Ca2+]m
in only a few mitochondria (Fig.
9C); these myocytes, however, also
contracted (Fig. 9D, compared with
Fig. 9C). Subsequent stimulation of
the same myocytes with 100 µM ATP again increased
[Ca2+]cyt
(Fig. 9F) but then markedly
increased
[Ca2+]m
in many mitochondria (dark spots in Fig.
9G) and induced a large contraction
(see Fig. 9H).
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DISCUSSION |
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Rhod 2 fluorescence bright spots are mitochondria. Using high spatial resolution digital imaging, we observed that brief exposure of primary cultured rat aortic myocytes to physiological agonists increased mitochondrial fluorescence (equivalent to elevating [Ca2+]m) substantially in some mitochondria. The fraction of mitochondria affected appears to be directly related to the magnitude of the evoked rise in [Ca2+]cyt (40). These increases in [Ca2+]m were detected as small, often rod-shaped, spots of intense rhod 2 fluorescence that appeared a few seconds after the rise in [Ca2+]cyt and persisted after recovery of [Ca2+]cyt. Several observations indicate that these bright spots correspond to mitochondria: 1) mitochondria preferentially accumulate rhod 2 (1, 12, 16, 28), and ionomycin, which saturates the dye with Ca2+ (Ca2+-bound rhod 2 is the fluorescent form), induced much more fluorescence in these organelles than in the surrounding cytoplasm (Figs. 3 and 4); 2) the bright spots often had characteristic rod-shaped morphology, and many were located in the perinuclear region; 3) the bright spots were stained by the mitochondrial marker, MTG. Furthermore, consistent with previous studies (8), ATP-induced increases in fluorescence intensity in the small spots were prevented by pretreating myocytes for 30 s with the mitochondrial uncoupler FCCP (2 µM; not shown). Previously, we demonstrated that these FCCP-sensitive organelles are mitochondria, in part on the basis of their high Mg2+ and low free Ca2+ (200 nM) levels, their location and morphology, and their affinity for the lipophilic, cationic stain 3,3'-dihexyloxacarbocyanine (8).
Effects of modest elevation of [Ca2+]cyt on [Ca2+]m. The mitochondria and cytosol in quiescent myocytes exhibited negligible rhod 2 signal, relative to the Ca2+-saturated rhod 2 signal emanating from the SR (Fig. 1). The rhod 2 Kd for Ca2+ is 570 nM in Mg2+-free solution (Molecular Probes) and is probably much higher in situ. Therefore, these data imply that resting [Ca2+]m and [Ca2+]cyt were low, even though the rhod 2 signals were not calibrated because of uncertainties inherent in nonratiometric determinations. Our data are consistent with reported levels of [Ca2+]m in the range of 80-200 nM (1, 8, 26, 34) and similarly low levels of [Ca2+]cyt (82 nM; see Effects of 5-HT on [Ca2+]cyt and dynamic changes in [Ca2+]m).
Stimulation by 10 µM 5-HT elevated [Ca2+]cyt to 350 nM, on average, whereas the rhod 2 signal from most mitochondria in most cells was not detectably different from that in the surrounding cytosol. This implies that [Ca2+]m did not rise above [Ca2+]cyt in most of the mitochondria in these cells; nevertheless, this stimulus was sufficient to induce shortening in contraction-competent cells (Fig. 9). It is important to note, however, that the methods used here precluded detection of small changes in [Ca2+]m. Clearly, small increases and decreases in [Ca2+]m that paralleled the changes in [Ca2+]cyt would have been masked by the fluorescence signal from the cytosol. Any increases in [Ca2+]m, if they did occur, must have been smaller than the increases in [Ca2+]cyt, since results with ionomycin (Figs. 3 and 4) indicate that the concentration of rhod 2 in mitochondria is far greater than that in the cytosol (1, 14). This Ca2+ uptake could be mediated by the recently characterized mitochondrial "rapid uptake mode" (38); the kinetics also seem consistent with the rapid Ca2+ uptake detected with aequorin (20). The magnitude of this uptake may, however, have been overestimated with aequorin (20) because of difficulties in calibrating this photoprotein (1, 16).Effects of large elevations of [Ca2+]cyt on [Ca2+]m. Stimulation with 100 µM ATP increased [Ca2+]cyt to >1 µM, on average, which was about threefold higher than the level evoked by 10 µM 5-HT. In contrast to the limited effects of 10 µM 5-HT or 1 µM ATP, stimulation by 100 µM ATP was consistently associated with a substantial, sustained increase in [Ca2+]m (i.e., rhod 2 fluorescence) in numerous mitochondria in a majority of cells.
Responses of mitochondria to stimulation by both ATP and 5-HT were heterogeneous. The heterogeneity within individual cells (e.g., Fig. 2) suggests that the marked increases in [Ca2+]m were associated with microdomains of high [Ca2+]cyt. Proximity to SR Ca2+ release sites (31, 32) may be one of the underlying mechanisms, since 100 µM ATP also increased [Ca2+]m in cells bathed in Ca2+-free PSS (not shown). Mitochondria in smooth muscle (8, 15) and other cells (e.g., HeLa; see Ref. 32) often lie along elements of the SR/ER. Thus the mitochondria may be exposed to local microdomains of particularly high [Ca2+]cyt when the stored Ca2+ is mobilized. Indeed, Rizzuto and colleagues (32) used selectively targeted aequorin to obtain direct evidence for this phenomenon in HeLa cells. Our studies, with high temporal and spatial resolution, indicate directly that this mitochondrial Ca2+ uptake can be delayed by a few seconds.Kinetics of mitochondrial Ca2+ uptake and release. Comparison of absolute levels of free Ca2+ in cellular subcompartments is difficult because the local environment may affect dye spectral properties (30). Temporal differences on the order of seconds, such as those we observed between changes in [Ca2+]cyt and [Ca2+]m, are, however, unlikely to arise from differences in on and off rates for Ca2+ binding to the dye. Our imaging studies of individual mitochondria directly confirm PMT data (1, 16) showing that the rise in [Ca2+]m, detected with rhod 2, lags behind the rise in [Ca2+]cyt. Indirect studies (using cytosolic fura 2) of mitochondrial Ca2+ uptake after large Ca2+ loads also reveal slow onset kinetics (24), as do studies with indo 1 (40). The mechanism responsible for this delayed mitochondrial Ca2+ uptake apparently differs from the rapid uptake mode (38) mentioned above, which seemingly has little detectable delay. The lag might be due to the slow kinetics of the allosteric activation of mitochondrial Ca2+ uptake by Ca2+ (Ref. 19; see Ref. 10). One possibility is that the same uniporter may mediate both the rapid and delayed modes of Ca2+ uptake (38). This could occur if a fraction of the uniporter molecules were already activated by the ambient (resting) [Ca2+]cyt (i.e., the rapid component) and the remainder needed to be activated by slow, allosteric Ca2+ binding after elevation of [Ca2+]cyt. Interestingly, an analogous delay (or slow onset) in the activation of plasma membrane (PM) Na+/Ca2+ exchange by intracellular Ca2+ has been described (22). Such a delay would prevent the mitochondria close to the SR Ca2+ release sites from "working against" the SR and rapidly buffering the released Ca2+ during the rising phase of the [Ca2+]cyt transients. This would also, of course, bias this mitochondrial Ca2+ uptake pathway toward operating on the larger, longer-lasting [Ca2+]cyt transients (i.e., "amplitude modulation"; Ref. 2), as predicted by Gunter and Pfeiffer (10).
The recovery of [Ca2+]m is also slower than the decline of [Ca2+]cyt following large [Ca2+]cyt transients. Similar slow mitochondrial recovery after physiological stimuli has been reported in some low spatial resolution rhod 2 studies (Refs. 1, 16; but see Ref. 12). Indeed, delays between [Ca2+]m and [Ca2+]cyt in the time to peak and the time to recovery were predicted in early models of mitochondrial Ca2+ homeostasis (e.g., Ref. 11). Differences between cell types, stimuli, and individual mitochondria in lag, recovery, and the degree of uptake are indicative of the variety of factors that regulate mitochondrial Ca2+ uptake. For example, the rate of increase of [Ca2+]cyt, the level reached, the spatial inhomogeneities, and the duration of high [Ca2+]cyt may all influence not only the amount of Ca2+ available for sequestration in mitochondria but also the mode of Ca2+ uptake (rapid and/or slow; Ref. 38).Mitochondria and Ca2+ homeostasis in vascular smooth muscle. The main mechanisms for reduction of [Ca2+]cyt in cultured aortic myocytes after 5-HT stimulation are the PM Na+/Ca2+ exchanger and the PM and SR Ca2+ pumps (35). Our data imply that the mitochondria are not likely to act as a major sink for Ca2+ following modest physiological stimulation. In fura 2-loaded bovine artery myocytes, 1 µM norepinephrine induced marked contraction and prolonged elevation of intracellular Ca2+ concentrations in small microdomains that were believed to be located in the SR (6). However, recent evidence (8, 27) indicates that fura 2 within the SR should be saturated with Ca2+; thus these microdomains were more likely mitochondria than SR, as observed here with 100 µM ATP. Nevertheless, it appears that large accumulations of Ca2+ by mitochondria are not a necessary accompaniment of agonist-induced activation of arterial myocytes. Perhaps small elevations of [Ca2+]m, mediated by the rapid uptake mode, are sufficient to modify the activity of Krebs cycle enzymes during these physiological responses. In contrast, more intense stimulation (e.g., with 100 µM ATP) not only elevates bulk [Ca2+]cyt to the 1,000 nM range but also markedly increases [Ca2+]m and greatly stimulates mitochondrial dehydrogenases (12) when this is most needed. Whether the latter activities represent a part of the physiological spectrum or border the pathophysiological range is unclear. We do show, however, that even these pronounced mitochondrial responses are reversible and repeatable. The ability to detect changes in [Ca2+]m in individual mitochondria with high spatial and temporal resolution has revealed a complex spatial heterogeneity and has provided direct evidence that, during mobilization of large amounts of Ca2+, mitochondrial accumulation is delayed by a few seconds. This delay may prevent the mitochondria from interfering with rapid signaling events but enable them to respond to large, prolonged elevations of [Ca2+]cyt.
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ACKNOWLEDGEMENTS |
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We thank Drs. M. Juhaszova, J. P. Y. Kao, and W. G. Wier for comments on a preliminary version of this paper, K. Strauss for cell cultures, and R. S. Rogowski for technical assistance with files and figures.
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FOOTNOTES |
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This research was supported by National Heart, Lung, and Blood Institute Grant HL-45215, by a University of Sydney Eleanor Sophia Wood Fellowship to G. R. Monteith, and by funds from the University of Maryland School of Medicine and the University of Maryland, Baltimore Graduate School.
Present address of G. R. Monteith: School of Pharmacy, University of Queensland, St. Lucia, QLD 4072, Australia.
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. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: M. P. Blaustein, Dept. of Physiology, University of Maryland School of Medicine, 655 W. Baltimore St., Baltimore, MD 21201.
Received 20 November 1998; accepted in final form 11 February 1999.
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