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The Journal of Neuroscience, July 15, 2001, 21(14):5054-5065
Spontaneous Changes in Mitochondrial Membrane Potential in
Cultured Neurons
Jennifer F.
Buckman and
Ian J.
Reynolds
Department of Pharmacology, University of Pittsburgh, Pittsburgh,
Pennsylvania 15261
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ABSTRACT |
Using the mitochondrial membrane potential
(  m)-sensitive fluorescent dyes
5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolocarbocyanine iodide (JC-1) and tetramethylrhodamine methyl ester (TMRM), we have
observed spontaneous changes in the m of cultured
forebrain neurons. These fluctuations in m appear to
represent partial, transient depolarizations of individual
mitochondria. The frequency of these m
fluctuations can be significantly lowered by exposure to a
photo-induced oxidant burden, an ATP synthase inhibitor, or a
glutamate-induced sodium load, without changing overall JC-1 fluorescence intensity. These spontaneous fluctuations in JC-1 signal
were not inhibited by altering plasma membrane activity with
tetrodotoxin or MK-801 or by blocking the mitochondrial permeability transition pore (PTP) with cyclosporin A. Neurons loaded with TMRM
showed similar, low-amplitude, spontaneous fluctuations in m. We hypothesize that these m
fluctuations are dependent on the proper functioning of the
mitochondria and reflect mitochondria alternating between the active
and inactive states of oxidative phosphorylation.
Key words:
mitochondria; membrane potential; ATP; JC-1; TMRM; F1FO ATPase
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INTRODUCTION |
Mitochondria have been implicated in
excitotoxic injury pathways, as well as injury mechanisms manifested as
apoptotic or necrotic death processes. The mitochondrial membrane
potential (m) has often been used as a
marker for mitochondrial activity and neuronal viability during the
various cell death cascades (for review, see Kroemer et al., 1998 ;
Nicholls and Ward, 2000). Injurious stimuli, leading to either
excitotoxicity or apoptosis, can lead to profound depolarization of
m resulting from abnormalities in neuronal
processes, including alterations in intracellular calcium dynamics and
the opening of the mitochondrial permeability transition pore (PTP)
(Ankarcrona et al., 1995; Nieminen et al., 1996; Schinder et al., 1996;
White and Reynolds, 1996; Vergun et al., 1999; Budd et al., 2000).
Although a loss of m may be linked to
various inducers of cell death, these are observed as large and
possibly catastrophic changes in mitochondrial function.
Mitochondria under physiological conditions also play active roles in
the maintenance of normal cellular functioning. A key feature of
mitochondria that allows them to participate in cell survival is proton
pumping across the impermeable inner membrane. This generates an
electrochemical gradient, composed of m and pH, which is used for ATP synthesis, ADP-ATP exchange, uptake of
respiratory substrates and inorganic phosphate, transport of K+, Na+, and
anions to regulate volume, and regulation of protons to control heat
production (for review, see Bernardi, 1999). Mitochondria also play
protective roles by buffering cells against high concentrations of
calcium (Budd and Nicholls, 1996; White and Reynolds, 1997; Stout et
al., 1998) and sequestering proapoptotic agents, such as cytochrome c
(for review, see Green and Reed, 1998; Desagher and Martinou, 2000).
Compared with the catastrophic changes in acute injury states, healthy
mitochondria may exhibit smaller functional changes in ion transport,
ATP production or consumption, volume, and permeability, all of which
may impact m, to optimize the balance
between the need for respiration and ATP synthesis and the production
of free radicals (Nicholls and Budd, 2000). In the present experiments,
the physiological activity of mitochondria has been assessed using
m-sensitive fluorescent dyes. These dyes
exhibit exceptional sensitivity to small changes in
m (Ward et al., 2000) and offer the
opportunity to study subtle activities inherent in mitochondria.
Several laboratories have reported fluctuations in
m in isolated mitochondria (Ichas et al.,
1997; Huser and Blatter, 1999), cardiomyocytes (Duchen et al., 1998),
neuroblastoma (Loew et al., 1994; Fall and Bennett, 1999), vascular
endothelial (Huser and Blatter, 1999), and pancreatic B-cells
(Krippeit-Drews et al., 2000). We report here that spontaneous,
low-amplitude changes in the m occur in
neuronal mitochondria. The widespread occurrence of these spontaneous
fluctuations have prompted us to hypothesize that mitochondria exhibit
partial, transient depolarizations that represent an inherent
physiological function of mitochondria thus far undescribed in neurons.
A significant role for the functional state of mitochondria in these
fluctuations in m is discussed.
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MATERIALS AND METHODS |
Cell culture
All procedures were in strict accordance with the NIH
Guide for the Care and Use of Laboratory Animals and were
approved by the Institutional Animal Care and Use Committee of the
University of Pittsburgh. Primary forebrain neurons were prepared as
described previously (White and Reynolds, 1995). Briefly, forebrains
from embryonic day 17 Sprague Dawley rats were removed and dissociated. Cells were plated on poly-D-lysine-coated 31 mm
glass coverslips at a density of 450,000 per milliliter (1.5 ml/coverslip) and inverted after 24 hr to decrease glial growth.
Experiments were performed when cells were 12-14 d in culture.
Solutions
Coverslips were perfused with HBSS containing (in
mM): 137 NaCl, 5 KCl, 10 NaHCO3, 20 HEPES, 5.5 glucose, 0.6 KH2PO4, 0.6 Na2HPO4, 1.4 CaCl2, and 0.9 MgSO4, pH
adjusted to 7.4 with NaOH. (+)-5-Methyl-10,11-dihydro-5H-dibenzo
[a,d] cyclohepten-5,10-imine maleate (MK-801) (10 mM stock in dH2O) was purchased from
Research Biochemicals (Natick, MA), cyclosporin A (20 mM
stock in methanol) from Calbiochem (San Diego, CA), kainic acid (10 mM stock in dH2O), oligomycin (10 mM stock in ethanol), and
p-(trifluoromethoxy)phenylhydrazone (FCCP) (750 µM in methanol) from Sigma (St. Louis, MO), and
tetrodotoxin (TTX) (200 mM stock in
dH2O) from Alomone Labs (Jerusalem, Israel). Tetramethylrhodamine methyl ester (TMRM) and
5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolocarbocyanine iodide (JC-1) were purchased from Molecular Probes (Eugene, OR).
Fluorescence imaging
Coverslips were mounted on a BX50WI Olympus Optical (Tokyo,
Japan) light microscope fitted with an Olympus Optical LUMPlanFI 60× water immersion quartz objective. All recordings were made at room
temperature while cells were perfused with 10 ml/min HBSS (alone or
containing various drugs, as described below). Imaging was performed
using a 75 W xenon lamp-based monochromator light source (T.I.L.L.
Photonics GmbH, Martinsried, Germany), and light was detected using a
CCD camera (Orca; Hamamatsu, Shizouka, Japan). Data acquisition was
controlled using Simple PCI software (Compix, Cranberry, PA). For JC-1
(see below), cells were illuminated with a 485 ± 12 nm light
(incident light is attenuated with neutral density filters; Omega
Optical, Brattleboro, VT), and emitted fluorescence was passed through
500 nm long-pass dichroic mirror. The aggregate signal was obtained
using a 605/55 nm filter, and the monomer signal was collected using a
535/25 nm filter. For TMRM (see below), cells were illuminated with a
550 ± 12 nm light, emitted fluorescence was passed through a 570 nm long-pass dichroic mirror, and the single emission signal was
obtained using a 605/55 nm filter. An image was collected every 5 sec
for the duration of the 10 min experiment.
JC-1. Coverslips were incubated for 20 min at 37°C with a
3 µM JC-1 and then rinsed in HBSS for 15 min at
room temperature. Coverslips were mounted on the microscope and
perfused with HBSS. A region of cell processes (adjacent to healthy
cell bodies) was chosen, and a differential interference contrast image
of this field was digitally captured. Mitochondria within neuronal
processes, but not cell bodies, were used in these analyses because the
dimensions of these processes are such that the movement of
mitochondria is constrained to the x-y plane (no
z-plane depth) and mitochondrial movement is readily observable.
The length of time cells were illuminated was minimized and kept
constant across coverslips. The coverslip was briefly illuminated with
485 nm light, an image of the aggregate signal was captured, and the
monomer signal was focused. The coverslip was then left unilluminated
for 3 min while the dye reequilibrated (after light exposure). For
post-treated coverslips, basal fluorescence was recorded for 4 min,
followed by a 5 min drug exposure and a 1 min recovery. Data were
collected from neurons from at least four separate culture dates
(except for BAPTA experiments, in which two culture dates were tested).
TMRM. Optimal conditions for TMRM were observed when cells
were loaded for 30 min with 200 nM TMRM in HBSS
and perfused with 20 nM TMRM during the
experiment. At this concentration of TMRM, the dye that
accumulates in the mitochondria becomes quenched and a depolarization
leads to an increase in fluorescence (Ward et al., 2000). The TMRM
experiments were identical to those with JC-1, except that cells were
not exposed to light before the initiation of the experiment because
TMRM does not appear to reequilibrate after light exposure as JC-1 does.
Data analysis
A 640 × 512 pixel field of neuronal processes was imaged,
and a "mask" that identified regions of interest (ROIs) that
correlated with the expected number, appearance, and distribution of
mitochondria within these neuronal processes was generated. The
mask was made using a single JC-1 aggregate image taken before
the initiation of the monomer imaging or from a stacking of TMRM
images. The mask isolated individual spots of fluorescence that had a
fluorescence intensity indicative of physiological
m and were more than eight contiguous
pixels. Cell bodies present in the imaged field were excluded from the
mask to prevent analysis of mitochondrial clusters often observed
within these regions. Using the fluorescence images from these dyes,
~1000 ROIs per field were detected. The ROIs generated from the
punctate JC-1 aggregate signal were transferred onto the diffuse JC-1
monomer signal, and the fluorescence intensity within each individual
ROI was analyzed over time. TMRM, unlike JC-1, is a single wavelength
dye that has nonfluorescent, quenching aggregates. Thus, to locate
mitochondria, fluorescent images collected during the 10 min experiment
were stacked, and a mask was generated based on the brightest spots of
fluorescence. ROIs were further qualified based on size, as they were
with JC-1 (more than eight pixels, not within cell bodies).
The differences in the techniques used to identify mitochondria with
JC-1 and TMRM were useful in determining whether mitochondrial motility
was a significant factor in the assessment of
m fluctuations. With JC-1, mitochondrial
location was identified only at the onset of the experiment, a
percentage of mitochondria moved during the experiment (for review, see
Overly et al., 1996), and a decrease in the number of
m fluctuations was observed. This decrease was at least partially attributable to mitochondria moving out of the defined ROI. Using an image-stacking procedure to identify mitochondria in TMRM-loaded cells, the number of mitochondria was
overestimated because the same mitochondrion could be identified at
more than one location. With TMRM, we corrected for mitochondrial motility (there was no decrease in the number of fluctuations) but, in
turn, underestimated the number of fluctuations occurring per 1000 ROIs. By using both dyes, we were thus able to determine the overall
pattern of spontaneous m fluctuations.
The fluorescence intensity data from each ROI were analyzed
separately, allowing us to determine m in
individual mitochondria within cultured neuronal processes and measure
changes in that m over the course of 10 min. Several mathematical criteria were set to detect the spontaneous
fluctuations in mitochondrial fluorescence intensity. The first
criterion was set to determine whether the changes in the raw
fluorescence values exceeded basal fluctuations. Preliminary data
suggested that inherent variability within the system accounted for
fluorescent fluctuations of 4 units or less. Therefore, the
first criterion was that changes in raw fluorescence between two
sequential images (taken 5 sec apart) were >4 fluorescent units. The
second criterion for detection of a spontaneous
m fluctuation was that the slope of the
fluorescence change had to be >0.3 fluorescent units per second for
both the rise and fall of the fluctuation. The slope was determined
using a moving average of three sequential images. This criterion was
set to distinguish fluctuations from changes in basal fluorescence
attributable to photo-oxidation, focus drifts, debris in the field
temporarily impeding fluorescence detection, or other spurious changes
in signal. Using these two criteria, preliminary analysis showed substantial correspondence between the observation of changes in
fluorescence (on the computer screen) and the statistical detection of
a spontaneous m fluctuation.
The number of a spontaneous m fluctuation
detected in a field of neuronal processes was determined and normalized
as the number of spontaneous m fluctuations
occurring per minute per 1000 ROIs. Data were graphed as either the
percentage of fluctuations in a drug-treated field versus an untreated
control field (for pretreatment experiments) or as a ratio of the
number of spontaneous m fluctuations per
minute per 1000 ROIs before versus after drug treatment (for
post-treatment experiments). Presentation of a ratio was chosen
because, as a result of mitochondrial motility, all JC-1-loaded
neuronal fields analyzed showed a decreasing number of fluctuations
with time. For the post-treatment experiments, the number of
fluctuations occurring per minute per 1000 ROIs was counted, and the
average of minutes 2-4 (baseline) and minutes 6-8 (treatment) were
calculated. This corrected for any instability in the baseline at the
onset of the experiment and allowed treatment effects to stabilize for
2 min before analysis. Data from controls and individual drug
treatments were averaged across coverslip and culture date and
statistically compared using a t test or ANOVA.
To distinguish between long-term, global mitochondrial depolarizations
and the spontaneous, small-amplitude fluctuations in m, the average fluorescence intensity in
all mitochondria in a field of neurons was calculated. This enabled us
to determine whether there was an association between average
m and the occurrence of spontaneous
fluctuations. Average fluorescence intensity (in arbitrary units) is
presented for all pharmacological experiments.
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RESULTS |
Under physiological conditions, mitochondria located within
cultured neuronal processes exhibit a great deal of spontaneous activity, including repetitive, small-amplitude depolarizations. Our
previous measurements of m were limited in
both spatial and temporal resolution (White and Reynolds, 1996; Scanlon
and Reynolds, 1998), and thus these phenomena were undetected. Using the wide-field imaging system described here, we captured images at 0.2 Hz with resolution adequate to monitor signals from neuronal processes.
Replaying these images at a rate of 6 Hz revealed some unappreciated
dynamics in the dye signal. Mitochondria traverse several tens of
micrometers and, more surprisingly, exhibit extensive spontaneous
alterations in dye signal consistent with depolarization of
m over the period of the experiment. With
real-time imaging (0.2 Hz), fluctuations in
m could occasionally be observed; however,
on playback (6 Hz), this phenomenon was observed throughout the field
of neurons during the entirety of the experiment.
Figure 1 illustrates the
spontaneous m fluctuations in JC-1-loaded
(Fig. 1A) and TMRM-loaded (Fig. 1B)
neurons. This figure shows small regions of fluorescence
(arrowheads) within untreated neuronal processes that appear
to be spontaneously increasing and decreasing in intensity. We were
unable to determine whether there was a propagation of this signal down
a process, although this did not appear to be the case. The
fluctuations in fluorescence within individual mitochondria appear
independent, and no wave-like activity was noted. Movies of JC-1- and
TMRM-loaded neurons illustrating this phenomenon are included in the
supplemental data section located in the on-line version of this
article. For details of these movies, see the legend for Figure
1.
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Figure 1.
Representative fluorescent images of
m fluctuations. A, JC-1-loaded
neuronal processes. B, TMRM-loaded processes. These
panels show spontaneous changes in fluorescent intensity
occurring in small regions of neuronal processes. Images show a
200 × 200 pixel field. Images were taken 30 sec apart.
Arrowheads identify examples of regions of fluorescence
that correspond to the expected size and shape of neuronal mitochondria
and show readily observed fluctuations in intensity. Increases in
fluorescence imply depolarization. TMRM-loaded cells have
lower basal fluorescence because light levels were kept low to avoid
light-induced increase in fluorescence. Scale bar, 10 µM.
Movie files corresponding to JC-1-loaded (A) and
TMRM-loaded (B) neurons have been included in the
supplemental data section located in the on-line version of this
article. Movie images were taken 5 sec apart, and time is shown in
minutes and seconds. Both spontaneous fluctuations and dye intensity
and mitochondrial motility are evident.
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Measurement of spontaneous m fluctuations
To provide a quantitative analysis of this phenomenon, we
developed a technique for identifying individual mitochondria within a
field of neuronal cell bodies and processes (Fig.
2). The majority of the data presented
were collected using the potentiometric dye JC-1, although
qualitatively similar results were obtained with TMRM. JC-1 was
preferred for these experiments because its aggregates accumulate
within the mitochondria based on m and are
fluorescent. This allowed us to use the aggregate fluorescent signal to
locate individual mitochondria and the monomer signal to detect changes
in m. A loss of
m results in an increase in JC-1 monomer
fluorescence in the regions in and around the mitochondria.
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Figure 2.
Imaging technique used to measure spontaneous
changes in mitochondrial membrane potential using JC-1.
A, Bright-field image, using differential interference
contrast. The arrow shows a healthy neuronal cell body
with processes within a field of neurites. B-E
represent fluorescent images from this same field. B,
The JC-1 aggregate image. Note the punctate nature of the label within
the processes. C, The mask (red) created
to discern ROIs thought to correspond to mitochondria. The
bright fluorescent spots observed with the JC-1 aggregate image (same
as B, now white) are detected using image
analysis software based on size criteria. Regions uniformly labeled
(e.g., cell bodies) are excluded from the mask based on size.
D, The JC-1 monomer image. Note the diffuse nature of
the label within the processes. E, The "overlay."
The mask generated in C (red) is overlaid
onto the JC-1 monomer image from D
(green). Scale bar, 20 µM.
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A differential interference contrast image was taken (Fig.
2A), followed by a single fluorescent image of the
JC-1 aggregate signal (Fig. 2B). We then isolated
each individual fluorescent area and identified it as an ROI depicting
what is likely to be a single mitochondrion (Fig. 2C).
Because of the thickness of the cell bodies and the wide-field
microscopy used, it was difficult to distinguish individual
mitochondria within the soma. This is evidenced by the large regions of
aggregate fluorescence present in the cell body (arrowhead),
most likely resulting from numerous mitochondria overlapping in the
z-axis. These large regions of fluorescence, most typically
seen in cell bodies, were excluded from analysis based on size. The
mask containing the ROIs (Fig. 2C) was overlaid onto the
JC-1 monomer signal (Fig. 2D), and the fluorescence
intensity from each individual ROI was recorded (Fig. 2E). This technique allowed us to measure the
fluorescence intensity of ~1000 ROIs per field from images collected
every 5 sec for 10 min.
A graphical illustration of the spontaneous
m fluctuations is presented in Figure
3, A and B. There
appears to be no regularity in amplitude or frequency of the
spontaneous depolarizations. Note that the individual mitochondria in
each of these graphs (traces A-D) exhibit
numerous spontaneous depolarizations and repolarizations over the
course of the experiments, with the overall basal fluorescence
intensity remaining relatively stable. However, not all mitochondria
exhibit spontaneous fluctuations (trace E on both graphs). Over 35,000 mitochondria from untreated cells were
assessed for spontaneous m fluctuations in
these experiments. The percentage of mitochondria that exhibit at least
one m fluctuation was calculated from
~15,600 mitochondria in JC-1-loaded control fields, and 55% of these
mitochondria were inherently active.
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Figure 3.
Quantification of m
fluctuations within individual mitochondria. A, JC-1
monomer traces. B, TMRM fluorescence traces. Each
trace represents in a single ROI corresponding to an
individual mitochondrion. An increase in fluorescence is associated
with depolarization. Note that the basal fluorescence in both graphs
remains stable and that fluctuations are not all of the same magnitude
or of regular frequency. Approximately 1000 such traces were collected
per field. The y-axis represents arbitrary fluorescence
units, with each tick mark equaling 10 units.
Traces have been offset from one another for display
purposes. A, Trace A shows a
mitochondrion that fluctuates repeatedly followed by a loss of
fluorescence, most likely indicating the movement of the mitochondrion
out of the ROI. Trace B shows a
mitochondrion that fluctuates especially as the experiment continues.
Trace C shows large fluctuations and then a
slow increase in fluorescence suggesting a gradual depolarization.
Trace D initially exhibits fluctuations, but
by 7 min activity ceases. B, Traces
A and B show
mitochondria that fluctuate repeatedly throughout the experiment.
Trace C shows a mitochondrion that
fluctuates especially as the experiment continues. Trace
D initially has fluctuations, but by 5 min activity
ceases. Trace E from both figures shows
background noise in the system and reflects mitochondria that do not
exhibit spontaneous fluctuations. C, Average number of
spontaneous m fluctuations in JC-1- or
TMRM-loaded neurons from traces such as those presented
in A and B. Criteria for a spontaneous
fluctuation in m were set a priori: fluorescence
changes >4 fluorescent units and a slope >0.3 fluorescent units per
second. The number of m fluctuations that occurred
per minute per 1000 ROIs is presented. A decrease in the number of
m fluctuations over time was observed in JC-1-loaded,
but not in TMRM-loaded, neurons (see Results). Data are presented as
means ± SEM. The number of fields (n)
imaged (~1000 ROIs per field) is presented in the key.
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From Figure 3, it is evident that the fluctuations appear different in
JC-1- and TMRM-loaded mitochondria. The m
fluctuations appear slower in JC-1-loaded neurons, probably because of
the differences in the properties of the dyes. TMRM equilibrates faster across membranes, and thus the fluctuations appear more rapid. This
difference in the rate of reequilibration of these dyes thus results in
the same phenomenon having a somewhat different appearance, depending
on the dye used (Nicholls and Ward, 2000).
JC-1- and TMRM-loaded mitochondria can also be distinguished by the
fluctuation frequency, which appears to decrease over time with JC-1
but remain stable or increase with TMRM (Fig. 3C). This may
be explained by the rapid distribution of TMRM across membranes, which
necessitated the addition of TMRM into the perfusate to maintain the
level of fluorescence within the neurons (Fig. 3B, note the
stability of the basal fluorescence within the ROIs). In contrast, JC-1
was not included in the perfusate, making it likely that there is only
a partial reaggregation of the dye during repolarization, with some dye
diffusing out of the cell. This could lead to a decrease in JC-1
aggregate fluorescence, which could imply less monomer release until
eventually the magnitude of the increase in monomer fluorescence would
fall below the criterion needed to indicate a fluctuation. This
explanation however seems unlikely because, when JC-1 aggregate
fluorescence in neurons was decreased by high-light exposure,
depolarization could still be observed as a rise in monomer
fluorescence (data not shown).
Another factor that may impact the number of fluctuations is that a
percentage of mitochondria are known to be motile within neurons
(Overly et al., 1996). In fact, we note that a fraction of the
mitochondria move throughout the processes of our cultured neurons,
regardless of which dye we used. With JC-1, mitochondria were located
and ROIs identified only at the onset of the experiment, using the JC-1
aggregate signal. Thus, over time, the regions being analyzed may no
longer correlate with the regions exhibiting the changes in
fluorescence (i.e., a mitochondrion may move out of the defined ROI).
TMRM-loaded mitochondria were located using image stacking; thus, when
a mitochondrion moves, each location may be associated with an
independent ROI. This would suggest that TMRM overestimates the number
of ROIs and thus underestimates the number of fluctuations per 1000 ROIs. These technical differences can explain why TMRM-loaded
mitochondria appear more active than JC-1-loaded mitochondria, a fact
that is not reflected in the average number of fluctuations reported in
Figure 3C. In addition, they suggest that the percentage of
mitochondria that move is stable because the decrease in fluctuations
observed with JC-1 is very consistent. It is important to note,
however, that regardless of the cause, we have corrected for this loss
of fluctuation detection in subsequent pharmacological experiments by
analyzing only the ratio of spontaneous m
fluctuations at the beginning versus end of the experiment.
It was important to rule out nonphysiological influences on the
fluctuations in m. Fluorescence imaging of
organelles as small as mitochondria requires careful maintenance of the
correct focal plane. Drifts in focus could lead to the appearance
and/or disappearance of m fluctuations. To
minimize inclusion of focus changes in our quantitation of
m fluctuations, we used a moving average
paradigm to smooth out brief irregularities in whole-field fluorescence
(see Materials and Methods). However, because the cells were also being
perfused throughout the experiment, it was possible that smaller
regions of the field were drifting in and out of focus, thus leading to
the graphical appearance of fluctuations. To show that the
m fluctuations were not attributable to
focus drift, we identified three adjacent mitochondria located within a
single neuronal process and analyzed them for spontaneous
m fluctuations (Fig.
4). If the m
fluctuations were the result of drifts in focus, then mitochondria in
close proximity to one another should all exhibit simultaneous
fluctuations. This was not observed. Three neighboring ROIs were
identified using the JC-1 aggregate signal (Fig. 4A,
red), and the fluorescence intensity of JC-1 monomer signal
(Fig. 4A, green) within each ROI was
determined. As the monomer signal increases (indicating
depolarization), the ROIs appear increasingly yellow (Fig.
4A) and an increase in fluorescence intensity is
observed graphically (Fig. 4B). The fluctuations in
fluorescence intensity observed qualitatively and quantitatively appear
to correspond, with no evidence of changes in focus in any of the ROIs.
Moreover, two mitochondria demonstrate fluctuations (ROIs 1 and 3),
whereas the ROI in the middle does not (ROI 2). This strongly argues
against a drift in focus or movement of the process as the underlying
mechanism of this phenomenon.
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Figure 4.
m fluctuations are not
attributable to focus drift. Three individual ROIs located within a
single neuronal process analyzed for spontaneous m
fluctuations. A, In this series of images, the JC-1
aggregate signal (red, depicting individual ROIs) has
been overlaid on the corresponding JC-1 monomer
(green) images (as shown in Fig.
2E). The first image in this series identifies
three mitochondria at the onset of imaging (arrows). The
numbers associated with each mitochondrion correspond to
the traces in B. In images
a-f, a static aggregate (red) image has
been used; thus, changes in the ratio of green to
red represent changes in JC-1 monomer fluorescence.
(i.e., when mitochondria appear yellow, monomer signal
has increased, suggesting depolarization). Note that fluctuations in
JC-1 monomer fluorescence intensity can be observed qualitatively over
time. B, Fluorescence intensity of the JC-1 monomer
signal within each of the three ROIs identified in A was
quantified (arrowheads a-f show the time
points when the images in A were taken). Note that
traces 1 and 3 show significant
fluctuations in fluorescence intensity, but trace 2 does
not. This illustrates that changes in fluorescence intensity are not
simply attributable to a drift in focus or movement of the
process. If this were the case, then all three mitochondria within this
process should exhibit similar m fluctuations.
C, A differential interference contrast image showing
the neuronal process from which the fluorescent images were taken. The
arrows correspond to the ROIs analyzed. Scale bar, 5 µM.
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Illumination of mitochondria loaded with cationic fluorescence dyes has
been reported to lead to the generation of reactive oxygen species
(ROS) and thus mitochondrial damage (Bunting, 1992). Photo damage to
the mitochondria could therefore be a potential cause of the observed
fluctuations in m. Typically, during
imaging, neurons were exposed to attenuated fluorescent light (5%
transmitted light) for ~0.1 sec every 5 sec over the course of a 10 min experiment. To determine the relative contribution of light, we
took JC-1-loaded neurons and exposed a portion of the field to a brief,
intense light. For our high-light conditions, we maintained the
exposure time of 0.1 sec (per 5 sec) but increased transmitted light to 100%. In these experiments, JC-1-loaded neuronal processes were imaged
under standard conditions for 50 frames (one frame per 5 sec). The
microscope aperture size was then decreased so that light exposure was
limited to the center of the neuronal field and the processes in this
central region were illuminated, for 20 frames (one frame per 5 sec),
with a 20-fold higher light intensity. Light conditions were then
returned to normal. We separated the imaged field into the exposed
center region (illuminated under high-light conditions) and the
peripheral, control region (unilluminated for the same 20 frames). The
ratio of fluctuations occurring before versus after intense light
exposure were compared across the control and exposed regions. The
region exposed to high-light conditions showed significantly fewer
spontaneous m fluctuations (Fig. 5A), without a concurrent
change in average fluorescence intensity (Fig. 5B). Thus,
photo damage cannot account for the observation of these phenomena; in
fact, these data suggest that light exposure can lead to the loss of
m fluctuations, potentially through a
photo-induced oxidant burden.
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Figure 5.
m fluctuations in JC-1-loaded
neurons are not caused by light. Untreated neurons were loaded with
JC-1 and imaged under our standard light conditions for 50 frames
(neutral density attenuating transmitted light to 5%). The aperture on
the microscope was then closed to allow light to hit only the center
portion of the field, and the cells were imaged for 20 frames. Under
this condition, the middle portion of the coverslip was exposed to
intense light (100% transmitted light; EXPOSED AREA),
whereas the periphery of the coverslip was left unexposed
(CONTROL AREA). The imaging conditions were then
returned to the standard conditions from the beginning of the
experiment for 50 more frames. A, The number of
fluctuations in m per minute per 1000 ROIs was
determined in the control and exposed regions, and the ratio of
fluctuations occurring before versus after intense light exposure are
presented (note that the control area received no light during this
period). Data are presented as means ± SEM, and the
numbers above each bar equal the number
of fields imaged. The region exposed to high-light conditions showed
significantly fewer spontaneous m fluctuations
(paired t test; t = 3.77; df = 5; p < 0.05). B, Average
fluorescence intensity of all mitochondria imaged, with representative
error bars indicating SEM. Basal fluorescence did not change after
light exposure.
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Pharmacological analysis of spontaneous
m fluctuations
Having established methods to measure spontaneous fluctuations in
fluorescence intensity in JC-1-loaded mitochondria in untreated neurons, we explored mechanisms underlying this phenomenon. We first
tested the possibility that the m
fluctuations were the result of synaptic activity and thus were
reflecting plasma membrane potential changes or calcium influx. Neurons
were treated with TTX (200 nM), a sodium channel blocker,
to inhibit spontaneous synaptic activity. We also tested an NMDA
receptor antagonist, MK-801 (10 µM), to block a major
route of calcium entry. JC-1-loaded neurons were perfused with HBSS for
4 min to establish basal activity. Neurons were then exposed to the
inhibitors for 5 min. The ratio of fluctuations occurring before versus
those occurring after drug treatment was graphed to correct for the
decrease in the number of fluctuations that occurs even in untreated
control mitochondria (Fig. 3C). There was no
difference in the ratio of fluctuations between control mitochondria
and those treated with TTX or MK-801 (Fig.
6A). These treatments
did not alter the average fluorescence intensity of the JC-1 monomer
signal within the mitochondria (Fig. 6B). This
suggests that drugs that inhibit plasma membrane activities do not
inhibit spontaneous fluctuations or change the resting level of
m.
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Figure 6.
m fluctuations are not
attributable to synaptic activity. Untreated neurons were loaded with
JC-1 and imaged for 4 min before the addition of drugs
(arrow). TTX (200 nM) (a
Na+ channel blocker) or 10 µM MK-801
(NMDA antagonist) were perfused over the cells for 5 min, and
fluorescence intensity and number of fluctuations per minute per 1000 ROIs were calculated. A, The ratio of fluctuations
occurring during the baseline to those occurring after drug treatment.
This corrected for the consistent decrease in fluctuations observed in
untreated mitochondria (see Fig. 3). Data are presented as means ± SEM, and the numbers above each bar
equal the number of fields imaged (~1000 ROIs per field). Neither
treatment had a significant impact on m fluctuations
(TTX, t = 1.03, df = 14, p > 0.05; MK-801, t = 1.98, df = 16, p > 0.05). B, Average fluorescence
intensity of all mitochondria imaged, with representative error bars
indicating SEM. Neither treatment altered basal fluorescence.
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There are several additional routes of calcium entry beyond NMDA
receptor activation. Mitochondria are intricately involved in shaping
calcium dynamics (Duchen, 1999), and calcium has also been suggested to
regulate m fluctuation in other cells (Loew et al., 1994; Ichas et al., 1997; Duchen, 1999; Krippeit-Drews et al.,
2000). BAPTA, an intracellular calcium chelator, interrupts calcium-mediated events that result from calcium influx and release from intracellular stores. Because of the close apposition of mitochondria to endoplasmic reticulum, release of calcium from the
reticular stores can alter mitochondrial function by creating microdomains of high calcium (Rizzuto et al., 1993, 1998; Hajnoczky et
al., 1995). Thus, any involvement of calcium-mediated activity in the
observed spontaneous mitochondrial depolarizations should become
evident with BAPTA treatment. Neurons were preincubated with BAPTA-AM
(50 µM) showed a dramatic decrease in the number of
spontaneous fluctuations compared with controls (Fig.
7A). However, this
BAPTA-induced inhibition was associated with a dramatic rise in
fluorescence intensity (Fig. 7B). This implies that
BAPTA-AM, at this concentration, leads to mitochondrial depolarization. The mechanism by which this occurs is unclear and cannot necessarily be
attributed to calcium chelation. Respiratory complex inhibitors and the
uncoupler FCCP depolarized m and decreased
the number of spontaneous fluctuations (data not shown). This raises
the possibility that the effects of BAPTA are attributable to general mitochondrial depolarization, which occludes the smaller fluctuations rather than a result of modifying intrinsic calcium changes.
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Figure 7.
An intracellular calcium chelator alters
m fluctuations and basal fluorescence. Neurons loaded
with JC-1 were treated with 50 µM BAPTA-AM for 15 min
before imaging. The fluorescence intensity and number of fluctuations
per minute per 1000 ROIs were calculated. A, The
percentage of fluctuations occurring in BAPTA-pretreated mitochondria
to untreated mitochondria. Data are presented as means ± SEM.
Numbers above each bar equal the number
of fields imaged (~1000 ROIs per field). BAPTA decreased
m fluctuations (t = 5.17; df = 7; *p < 0.05; but see Discussion).
B, Average fluorescence intensity of all mitochondria
imaged, with representative error bars indicating SEM. BAPTA increased
the average fluorescence intensity. (Note that the scale is twice that
of Figs. 8-10).
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Activity of the respiratory chain complexes leads to the extrusion of
protons from the mitochondrial matrix, whereas synthesis of ATP through
the F1FO ATPase
(ATP synthase) is driven by proton reentry. Thus, whereas inhibition at
the respiratory chain complexes may lead to a slow loss of
m, inhibition at the ATP synthase should
lead to a subtle hyperpolarization in healthy, ATP-generating mitochondria and a depolarization in damaged, ATP-consuming
mitochondria (Scott and Nicholls, 1980; Nicholls and Ward, 2000).
Neurons pretreated with oligomycin (15 min) showed a significant
decrease in the number of spontaneous m
fluctuations compared with untreated control mitochondria (Fig.
8A) but no significant
change in initial basal fluorescence (Fig. 8B).
Because it is unlikely that all mitochondria within a field of neurons
are of equivalent membrane potential and ATP synthase activity and
~1000 mitochondria were averaged per field, small changes in overall
fluorescence intensity were not detected.
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Figure 8.
m fluctuations are decreased by
pretreatment with an ATP synthase inhibitor. Neurons loaded with JC-1
were treated with 10 µM oligomycin (OLIGO)
for 15 min before imaging to ensure substantial inhibition of the
synthase. Fluorescence intensity and number of fluctuations per minute
per 1000 ROIs were calculated. A, Percentage of
fluctuations in oligomycin-pretreated to untreated mitochondria
(means ± SEM). Oligomycin significantly decreased
m fluctuations (t = 3.03;
df = 14; **p < 0.01).
Numbers above bars equals the number of
fields imaged (~1000 ROIs per field). B, Average
fluorescence intensity of all mitochondria, with representative error
bars indicating SEM. Oligomycin had no effect.
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The PTP is thought to regulate matrix
Ca2+, pH, m,
and volume (Susin et al., 1998). Its opening is triggered by a wide
variety of treatments, including increased
Ca2+ (Zoratti and Szabo, 1995), and is
inhibited by cyclosporin A (Crompton et al., 1988). Two open states for
this pore have been proposed, with the low-conductance state causing
transient openings that aid in stabilizing
m and volume (Ichas et al., 1997) and the
high-conductance state involved in irreversible openings, complete
dissipation of the m, extensive swelling,
and cell death (Petit et al., 1996; Susin et al., 1997). Transient
depolarizations in m have been reported to
coincide with activation of the low-conductance state of the PTP in
isolated mitochondria, which could represent a physiological process
necessary for regulating m (Ichas et al.,
1997) and limiting the generation of ROS in hyperpolarized mitochondria
(Skulachev, 1996). We tested whether cyclosporin A influenced the
frequency of the m fluctuations in our
cultured neurons. No effect was observed at any concentration tested
(Fig. 9A); however,
cyclosporin A caused a modest decrease in fluorescence intensity at
lower concentrations (Fig. 9B).
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Figure 9.
m fluctuations are not altered
by treatment with cyclosporin A. Neurons loaded with JC-1 were treated
with 0.2-20 µM cyclosporin A (CsA) for 15 min before imaging. Fluorescence intensity and number of fluctuations
per minute per 1000 ROIs were calculated. A, Percentage
of fluctuations in cyclosporin A-pretreated mitochondria to untreated
mitochondria. Cyclosporin A did not significantly alter
m fluctuations
(F(3,34) = 1.27; p > 0.05). Data are presented as means ± SEM, and the
numbers above bars equal the number of
fields imaged (~1000 ROIs per field). B, Average
fluorescence intensity, with representative error bars indicating SEM.
Cyclosporin A did not substantially alter the average fluorescence
intensity.
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Taxing neurons by increasing synaptic activity may lead to an increase
in mitochondrial activity. Stimulation of neurons by high
concentrations of glutamate causes large calcium fluxes that directly
influence mitochondrial function (White and Reynolds, 1995) and have
been proposed to increase ATP demand by the plasma membrane ATPases.
Although inhibition of basal synaptic activity with a sodium channel
blocker or a glutamate receptor antagonist did not alter the frequency
of m fluctuations (Fig.
6A), we tested whether increasing synaptic activity,
and presumably increasing the demand placed on the mitochondria, would
alter the frequency of fluctuations. Treatment with glutamate in the
presence of extracellular calcium leads to an increase in fluorescence
intensity in a similar manner to that observed with BAPTA-AM. In fact,
stimulation with glutamate is known to cause mitochondrial calcium
influx and depolarization (Ankarcrona et al., 1995; White and Reynolds,
1996). We attempted to circumvent the problem of the superimposition of
the catastrophic depolarization on the smaller spontaneous changes
using two approaches. First, we treated neurons with kainic acid, an
agonist of the AMPA-kainate subtype of glutamate receptors. Activation
of glutamate receptors with kainic acid results in increased
intracellular sodium and calcium but does not lead to mitochondrial
depolarization (Courtney et al., 1995, Hoyt et al., 1998). Accumulation
of calcium within mitochondria is also significantly lower using kainic
acid than it is with NMDA (Budd and Nicholls, 1996; Stout et al.,
1998). Therefore, neurons were treated with 100 µM kainic acid for 5 min, in either the
presence or absence of calcium, but no evidence of altered
m fluctuation frequency (Fig.
10A) or overall
fluorescence intensity (Fig. 10B) was observed.
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Figure 10.
m fluctuations are not altered
by treatment with kainic acid. Untreated neurons were loaded with JC-1
and imaged for 4 min before the addition of drug
(arrow). Kainic acid (100 µM) was perfused
over the cells in a buffer containing 1.4 mM
(+Ca2+) or 0 mM
(no Ca2+) calcium for 5 min, and
fluorescence intensity and number of fluctuations per minute per 1000 ROIs were calculated. A, The ratio of fluctuations
occurring during baseline to those occurring after drug treatment. Data
are presented as means ± SEM, and the numbers
above each bar equal the number of fields imaged
(~1000 ROIs per field). Neither treatment had a significant impact on
m fluctuations (with Ca2+,
t = 1.90, df = 9, p > 0.05; without Ca2+, t = 0.48, df = 9, p > 0.05). B, Average
fluorescence intensity of all mitochondria imaged, with representative
error bars indicating SEM. Neither treatment altered basal
fluorescence.
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We then attempted to increase synaptic activity and mitochondrial
demand without flooding mitochondria with calcium and depolarizing them
by treating neurons with glutamate (in the presence of its coagonist
glycine) in buffer that is nominally calcium free. This should lead to
the activation of the glutamate receptors without the concurrent
increase in intracellular and intramitochondrial calcium
concentrations. Receptor activation should, however, lead to an influx
of sodium and thus manipulate mitochondrial ion concentrations through
sodium-related pathways. JC-1-loaded neurons were treated with 100 µM glutamate and 10 µM glycine in
calcium-free buffer for 5 min. This treatment led to a significant
decrease in m fluctuations (Fig.
11A) without a
concomitant rise in fluorescence intensity (Fig.
11B). The differences between glutamate and kainate on m fluctuations are not surprising in
light of the observation that their ability to activate glutamate
receptors is not equivalent qualitatively or quantitatively (Hoyt et
al., 1998; Sattler et al., 1998; Brocard et al., 2001).
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Figure 11.
m fluctuations are decreased
after treatment with glutamate in a calcium-free buffer. Untreated
neurons were loaded with JC-1 and imaged for 4 min before the addition
of drug (arrow). Glutamate (100 µM,
with 10 µM glycine) was perfused over the cells in
a Ca2+-free buffer for 5 min. Fluorescence intensity
and number of fluctuations per minute per 1000 ROIs were calculated.
Glutamate in the presence of Ca2+ caused
mitochondrial depolarization and thus was not tested. A,
The ratio of fluctuations occurring during baseline to those occurring
after drug treatment. Data are presented as means ± SEM, and the
numbers above each bar equal the number
of fields imaged (~1000 ROIs per field). Glutamate in
Ca2+-free buffer significantly decreased the number
of spontaneous fluctuations (t = 4.04; df = 8;
**p < 0.01). B, Average
fluorescence intensity of all mitochondria imaged, with representative
error bars indicating SEM. Glutamate (no
Ca2+) treatment did not alter basal
fluorescence.
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DISCUSSION |
In this study, we report that mitochondria in neuronal cultures
display small, spontaneous fluctuations in
m and that these fluctuations can be
dramatically decreased, without a concurrent change in basal
fluorescence, by treatments that alter mitochondrial activity. Although
there has been a great deal of recent interest in large-scale changes
in mitochondrial function associated with neuronal injury, the present
findings reveal a previously unappreciated property of mitochondria in
resting, uninjured neurons.
The m is a key marker of mitochondrial
function, generated by the pumping of protons across the inner
mitochondrial membrane in association with electron transport. In turn,
m drives many key mitochondrial functions,
including ATP synthesis, calcium accumulation, and maintenance of ion
gradients that permit the influx of substrates and egress of metabolic
products. Clearly, m has a number of
important functions, and thus a variety of activities could account for
the spontaneous changes in m reported here.
Our observations of m fluctuations could
reflect inherent changes in mitochondrial ion transport, ATP
production-consumption, respiration, or volume, all of which are
essential for proper mitochondrial function. We believe that
spontaneous m fluctuations represent a
normal physiological feature of neuronal mitochondria such that the
presence or absence of these fluctuations may be useful as a novel
marker for mitochondrial activity.
The pharmacological approach of using tetrodotoxin or MK-801 clearly
excludes alterations in plasma membrane potential and ion fluxes as the
basis for the change in the mitochondrial dye signal. However, the
mitochondrial mechanism that causes the fluctuations is less clear.
Attempts to regulate the permeability transition pore with cyclosporin
A had no effect on either m fluctuations or
fluorescence intensity, which ostensibly excludes low-amplitude transition as a basis for these changes (Ichas et al., 1997). Oligomycin, however, significantly decreased the frequency of fluctuations, which argues that the fluctuations are associated with
oxidative phosphorylation.
Oxidative phosphorylation may impact m
fluctuations by one of two mechanisms. The first mechanism would
suggest that, as mitochondria go from a resting state to active
phosphorylation (state 4 to state 3), the fluctuations reflect the
transient loss of m as proton flux
increases (Nicholls and Ferguson, 1992). In this case, the increase in
dye signal would be attributable to the disaggregation and subsequent
dissipation of dye from mitochondria. Alternatively, because state 3 mitochondria adopt a condensed configuration whereas state 4 mitochondria adopt the larger orthodox configuration (Scalettar et al.,
1991), the fluctuations would reflect changes in mitochondrial matrix
volume. In this case, the increase in dye signal attributable to the
increased volume would lead to a disaggregation of dye that is retained
within the mitochondrial matrix. Although these mechanisms are
contradictory in that the first proposes that a fluctuation is
associated with the onset of phosphorylation whereas the second
suggests that the signal should be associated with the termination of
active phosphorylation, the key feature of both mechanisms is that the fluctuations represent an on-off transition. Thus, either mechanism could explain the decrease in fluctuations observed with a
glutamate-induced sodium load. In this condition, an increase in ATP
demand presumably occurs causing the mitochondria to spend a greater
fraction of the time engaged in active phosphorylation rather than
switching on and off. One could also argue that light-induced damage
places greater demands on the mitochondria to be met by increasing ATP synthesis (or perhaps by decreasing synthetic capacity).
Predicting manipulations that increase the frequency of fluctuations is
harder. We have observed differences in the number fluctuations on a
culture-to-culture basis but have not yet established a mechanism for
these differences. We are not aware of any previous studies that have
suggested that individual mitochondria can be observed to change
between states of resting and active phosphorylation. However, the
architecture of neurons is uniquely suited to making such observations
because optically isolating individual mitochondria in neuronal
processes is straightforward (Figs. 1, 2).
Cationic dyes used to measure m (such as
JC-1 and TMRM) can lead to toxicity resulting from light-induced
singlet oxygen generation (Bunting, 1992) and inhibition of
respiration. Although light exposure was carefully controlled and
minimized to that necessary for adequate recordings, some effects of
light exposure were observed even under these low-light conditions.
With JC-1, exposure to light before recording stabilized the baseline,
and with TMRM, increasing light exposure tended to increase the overall fluorescent signal. Both of these light-induced changes in dye signal
could be indicative of a potential impact of phototoxicity in
m measurements. A more immediate concern
was that the spontaneous changes were triggered by light exposure.
Thus, if a brief, intense light augmented the frequency of
m fluctuations, it would suggest that these
fluctuations were merely reflecting photo damage. We saw, however, a
decrease in the number of fluctuations under high-light conditions,
suggesting that dye-loaded mitochondria exhibit spontaneous fluctuations in m, which were not a
consequence of illumination. These high-light conditions led to a
decrease in the JC-1 aggregate signal without a concurrent change in
the monomer signal (data not shown), similar to what is observed with
oxidant treatments, such as hydrogen peroxide (Scanlon and Reynolds,
1998; Chinopoulos et al., 1999). However, treatment with the uncoupler
FCCP at the end of the high-light experiments led to an increase in
monomer fluorescence of the same magnitude in both the exposed and
unexposed regions (data not shown). This supports the hypothesis that
JC-1 aggregate fluorescence responds to more than just changes in
m (Scanlon and Reynolds, 1998; Chinopoulos
et al., 1999) but indicates that light-induced changes in aggregate
fluorescence do not change the ability of the JC-1 monomer signal to
respond to changes in m.
Calcium may have a profound impact on mitochondrial function in general
and on membrane potential in particular. Small transient changes in
m observed in cardiomyocytes were reported
to be the result of mitochondrial calcium transport (Duchen et al., 1998; Fall and Bennett, 1999). Furthermore, calcium can induce the
generation of ROS, alter respiration (McCormack et al., 1990), and
possibly open the large, nonselective PTP (Zoratti and Szabo, 1995).
This suggests that calcium could be a key mediator of changes in
m. Inhibiting the NMDA receptor, thus
decreasing the entry accumulation of calcium (Fig.
6A), chelating extracellular calcium with EGTA (data
not shown), and inhibiting the PTP with cyclosporin A (Fig. 9) all
failed to change m fluctuations, which
argues against this possibility. Chelating intracellular calcium with BAPTA (Fig. 7A) did decrease fluctuations, but this occurred
in conjunction with a considerable mitochondrial depolarization. The
large rise in monomer fluorescence induced by BAPTA most likely occluded our ability to detect small fluctuations and was similar to
the effects of the protonophore FCCP. However, it remains unclear how
adding BAPTA influences the free calcium because, under resting conditions, the calcium concentration in both the cytoplasm and mitochondria of these neurons is low (Brocard et al., 2001).
We believe that these are the first experiments that illustrate
spontaneous changes in m in neurons.
Previous studies have suggested cyclosporin A-stimulated changes
in whole-cell TMRM signal in neuroblastoma cells (Fall and Bennett,
1999), which are obviously distinct from the single organelle signals
reported here. In cardiomyocytes, transient depolarizations in single
mitochondria have been seen (Duchen et al., 1998), but these changes
were the consequence of calcium movements. Other studies have
investigated single mitochondria but only after their isolation from
cells (Ichas et al., 1997; Huser and Blatter, 1999). Apparently none of
these studies have reported oligomycin sensitivity of the fluctuations.
Herein, we have documented a novel feature of mitochondrial physiology
in neurons. Unlike previous studies that have reported large changes in
m associated with neuronal injury, the
spontaneous changes in m reported here
appear to be a normal characteristic of mitochondrial function in
neurons and may reflect alterations in the activity of individual
mitochondria associated with the transition between rest and active
oxidative phosphorylation. We suggest that this phenomenon that may
prove to be a useful marker of mitochondrial function in neurons and
hypothesize that these fluctuations in m
reflect variations in the cellular environment associated with altered
states of respiratory control or ion-induced matrix swelling.
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FOOTNOTES |
Received March 8, 2001; revised April 25, 2001; accepted April 30, 2001.
This work was supported by United States Army Medical Research and
Materiel Command Grant DAMD-17-98-1-8627 (I.J.R.), the Scaife
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TOP
ABSTRACT
INTRODUCTION
