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INTRODUCTION |
Higher brain functions are susceptible to damage through exposure
to the prolonged hypoxia of ischemia or chronic cardiorespiratory disease (1-3). Indeed, there is a well documented increased incidence of dementias in patients who have previously suffered prolonged hypoxic
or ischemic episodes arising as a consequence of cardiovascular dysfunction such as stroke or arrhythmia (4-6). Such a clear link
between hypoxic/ischemic episodes and increased incidence of dementias
strongly suggests that lack of oxygen is a contributory factor in the
precipitation of such diseases.
Many non-neuronal cell types (particularly astrocytes) contribute to
intercellular signaling in the central nervous system at several levels
(7-9). Astrocytes, as well as other glia, are in intimate contact with
neurones and have projections that are located at neuronal synapses
(10). Indeed, chemical synapses and gap junction connections between
astrocytes and neurones have been identified (11, 12). Astrocytes
possess receptors for numerous transmitters (e.g. glutamate,
-aminobutyric acid (GABA), acetylcholine, ATP, bradykinin; reviewed
in Ref. 7) and so play important, active roles in synaptic activity.
Activation of astrocytes by transmitters released from neurones has
been reported at levels of transmitter concentrations found outside (but adjacent to) synaptic clefts (13, 14). Astrocytic activation is
usually manifest as a rise of intracellular Ca2+
concentration ([Ca2+]i) due to
release of Ca2+ from internal stores as well as
Ca2+ uptake from the extracellular space (9, 14-16). This
fundamental initial response correlates with neuronal synaptic
activity, and a rise of [Ca2+]i in
one astrocyte can initiate Ca2+ waves that propagate across
significant distances via adjacent astrocytes (17-19). This represents
a means of intercellular signaling in the brain that parallels and
modulates classical neuronal synaptic communication and, as such, is of
fundamental importance to central neuronal activity (8, 20). Within an
individual astrocyte, a rise of
[Ca2+]i can also initiate
important processes. In particular, elevated
[Ca2+]i triggers glutamate release
which modulates neuronal activity via extrasynaptic metabotropic
glutamate receptors (19, 21). Indeed, astrocytes are capable of
releasing glutamate via regulated,
Ca2+-dependent exocytosis, in addition to
reverse-mode uptake systems (17).
In the present study, we have examined how intracellular
Ca2+ stores, key components in astrocyte Ca2+
signaling coupled to receptor activation via generation of inositol trisphosphate, are modulated by prolonged hypoxia. Hypoxia is a key
feature of numerous cardiorespiratory diseases associated with
disturbance of higher brain functions (see above), and is also a well
known regulator of gene expression (22). Our results indicate that
hypoxia dramatically modulates intracellular Ca2+ stores,
primarily by causing mitochondrial Ca2+ loading.
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MATERIALS AND METHODS |
Astrocyte Culture--
To obtain astrocytes, cerebral cortices
were removed from 6-8 day old Wistar rat pups and placed immediately
in ice-cold buffer solution consisting of 10 mM
NaH2PO4, 2.7 mM KCl, 137 mM NaCl, 14 mM glucose, 1.5 mM
MgSO4, and 3 mg/ml bovine serum albumin. Meninges were
removed using fine forceps, and whole cortices were then minced gently
with a mechanical tissue chopper (McIlwain) and dispersed into the same
buffer containing 0.25 µg/ml trypsin, at 37 °C for 15 min. Trypsin
digestion was halted by the addition of an equal volume of buffer
supplemented with 16 µg/ml soy bean trypsin inhibitor (SBTI, type
I-S; Sigma), 0.5 µg/ml DNase I (EC 3.1.21.1 type II from bovine
pancreas; 125 kilounits/ml; Sigma) and 1.5 mM
MgSO4. The tissue was then pelleted by centrifugation at
1300 rpm for 90 s following which the supernatant was removed and
the cell pellet resuspended in 2 ml of buffer solution containing 100 µg/ml SBTI, 0.5 µg/ml DNase I, and 1.5 mM
MgSO4. The tissue was subsequently triturated gently with a
fire polished Pasteur pipette. After allowing larger pieces of tissue
to settle for 5 min, the cell suspension was taken and centrifuged at
1300 rpm for 90 s before resuspension into 60 ml of culture medium
(Eagle's minimal essential medium supplemented with 10% fetal calf
serum and 1% penicillin-streptomycin (GIBCO)). The cell suspension was then aliquoted into 2 × 25 cm2 flasks and onto glass
cover slips in 6- and 24-well tissue culture plates. Cells were then
kept in a humidified incubator at 37 °C (95% air; 5%
CO2). This was designated passage 1 and cells were used up
to a passage of 2. 4-6 h following plating, cells were washed
vigorously several times with fresh medium to remove non-adhered cells.
This resulted in a culture of primarily type I cortical astrocytes (as
confirmed by positive immunostaining with an anti-GFAP antibody).
Culture medium was exchanged every 3-4 days, and cells were grown in
culture for up to 14 days. All recordings were made from cells between
days 5-12.
Cells exposed to chronic hypoxia were subcultured in the same way as
control cells but 24 h prior to experimentation were transferred
to a humidified incubator equilibrated with 2.5% O2, 5%
CO2 balanced with N2 (termed chronically
hypoxic (CH)1 conditions).
Following exposure to hypoxia, cells were kept in room air for no
longer than 1 h while microfluorimetric recordings took place.
Corresponding control cells were maintained in a 95% air, 5%
CO2 incubator for the same period.
MTT Assay--
Cell viability was assessed using the
3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT)
reduction assay (23). Absorbency was measured using a spectrophotometer
at a test wavelength of 570 nm and reference wavelength of 630 nm.
Student's t test (unpaired) was used to determine the
significance of differences between means, with p values of
less than 0.05 being considered significant.
Microfluorimetric Recordings--
To measure cytosolic
[Ca2+], glass coverslips onto which cells had grown were
incubated in 2 ml of control solution containing 4µM
Fura-2AM for 1 h at 21-24 °C in the dark, as previously
described (24). Control solution was composed of: NaCl 135 mM, KCl 5 mM, MgSO4 1.2 mM, CaCl2 2.5 mM, Hepes 5 mM, and glucose 10 mM (pH 7.4, osmolarity
adjusted to 300 mosM with sucrose, 21-24 °C). Following this incubation period, fragments of coverslips were transferred into an 80-µl recording chamber mounted on the stage of
an inverted microscope, where cells were continuously perfused under
gravity at a rate of 1-2 ml min
1.
[Ca2+]i was determined using an
Openlab System (Image Processing & Vision Company Ltd, Coventry, UK).
Excitation was provided using a Xenon arc lamp (75 watts) and
excitation wavelengths (340 and 380 nm) were selected by a
monochromator (Till Photonics, Planegg, Germany). A quartz fiber-optic
guide transmitted light to the microscope and was reflected by a
dichroic mirror (Omega Optical, Glen Spectra Ltd, Stanmore, UK) into
the objective. Emission was collected through the objective and a
510-nm filter (40-nm bandwidth). Digital images were sampled at 14-bit
resolution by an intensified charge-coupled device camera (Hamamatsu
Photonics, Hertfordshire, UK). Fura-2 was excited alternately at 340 and 380 nm for between 120 and 180 ms (this varied on a day-to-day
basis, depending on dye-loading efficiency, but never varied between
control and hypoxic cells on any given day), and ratios of the
resulting images were produced every 4 s. Regions of interest
(ROI) were used to restrict data collection to individual cells. All
the imaging was controlled by Improvision software that included
Openlab 2.2.5 (Image Processing & Vision Company Ltd, UK) and operated
on a Macintosh PowerPC. Drugs and agonists were applied to cells as
indicated under "Results" by switching the inflow with a 6-way
Hamilton tap to one supplied by a reservoir of the relevant
composition. All experiments were conducted at room temperature
(21-24 °C).
For cytosolic [Ca2+] measurements, several parameters
were determined from collected data. Changes in
[Ca2+]i were taken from measuring
peak or plateau values and expressing them as the change in
fluorescence ratio from basal levels, determined for each recording.
Decay times are expressed as t1/2 values
i.e. the time taken for a response to decline to 50% of its
peak value. The size of bradykinin-evoked stores was taken from the
integral of transient responses recorded in Ca2+-free
perfusate. All results are expressed as means ± S.E., together with sample traces, and statistical comparisons were made using unpaired Student's t-tests. For all experiments reported,
4-8 cells in any one field of view were selected at random before the
experiment was performed, and data obtained from all selected cells
were included in the analysis. At least three repeats of each
experiment were performed.
Mitochondrial Ca2+ levels were examined using a confocal
system (see below), but in cells loaded with Rhod-2 (by incubation of
cells with 1.5 µM Rhod-2AM for 1 h at
21 °C-24 °C in the dark, followed by a 1 h period of
maintaining cells in control perfusate solution for further dye
de-esterification). Rhod-2 was only excited at one wavelength (543 nm),
and emission light was collected through the objective and a 570-nm
filter (40-nm bandwidth). Mitochondrial membrane potential measurements
were also made, using tetramethylrhodamine ethyl ester (TMRE). To load
this indicator, cell monolayers incubated in 2 ml of control solution
containing 300 nM TMRE for 20 min at 21 °C-24 °C in
the dark. TMRE was excited at one wavelength (546 nm), and emission
light was collected through the objective and a 576-nm filter (40-nm
bandwidth). During TMRE fluorescence recordings, cells were
continuously perfused with the same concentration of TMRE in the
presence of 1 mM ascorbate (to prevent TMRE oxidation and
suppress autoquenching, Ref. 25). Under these conditions photobleaching
was significantly reduced, and the slow decrease in the fluorescence
level caused by dye bleaching or dye loss was not considered
depolarization. Changes in mitochondrial membrane potential
(
m) were determined from groups of mitochondria using the region of interest (ROI) tool. TMRE fluorescence was not calibrated to membrane potential and so is plotted as arbitrary units (256 greyscale). Confocal images were obtained from cells plated and loaded
with dye as described above. Fragments of coverslips on which cells
were grown were transferred to a recording chamber as described for
microfluorimetric recording. This chamber was mounted on the stage of a
Zeiss Axiovert 200 M inverted microscope, equipped with a
C-Apochromat ×63 water immersion lens (1.2 numerical aperture) and
fitted with a Zeiss LSM 510 laser-scanning module. The 543-nm laser
line of a helium/neon laser was used for excitation of either Rhod-2 or
TMRE-loaded cells, and the emitted light (581 nm or 574 nm
respectively) collected by photomultiplier. 8 bit images (monitored
using a 256 gray scale, then converted to
F/Fo values for comparison) were
acquired as single frames (frame time 7 s), and 4 images were
averaged for each frame. All experiments were performed at room
temperature (21-24 °C). Loading conditions and illumination
intensities and duration were kept constant for all recordings.
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RESULTS |
Augmentation of [Ca2+]i Responses to
Bradykinin Caused by Chronic Hypoxia--
Fig.
1A shows representative,
bright-field images of type I cortical astrocytes cultured under
normoxic (upper) or hypoxic (lower) conditions. The cells were
typically flat and elongated in appearance, and hypoxia did not
noticeably alter their morphology. In order to examine cell viability
under hypoxic conditions, we performed colorimetric MTT assays (23),
and found that hypoxia (24-48 h) had no significant effect on cell
viability as compared with cells cultured under normoxic conditions
(Fig. 1B).

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Fig. 1.
Chronic hypoxia augments
[Ca2+]i responses to bradykinin without
affecting astrocyte viability. A, bright-field phase
contrast images of astrocytes used in the studies presented herein.
Cells were cultured under either normoxic (upper image) or
chronically hypoxic (C.H., lower image)
conditions. Scale bar applies to both images.
B, cell viability, determined using the MTT assay,
following culture under normoxic conditions (open bar) or
chronically hypoxic conditions (hatched bars) for the time
periods indicated. Data are normalized to control levels of cell
viability. C, sample increases of
[Ca2+]i evoked by application of
100 nM BK for the period indicated by the horizontal bars
in control (upper) and chronically hypoxic (C.H.,
lower) type I cortical astrocytes. Scale bars
apply to both traces. D, concentration-response
relationships indicating peak rises in
[Ca2+]i evoked by varying
concentrations of BK in control (open circles) and
chronically hypoxic (filled circles) astrocytes. Each point
plotted represents the mean ± S.E. response, n = 3-6 experiments in each case, from each of which at least three cells
were measured.
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Basal Ca2+ levels (determined before bradykinin (BK)
application) were not significantly different between chronically
hypoxic (CH) and control cells, being 0.53 ± 0.03 r.u. in
control cells (n > 12) and 0.58 ± 0.02 in CH
cells (n > 12). When astrocytes were perfused with a
solution containing 2.5 mM Ca2+, bath
application of 100 nM BK evoked a rapid rise of
[Ca2+]i due to release of
Ca2+ from internal stores mediated by production of
inositol trisphosphate, which declined with an extremely slow time
course in both control and CH cells (Fig. 1C). This slow
decline is likely due to Ca2+ influx (see Fig.
2, below). Full
concentration-response relationships are presented in Fig.
1D, which indicates that the EC50 for BK was
similar in both groups of cells, at ~5 nM. However, it
was notable that peak BK-evoked responses were consistently greater in
CH cells. Since the peak of the response reflects Ca2+
release from intracellular stores, rather than subsequent
Ca2+ influx, the rest of the study employed
Ca2+-free perfusates (containing 1 mM EGTA) to
examine release from stores in isolation, and we selected 100 nM BK as a maximally effective agonist concentration.

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Fig. 2.
Chronic hypoxia causes apparent augmentation
of Ca2+ liberation from intracellular stores by
bradykinin. A, sample rises in
[Ca2+]i evoked by application of
100 nM BK for the period indicated by the horizontal bars
in control (left) and chronically hypoxic (C.H.,
right) type I cortical astrocytes. For these experiments,
the perfusate was nominally Ca2+-free. Scale
bars apply to both traces. B, bar graphs
indicating mean values of parameters measured from recordings
exemplified in A: basal
[Ca2+]i levels, peak responses to
100 nM BK, the integral of the transient rise of
[Ca2+]i evoked by BK and the time
taken for the peak to decline to 50% of its maximal value. Data are
means ± S.E. taken from control recordings (open bars)
and chronically hypoxic cells (hatched bars). Statistical
differences between control and chronically hypoxic cells are indicated
by p values above each graph (n = 3-6
experiments in each case, from each of which at least four cells were
measured).
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Chronic Hypoxia Potentiates Liberation of Ca2+ from
Intracellular Stores by Bradykinin--
In Ca2+-free
solution, responses to 100 nM BK were transient in both
control (Fig. 2A, left) and CH (Fig.
2A, right) cells. However, a clear enhancement of
responses was observed in CH cells. The bar graphs of all measured
parameters are presented in Fig. 2B. Thus, as was the case
in Ca2+-containing perfusate, basal levels recorded in
Ca2+-free solution were not significantly different in the
two cell groups. The peak rise of
[Ca2+]i and the total amount of
Ca2+ liberated by BK (determined by integration of the
transient response) were both significantly greater in CH cells.
Finally, a significant slowing of the decay rate was also observed in
CH cells, as compared with controls.
The enhancement of BK-evoked rises of
[Ca2+]i seen in CH cells could
have been due to a more complete discharge of Ca2+ from
internal stores in this cell group, or due to CH cells having a greater
store size. To investigate this, we discharged intracellular stores
completely, by exposing cells to 10 µM cyclopiazonic acid (CPZ). We found in these cells that this agent depleted stores more
rapidly than the irreversible Ca2+-ATPase inhibitor,
thapsigargin. As illustrated in the sample traces of Fig.
3A and the mean data of Fig.
3B, the amount of Ca2+ discharged from
intracellular stores was clearly significantly greater in CH cells than
in controls. In addition, subsequent application of BK following
depletion of stores with either CPZ (Fig. 3C) or
thapsigargin (Fig. 3D) failed to evoke a rise in [Ca2+]i in either cell group,
indicating that both CPZ and thapsigargin fully discharged
intracellular stores.

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Fig. 3.
Cyclopiazonic acid and thapsigargin fully
deplete BK-sensitive intracellular Ca2+ stores in
astrocytes. A, representative rises in
[Ca2+]i evoked by bath application
of cyclopiazonic acid (CPZ; 10 µM which was present for
the period indicated by the horizontal bar) in control
(upper trace) and chronically hypoxic (C.H.;
lower trace) astrocytes. Scale bars apply to both
traces. B, bar graph indicating mean
integrals (with S.E. bars) of the transient rise of
[Ca2+]i evoked by CPZ. Data are
taken from control recordings (open bars) and chronically
hypoxic cells (hatched bars). Statistical difference between
control and C.H. cells is indicated by p value shown
(n = 4-6 experiments in each case, from each of which
at least four cells were measured). C, sample lack of
response to BK in control (left) and chronically hypoxic
(right) astrocytes following 5-min pretreatment with 10 µM CPZ. D, as in C, except
that cells were previously exposed to 1 µM thapsigargin
for 20 min prior to recording. Scale bars apply to
C and D.
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Chronic Hypoxia Inhibits Plasmalemmal
Na+/Ca2+ Exchange--
While the data of Fig.
2 suggest that BK-sensitive Ca2+ stores are greater in CH
cells, the enhanced responses could be due to altered buffering of
cytosolic Ca2+ once liberated from stores. One such
mechanism to account for the transient responses of cells to BK in
Ca2+-free perfusate is Ca2+ extrusion across
the plasma membrane on transporters such as the
Na+/Ca2+ exchanger (NCX). To investigate the
role of NCX in shaping the responses to BK, we applied the agonist in
Na+-free solutions (replaced with NMDG). As illustrated in
Fig. 4A and the averaged data
of Fig. 4B, this maneuver caused a small increase in the
peak [Ca2+]i response, which did
not reach statistical significance. However, there was a significant
increase in the transient integral, due to a slowing of the time course
of decay (Fig. 4B). In CH cells, no significant differences
were observed in either the amplitude or time course of the response
(Fig. 4, A and B). These results indicate that
NCX is not the major mechanism for Ca2+ extrusion following
release from intracellular stores in astrocytes, and also that
modulation of such exchange cannot account for the enhanced responses
observed in CH cells. However, perhaps more importantly, the
significant influence of NCX in shaping the responses seen in control
cells was not present in CH cells.

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Fig. 4.
Chronic hypoxia inhibits plasmalemmal
Na+/Ca2+ exchange. A,
sample rises in [Ca2+]i evoked by
application of 100 nM BK for the period indicated by the
horizontal bars in control (left) and chronically hypoxic
(C.H., right) type I cortical astrocytes. In each
case, the perfusate was Ca2+-free and either contained
Na+, or Na+ was replaced with NMDG
(Na+-free, as indicated). Scale bars apply to
both traces. B, bar graphs indicating mean
values of parameters measured from recordings exemplified in
A: peak responses to 100 nM BK, the integral of
the transient rise of [Ca2+]i
evoked by BK and the time taken for the peak to decline to 50% of its
maximal value. Data are means ± S.E. taken from six control
recordings in the presence of Na+ (open bars) or
in its absence (shaded bars) and nine chronically hypoxic
cells in the presence of Na+ (hatched bars) or
in its absence (hatched, shaded bars), as
indicated. Statistical differences between control and chronically
hypoxic cells are indicated by p values above each graph.
(n = 4-6 experiments in each case, from each of which
at least four cells were measured).
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Chronic Hypoxia Potentiates Mitochondrial Ca2+
Loading--
Recent studies have indicated that inhibition of NCX can
arise due to excessive Ca2+ loading of mitochondria (26).
We therefore explored the possible involvement of mitochondria in the
enhancement of BK-evoked rises in
[Ca2+]i. Fig.
5A (left)
illustrates a recording from a control cell, which was firstly exposed
to the mitochondrial inhibitor FCCP (10 µM), which
was applied together with 2.5 µg/ml oligomycin (to prevent ATP
consumption by the F1F0-ATP synthase
functioning in reverse mode). This caused a transient rise of cytosolic
[Ca2+]. In the continued presence of FCCP and oligomycin,
application of 100 nM BK evoked rises in control cells that
were significantly greater than those evoked without mitochondrial
inhibition (Fig. 2). Responses in CH cells (e.g. Fig.
5A, right) differed from those seen in control
cells in two important aspects. Firstly, mitochondrial inhibition
caused significantly greater rises of [Ca2+]i (mean data shown in Fig.
5B) and secondly, the subsequent application of BK evoked
rises in [Ca2+]i, which were not
significantly different from those observed in CH cells in the absence
of mitochondrial inhibitors. It is also noteworthy that during
mitochondrial inhibition, there were no significant differences in the
responses to BK observed between control and CH cells, as determined by
integration of the transients.

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Fig. 5.
Chronic hypoxia enhances mitochondrial
Ca2+ content. A, sample rises in
[Ca2+]i evoked by application of
10 µM FCCP together with 2.5 µg/ml oligomycin, and in
the additional presence of 100 nM BK. Following washout,
cells were exposed to 1 µM thapsigargin (TG).
Drug applications were for the periods indicated by the horizontal bars
in control (left) and chronically hypoxic (C.H.,
right) astrocytes. In each case, the perfusate was
Ca2+-free, and scale bars apply to both traces.
B, bar graph indicating mean integrals (with S.E.
bars) of the transient rise of
[Ca2+]i evoked by FCCP and
oligomycin (left), and by BK in the continued presence of
FCCP and oligomycin (right). Data are taken from control
recordings (open bars) and chronically hypoxic cells
(hatched bars). Statistical difference between control and
C.H. cells is indicated by p value shown. (n = 4-6 experiments in each case, from each of which at least three
cells were measured).
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Data presented in Fig. 5 suggest that the cytosolic Ca2+
response to BK may be attenuated in control (but not CH) cells due to
Ca2+ uptake by mitochondria. In further support of this
idea, responses to 10 µM CPZ were also significantly
potentiated (from 9.15 ± 0.84 r.u. (n = 14)
to 19.1 ± 2.56 r.u., p < 0.002, n = 18) during mitochondrial inhibition with FCCP (10 µM) and oligomycin (2.5 µg/ml), as Fig.
6 illustrates.

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Fig. 6.
Mitochondrial inhibition enhances CPZ-evoked
Ca2+ transients. Representative rises in
[Ca2+]i evoked by bath application
of cyclopiazonic acid (CPZ; 10 µM which was present for
the period indicated by the horizontal bar) in untreated
controls (taken from Fig. 3 for clarity) and in control cells during
continued exposure to 10 µM FCCP and 2.5 µg/ml
oligomycin (which were applied 1 min before the commencement of the
traces), as indicated.
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The enhanced response to FCCP and oligomycin observed in CH cells (Fig.
5A, right) strongly suggested that CH caused
excessive mitochondrial Ca2+ accumulation, as compared with
controls. To investigate this in more detail, we examined confocal
images of control and CH cells loaded with the mitochondrial
Ca2+ indicator, Rhod-2. To visualize the structure of
mitochondria in these cells, we firstly labeled mitochondria with
Mitotracker (Fig. 7A).
Clearly, mitochondria form extensive, complex networks in both control
and chronically hypoxic cells, and the mitochondrial density did not
appear altered by chronic hypoxia. Fig. 7B shows typical
images taken under identically matched exposure conditions in control
and CH astrocytes, at two levels of magnification. Mitochondria are
evident from their distribution and shape throughout the cell (see for
example Ref. 27), although the Rhod-2 images appear more fragmented
that the images obtained with Mitotracker. Clearly, fluorescence is
greater in the CH cells. The dashed lines in each upper trace show the
point at which a line scan was performed, and the corresponding pixel
intensities are plotted in Fig. 7C. Peaks in each trace
correspond to the line scan crossing mitochondria, and such peaks are
consistently greater in CH cells, reflecting a higher mitochondrial
Ca2+ content. It is noteworthy that while the peaks in the
line scans were greater in amplitude in CH astrocytes, the absolute
number was similar, indicating that hypoxia did not increase the number or density of mitochondria. These traces were typical of at least six
different images of both control and CH astrocytes. While not
quantitative, these data further support the Fura-2 data of Fig. 5
indicating that CH causes increased mitochondrial Ca2+
loading.

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Fig. 7.
Chronic hypoxia increases the
Ca2+ content of mitochondria in astrocytes.
A, confocal images of control (left) and CH
(right) astrocytes stained with the mitochondrial marker,
Mitotracker. B, upper traces, confocal
images of control (left) and CH (right)
astrocytes loaded with the mitochondrial Ca2+ indicator,
Rhod-2. The dashed line in each case shows the line of scan
used to generate the plots in C, and in each case is 14 µm
in length. Lower traces, higher resolution images acquired
as for upper traces. Scale bars shown at the
bottom left of each pair of images in A and
B indicate 15 µm. C, plots of pixel intensity
along the lines of scan shown in B (upper
traces). Peaks in each trace correspond to line scan crossing
individual mitochondria.
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Chronic Hypoxia Hyperpolarizes Mitochondrial Membrane
Potential--
Accumulation of Ca2+ by mitochondria is
dependent on the maintenance of the mitochondrial membrane potential,
(
m). We therefore examined whether
m
differed between control and CH cells, using the
m
indicator, TMRE. As can be seen from Fig.
8A, confocal images were
consistently brighter in CH cells as compared with control cells. While
individual mitochondria could not be resolved with the clarity found
using Rhod-2 (Fig. 7), staining appeared more punctate in CH cells,
consistent with greater accumulation into specific organelles.
Furthermore, the regions of brightest intensity (in both control and CH
cells) were found in parts of cells close to nuclei. This is also
reflected in the line scan plots of Fig. 8B (made using
lines indicated in the corresponding images), where in each case peaks
are seen either side of a trough in the plot which represents scanning
over the nuclear region of the image. These peaks are clearly greater
in the CH cells, an observation consistently seen in at least six
separate experiments.

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Fig. 8.
Chronic hypoxia hyperpolarizes astrocyte
mitochondrial membrane potential. A, confocal images of
control (left) and CH (right) astrocytes loaded
with the m indicator, TMRE. The dashed line
in each case shows the line of scan used to generate the plots in
B, and in each case is 14 µm in length. B,
plots of pixel intensity along the lines of scan shown in
A.
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We also employed TMRE to monitor fluorescent signals using conventional
(non-confocal) imaging, while cells were under continual perfusion.
Again, fluorescence observed in CH cells was clearly greater than in
control cells, and the brightest regions of each cell were found in
cytosolic areas close to the nucleus (Fig. 9A). Despite continual loss of
signal, we could also observe an increased rate of signal loss in both
cell types when exposed to 10 µM FCCP in the presence of
2.5 µg/ml oligomycin (Fig. 9A, lower
traces, and Fig. 9B). These findings strongly suggest
that CH causes a hyperpolarization of 
m, an effect
which is most likely responsible for the increased mitochondrial
accumulation of Ca2+ indicated by experiments described
earlier.

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Fig. 9.
Dissipation of mitochondrial membrane
potential by FCCP. A, fluorescent images obtained
from control (left) and chronically hypoxic
(C.H., right) astrocytes. Images were obtained
before (upper) and after (lower) bath application
of 10 µM FCCP and 2.5 µg/ml oligomycin.
Calibration bar applies to all images. B, mean
time course of the decline in TMRE fluorescence in control (open
circles, n = 12 cells) and chronically hypoxic
(filled circles, n = 14 cells). At the point
indicated by the arrow, the Ca2+-free perfusate
was exchanged for one containing 10 µM FCCP and 2.5 µg/ml oligomycin.
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DISCUSSION |
Cellular effects of cerebral hypoxia/ischemia have received
intense interest over many years, not only because of the associated neuronal damage and death (28, 29), but also because prolonged, milder
hypoxic episodes can lead to deleterious effects on higher brain
functions (1-3) and also to an increased likelihood of subsequent
development of dementias (4-6). The majority of studies to date have
focused on the effects of hypoxia/ischemia on central neurones, with
perhaps less attention given to astrocytes or other, non-neuronal cell
types in the central nervous system. The importance of astrocyte
function to central intercellular signaling is currently receiving
increasing attention (see the Introduction), in part due to emerging
awareness of their anatomical proximity and important physiological
regulation of neuronal synaptic transmission (11-14).
Since Ca2+ signaling is one of the major forms of
communication between astrocytes and is also a major factor in the
physiological activity of individual astrocytes, we have addressed the
question of whether prolonged hypoxia can modulate such signaling,
using primary cultures of rat type I cortical astrocytes, a well
established system for studying astrocyte function, and their robust
responses to BK (30-32). Our initial observation was that BK evoked
significantly greater release of Ca2+ from intracellular
stores in CH cells than was observed in control cells (Fig. 2). This
was not due to altered intracellular signaling between BK receptors and
Ca2+ stores, since CPZ also liberated more Ca2+
from stores in CH cells (Fig. 3). Likely possibilities to account for
this were that the BK-sensitive (endoplasmic reticulum; ER) stores
contained greater levels of Ca2+ following chronic hypoxia
or that, once liberated from the ER, Ca2+ was less
efficiently cleared and so could accumulate in the cytosol to a greater
concentration. Mechanisms for cytosolic Ca2+ clearance
include re-uptake into ER stores or other organelles, and
Ca2+ extrusion via NCX or Ca2+-ATPase.
Re-uptake into the ER was deemed unlikely in the continued presence of
agonist, and so we firstly investigated a possible role of NCX. This
was also prompted by a recent report indicating that ongoing, acute
hypoxia inhibits NCX in vascular smooth muscle (33). Results presented
in Fig. 4 indicate that in control cells, NCX plays a significant role
in shaping the transient rise of cytosolic [Ca2+] when ER
stores are discharged with BK. By contrast, NCX appeared to be
non-functional in CH cells (Fig. 4). Thus, CH somehow appeared to
inhibit NCX in cortical astrocytes. Importantly, however, the absence
of functional NCX could not account fully for the enhanced cytosolic
rise in [Ca2+] seen in response to BK application in CH cells.
At present, the underlying mechanisms accounting for inhibition of NCX
by prolonged hypoxia remain to be determined. However, a recent report
has demonstrated that accumulation of Ca2+ by mitochondria
specifically inhibits NCX in COS cells transfected with a bovine
Na+/Ca2+ exchanger (26). On the basis of this
report, we investigated the Ca2+ content of mitochondria in
astrocytes. Fig. 5 clearly demonstrates that CH leads to excessive
Ca2+ loading of mitochondria, as evidenced by the enhanced
response to application of FCCP and oligomycin, and this observation is reinforced by the confocal images of astrocytes loaded with the mitochondrial Ca2+ indicator, Rhod-2 (Fig. 7).
Additionally, the subsequent exposure to BK (and also to CPZ; Fig. 6)
in the continued presence of mitochondrial inhibitors caused a markedly
increased rise of cytosolic [Ca2+] in control cells,
while responses in CH cells were unchanged. These data indicate that,
in control astrocytes, BK-evoked rises in
[Ca2+]i are limited due to
Ca2+ buffering into mitochondria. This finding is in
accordance with numerous studies, which have documented the close
anatomical and functional interactions of mitochondria and the ER
(e.g. Refs. 34-36). Importantly, Ca2+ buffering
into mitochondria does not appear to occur in CH cells, presumably
because the mitochondria in these cells already contain excessive
amounts of Ca2+ and so are incapable of acquiring more.
Such a suggestion is supported strongly by the Rhod-2 confocal images
and associated pixel intensity lines scans of Fig. 7, which
demonstrated marked increases in punctate staining in CH cells. Such
mitochondrial accumulation of Ca2+ is most likely due to
mitochondrial hyperpolarization (upon which mitochondrial
Ca2+ accumulation is dependent), a view supported by both
confocal and conventional images acquired using the indicator TMRE.
This dye accumulates in mitochondria in proportion to the
m; the more hyperpolarized
m is, the more
dye accumulates. Clearly, in CH cells, increased (and more punctate)
fluorescence was observed which reflects a hyperpolarization of
mitochondria in astrocytes following a period of prolonged hypoxia.
Thus, the present study indicates that CH causes mitochondrial
hyperpolarization, which is likely to account for excessive accumulation of Ca2+, although a possible additional effect
of hypoxia to inhibit mitochondrial Na+/Ca2+
exchange has not been discounted, and is worthy of future study. Two
important consequences arise from this mitochondrial Ca2+
overload: firstly, via a mechanism yet to be identified, mitochondrial Ca2+ loading inhibits the plasmalemmal NCX (see also Ref.
26). Secondly, Ca2+-overloaded mitochondria are unable to
participate in buffering of Ca2+ liberated into the cytosol
from the ER following agonist application. The mechanism(s) by which
prolonged hypoxia leads to hyperpolarization of
m will
be the focus of future work. A recent study has suggested that
m can be hyperpolarized by
Ca2+-dependent dephosphorylation of cytochrome
c oxidase (37). This "molecular-physiological
hypothesis" (see also Ref. 38) is dependent on the mitochondrial
ATP:ADP ratio, which may well be altered under hypoxic conditions.
Importantly, this hyperpolarization in turn leads to increased
formation of reactive oxygen species (ROS), and ROS have been suggested
by others to cause irreversible inhibition of the plasmalemmal
Na+/Ca2+ exchanger (39), a suggestion
consistent with our observed lack of NCX function seen in CH astrocytes
(Fig. 4).
The concept that cellular ROS levels increase during prolonged periods
of hypoxia is not uncontested, but is currently gathering momentum.
Recently, a number of groups have suggested that the source of
increased cellular ROS during hypoxia is mitochondrial (37, 38,
40-43). The present study is also in accordance with these findings,
and indicates that hypoxia may increase ROS production via
mitochondrial hyperpolarization. Our findings are likely to have
important implications for the understanding of cellular damage and
death in the central nervous system following periods of hypoxia or
ischemia (44, 45).