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The Journal of Neuroscience, February 1, 1998, 18(3):932-947
Mitochondrial Membrane Potential and Nuclear Changes in Apoptosis
Caused by Serum and Nerve Growth Factor Withdrawal: Time Course and
Modification by (
)-Deprenyl
J. S.
Wadia1,
R. M. E.
Chalmers-Redman2, 3,
W. J. H.
Ju2,
G. W.
Carlile2,
J. L.
Phillips2,
A. D.
Fraser2, 3, and
W. G.
Tatton2, 3
1 Department of Physiology, University of Toronto,
Toronto, Ontario, Canada M55 1A8, 2 Department of
Physiology, Dalhousie University, Halifax, Nova Scotia, Canada B3H 4H7,
and 3 Department of Neurology, Mount Sinai School of
Medicine, New York, New York 10029-6511
 |
ABSTRACT |
Studies in non-neural cells have suggested that a fall in
mitochondrial membrane potential (
M) is one of
the earliest events in apoptosis. It is not known whether neural
apoptosis caused by nerve growth factor (NGF) and serum withdrawal
involves a decrease in M. We used epifluorescence and
laser confocal microscopy with the mitochondrial potentiometric dyes
chloromethyl-tetramethylrosamine methyl ester and
5,5
,6,6-tetrachloro-1,1,3,3-tetraethybenzimidazol carbocyanine
iodide to estimate M. PC12 cells were differentiated in media containing serum and NGF for 6 d before withdrawal of trophic support. After washing, the cells were incubated with media
containing serum and NGF (M/S+N), media without serum and NGF, or media
with the "trophic-like" monoamine oxidase B inhibitor, (
)-deprenyl. Mitochondria in cells without trophic support underwent a progressive shift to lower M values that was
significant by 3 hr after washing. The percentages of cells with
nuclear chromatin condensation or nuclear DNA fragmentation were not
significantly increased above those for cells in M/S+N until 6 hr after
washing. Replacement of cells into M/S+N or treatment with
()-deprenyl markedly reduced the proportion of mitochondria with
decreased M. Measurements of cytoplasmic peroxyl
radical levels with 2,7-dihydrodichlorofluorescein fluorescence and
intramitochondrial Ca2+ with
dihydro-rhodamine-2-acetylmethyl ester indicated that cytoplasmic peroxyl radical levels were not increased until after 6 hr, whereas increases in intramitochondrial Ca2+ paralleled the
decreases in M. ()-Deprenyl appeared to alter the
relationship between intramitochondrial Ca2+ levels
and M, possibly through its reported capacity
to increase the synthesis of proteins such as BCL-2.
Key words:
apoptosis; mitochondrial membrane potential; DNA
cleavage; chromatin condensation; JC-1; CMTMR; Rhod-2AM; DCFH
| |
INTRODUCTION |
Neuronal apoptosis in the developing
nervous system results from inadequate neurotrophic support (Lo et al.,
1995
). Apoptosis may also contribute to human neurodegeneration
(Chalmers-Redman et al., 1997), possibly because of increases in
reactive oxygen species (ROS) produced by defective mitochondrial
energy production (Richter et al., 1995; Beal, 1996). Various findings
link inadequate neurotrophic support and mitochondrial dysfunction in
the genesis of neuronal death (Frim et al., 1993; Mattson et al., 1993;
Kirschner et al., 1996).
A loss of mitochondrial membrane potential (M)
may be a critical mediator of apoptosis. The outward pumping of protons
across the inner mitochondrial membrane produces a proton gradient that drives the conversion of ADP to ATP and is reflected by
M (Sherratt, 1991). Measurements of whole-cell
potentiometric dye fluorescence have indicated that M
is reduced before the appearance of apoptotic nuclear changes in a
variety of blood, hepatic, and immune cell models (Susin et al.,
1996b). Similarly, decreased potentiometric dye uptake has suggested a
fall in M in sympathetoblasts after nerve growth
factor (NGF) withdrawal (Deckwerth and Johnson, 1993). Accordingly,
neurotrophic withdrawal may reduce M.
The relationship between M and apoptotic initiation
is uncertain. Decreased M induces opening of the
mitochondrial permeability transition pore (PTP), which may lead to the
release of mitochondrial apoptosis initiation factors (AIFs) (Marchetti
et al., 1996a; Susin et al., 1996a; Zamzami et al., 1996a). Cytochrome
C can facilitate apoptosis when released into the cell cytoplasm (Liu et al., 1996; Kluck et al., 1997). An overall decrease in
M, however, was reported to occur late in
apoptosis, well after the release of cytochrome C from mitochondria
(Yang et al., 1997). It has not been determined how changes in
M progress across populations of mitochondria in a
single cell. Loss of M in a small proportion of
mitochondria might be sufficient to initiate apoptosis.
()-Deprenyl reduces nerve cell death caused by a mitochondrial toxin
(for review, see Tatton and Chalmers-Redman, 1996), possibly by
increasing neurotrophic factor synthesis by astrocytes (Biagini et al.,
1994; Seniuk et al., 1994; Semkova et al., 1996). ()-Deprenyl also
increases reduced ubiquinone, suggesting that a mitochondrial action
may be involved in the increased neuronal survival (Koutsilieri et al.,
1996).
()-Deprenyl reduces the apoptosis of partially neuronally
differentiated PC12 cells caused by NGF and serum withdrawal (Tatton et
al., 1994b). We used epifluorescence and confocal laser microscopy in
that model to examine effects of NGF and serum-borne neurotrophic factors or ()-deprenyl on M in apoptosis. NGF and
serum withdrawal progressively shifted M across
mitochondrial populations to lower values at times that preceded any
increase in apoptotic nuclear events. Reexposure to NGF and serum or
treatment with 10
9 M ()-deprenyl
largely prevented the shifts. Increases in intramitochondrial Ca2+ levels appeared to parallel the decreases in
M, whereas increases in cytoplasmic peroxide
levels were delayed relative to decreases in M.
| |
MATERIALS AND METHODS |
Cell culture. PC12 cells (American Type Culture
Collection, Rockville, MD) were propagated in minimum essential medium
(MEM) (Life Technologies, Gaithersburg, MD) with 10% horse serum (Life Technologies), 5% fetal bovine serum (Life Technologies), 2 mM L-glutamine (Life Technologies), 50 U/ml
penicillin, and 50 µg/ml streptomycin (PC12 media).
For counts of intact nuclei, PC12 cells were differentiated in 24 well plates in the above media supplemented with 100 ng/ml 7 S NGF
(Upstate Biolotechnology, Lake Placid, NY). Cells were seeded at a
density of 8 × 104 cells per well to
facilitate neuronal differentiation. Seeding at higher densities
prevented the full elaboration of processes under the influence of NGF.
After 6 d in media containing serum and NGF (M/S+N), the cells
underwent three washes in HBSS to remove NGF and serum-borne trophic
agents, and the media were replaced with one of the following: MEM only
(M/O), MEM with 109 M ()-deprenyl
(M/d), or M/S+N. ()-Deprenyl at 109
M was used because that concentration induces the maximum
survival in the partially differentiated PC12 cells (Tatton et al.,
1994b) and is insufficient to cause monoamine oxidase (MAO) inhibition in PC12 cells (Youdim et al., 1986).
For imaging studies, the PC12 cells were plated on collagen-treated
(Sigma, St Louis, MO) or poly-D-lysine-treated
(Mr 30,000-70,000; Sigma) coverslips and
differentiated in the same media as above. Cells were seeded at 1 × 104 cells per 12 mm coverslip to facilitate
extensive process outgrowth (see Fig. 1A). After
6 d in M/S+N, the cells were washed three times in HBSS, and the
media was replaced with M/O, M/d, or M/S+N.
Estimates of cell survival. At 3, 6, 12, and 24 hr after
washing, cells were harvested by tituration from the 24 well plates and
centrifuged at 500 × g for 5 min; the supernatant was
removed, and the pelleted cells were lysed with 200 µl of lysing
buffer (Zap-o-globin II). Intact nuclei were counted in a hemocytometer by "blinded" observers according to the method of Soto and
Sonnenschein (1985). A second estimate of cell survival was obtained
for the same time points by lightly staining the cells on coverslips
with 1% methylene blue and counting intact cells under interference contrast optics (see Fig. 1A1,A2). Counts of intact
cells on each coverslip were taken from 25 100× fields, the
coordinates of which on an x-y grid were specified by two
computer-generated random numbers. The random method served to count
~50% of the coverslip area. The accuracy was assessed by performing
counts of all of the cells on 5% of the coverslips and comparing them
with the randomly generated counts for the same coverslips. The values differed by less than ±3% (p > 0.05).
ApopTag and 4,4-dichloro-4-bora-3a,4a-diaza-s-indacene
(BODIPY)-conjugated dUTP in situ end labeling of nuclear DNA
fragmentation. Two terminal deoxynucleotidyl transferase (TdT)
methods were used for the in situ labeling of 3-OH DNA ends
generated by DNA cleavage: (1) the ApopTag two-step method (Oncor,
Inc., Gaithersburg, MD) and (2) a BODIPY-conjugated dUTP (Molecular
Probes, Eugene, OR) single-step method. After fixation, the coverslips
were rinsed with PBS, placed in equilibrating buffer, and then
incubated in a reaction buffer consisting of TdT, dUTP, and necessary
ions for 30 min at 37°C. After rinsing, coverslips were incubated
with peroxidase-conjugated anti-digoxigenin (introduced together with Triton X-100 and a blocking agent). The coverslips were then exposed to
0.5 mg/ml diaminobenzidine and 0.05% hydrogen peroxide to generate a
brown reaction product and mounted onto slides with Aquamount (BDH
Chemicals, Poole, UK). The percentage of cells with ApopTag-positive nuclei were counted using interference contrast optics (see Fig. 1C). Negative controls were performed by substituting
distilled water for the TdT enzyme. Positive controls were performed by first nicking the DNA with DNase 1 before applying the ApopTag reagents.
In the single-step procedure (Li et al., 1996), fixed cells on
coverslips were rinsed with PBS and incubated in 50 µl of TdT reaction buffer containing the following: 10 µl of 5× concentrated buffer solution, 5 µl of 25 mM cobalt chloride, 5 µl
(25 U) of TdT (Promega, Madison, WI), and BODIPY-conjugated dUTP
(B-dUTP, Promega) for 60 min at 37°C. After rinsing with PBS, the
coverslips were incubated in the same medium containing 0.5 nM dideoxy-dUTP instead of deoxynucleotides for 10 min. The
coverslips were then treated with 15 mM EDTA and Triton
X-100 in PBS. The coverslips were mounted in glycerol-PBS on
microscope slides and examined using ordinary epifluoresence microscopy
(excitation, 450-490 nm; long-pass emission, 515 nm) or confocal laser
microscopy (excitation, 488 nm; emission 530/30 nm).
Hoechst 33258 staining of nuclear chromatin.
2-(4-Hydroxyphenyl)5-(4-methyl-1-piperazinyl)2-5-bi-1H-benzimidazole
trihydrochloride pentahydrate (Hoechst 33258, Molecular Probes) binds
to contiguous A-T bases in DNA. Treated cells were fixed on coverslips
and exposed to 1 µg/ml Hoechst 33258 in PBS for 15 min at room
temperature. The cells were then washed three times in PBS and viewed
with epifluorescence microscopy using 340-380 nm excitation coupled with 430 nm long-pass emission filtration. The nuclei of the partially neuronally differentiated PC12 cells appeared oval with a septate pattern of fluorescence (see Fig. 1B). Upon chromatin
condensation, the nuclei decreased in size and became intensely
fluorescent (see Fig. 1B1), often with ring-like
structures (see Fig. 1B3) and apoptotic bodies (see
Fig. 1B2).
Chloromethyl-tetramethylrosamine methyl ester estimation of
M in paraformaldehyde-fixed cells. Cationic,
lipophilic dyes, such as rhodamine 1,2,3 (Rh123) and
tetramethylrosamine ester (TMRE) accumulate in the negatively charged
matrix of mitochondria (Johnson et al., 1981). Experiments using
hepatocytes have shown that the ratio of Rh123 or TMRE fluorescence in
the cytoplasmic and nuclear compartments can yield a reliable estimate
of M in single cells with an accuracy of 10-20 mV
(Ubl et al., 1996) for values of up to 150 mV. With Rh123 or TMRE,
M must be measured ad hoc in living, metabolically
active cells so that immunocytochemistry for specific proteins or
in situ end labeling (ISEL) for DNA fragmentation cannot be
applied to cells identical to those examined for
M.
Recently, fluorescent probes for M have been
developed that are retained in cells at the time of fixation and can be
used to examine M together with levels and
localization of specific proteins or nuclear DNA fragmentation (see
Macho et al., 1996). One of those dyes,
chloromethyl-tetramethylrosamine methyl ester (CMTMR, Mitotracker
Orange; Molecular Probes) has the same basic structure as Rh123 and
TMRE with the addition of a chloromethyl moiety, which reacts with
thiol groups on peptides. CMTMR enters mitochondria proportionally to
the difference between the negativity of the cytoplasmic compartment
and mitochondrial matrix, and the chloromethyl group reacts with thiols
on proteins and peptides to form aldehyde-fixable conjugates. Once
M is lost, Rh123 and TMRE are washed out of the
mitochondria, but CMTMR remains bound even after permeabilization and
fixation (Calarco, 1995; Chen and Cushion, 1994). Therefore, CMTMR
fluorescence represents the highest level of negativity difference in
the mitochondria during the period of dye exposure before fixation.
At 3, 6, 12, and 24 hr after washing, the medium in each well was
supplemented with 50 nM CMTMR and incubated at 37°C for 15 min. The solution was replaced with dye-free media for a further 10 min before washing with PBS. The cells were then fixed on ice for 10 min with 4% paraformaldehyde. After fixation, the cells were rinsed
with PBS, and the coverslips were mounted onto slides using
Aquamount.
The mitochondria were visualized using a Reichert Polyvar II
epifluorescence microscope (Leica, Heidelberg, Germany) equipped with
an oil immersion, Fluotar 100×, 1.32 numerical aperture (NA) objective
(see Fig. 1D). CMTMR was excited at 540 nm, and the 590 nm emission image was captured using a charge-coupled device (Javelin Electronics Inc.) attached to an integrated
silicon-intensified target device (Hammamatsou Photonics, Inc.) and
digitized using a Matrox Image LC video digitization board connected to
a 486DX2/66 personal computer (PC) running Metamorph (Universal Imaging
Corporation, West Chester, PA) or Northern Exposure (Empix Ltd.)
imaging software. Images were acquired using 512 × 480 or
512 × 512 × 8 bits per pixel resolution using 32 frame
averaging with no background correction (see Fig.
1D).
Average CMTMR mitochondrial fluorescence intensity was estimated for
each cell by acquiring intensity values from 10 different 4 µm2 regions within the cytoplasm and calculating a
ratio of the average of the 10 values divided by the average
fluorescence intensity within the nucleus (see Ubl et al., 1996). An
intensity ratio of 1.0 represented cytoplasmic fluorescence equal to
that in the nucleus. The values for 50 cells under each treatment from
three different experiments were compared statistically and presented as frequency distributions.
Confocal microscopy was then used to resolve individual mitochondria
labeled with CMTMR at the time points of 6 and 24 hr after washing (see
Fig. 1E). A Leica true confocal scanning microscope coupled to an argon-krypton laser (Omnichrome, USA) was used. A
pinhole of 50 was used with an excitation filter wavelength of 488 nm
and a long-pass emission filter of 590 nm. Images were scanned using an
oil immersion, 100×, 1.3 NA objective at 512 × 512 × 8 bits per pixel resolution, background offset of 0, and averaged 16-32
times in bidirectional scan mode. The images were saved in tagged image
file format (TIFF) and transferred to a 100 MHz Pentium PC. Image Pro
Plus for Windows software (Media Cybernetics) was used to threshold
individual mitochondrial outlines and then to measure the mean
intensity within each mitochondrion. The value for each mitochondrion
was normalized against mean intensity for the immediately adjacent
cytoplasm, and the values were presented as frequency
distributions.
Immunocytochemistry for tubulin and histones. Cells
grown on coverslips were fixed for 15 min with 4% paraformaldehyde at 4°C. Nonspecific binding of protein was blocked by incubating the
coverslips in 10% normal goat serum for 1 hr. The cells were permeabilized simultaneously with 0.1% Triton X-100 in PBS.
Incubations with primary antibodies were done overnight at 4°C. Mouse
anti-tubulin (1:100, Sigma) (see Fig. 5A1) and mouse
anti-histone (1:200; Boehringer Mannheim, Indianapolis, IN) (see Fig.
5B1,C1) were used in the presence 0.2% normal serum and
0.05% Triton X-100. Coverslips were washed with PBS four times and
incubated for 1 hr with Texas Red-labeled anti-mouse antibody (1:500)
in the dark at room temperature. The coverslips were then washed four
times in PBS and mounted in PBS/glycerol and sealed before viewing.
Some of the coverslips had been exposed to CMTMR for
M measurements before fixation or were immunoreacted
for tubulin or histone identification before in situ
labeling of nuclear DNA fragmentation using BODIPY-conjugated dUTP.
JC-1 estimation of M in living cells.
5,5,6,6-Tetrachloro-1,1,3,3-tetraethybenzimidazol carbocyanine
iodide (JC-1, Molecular Probes) is a lipophilic, cationic dye that
enters mitochondria in proportion to the membrane potential and forms
J-aggregates at the high intramitochondrial concentrations induced by
higher M values. Formation of the J-aggregates is
associated with a Stoke's shift in emission (Smiley et al., 1991) from
527 nm for the monomer to 590 nm for the J-aggregate. The dye allows
for a dual measurement of dye concentration that does not require the
measurement of a nuclear or cytoplasmic reference value.
Studies using isolated cardiac myocyte mitochondria have shown that the
527 nm emission from monomeric JC-1 increases almost linearly with
M potentials ranging from 46 to 182 mV, whereas the 590 nm J-aggregate emission is less sensitive to
M values less negative than 140 mv and is strongly
sensitive to potential values in the range of 140 to 182 mV (Di
Lisa et al., 1995). Calculation of the 527:590 nm fluorescence ratio
eliminates many of the potential pitfalls of fluorescent dye
measurements, which require comparison between different cellular
compartments. Because the ratio can be calculated on a pixel-for-pixel
basis, it eliminates the possibility that changes in the volume of the
emitter account for changes observed for one experimental condition
relative to another. JC-1 imaging using a confocal microscope offers a
finer resolution and sensitivity of mitochondrial potential in living cells and allows M to be estimated for single living
mitochondria.
At 6 and 24 hr after washing, the media from each well containing PC12
cells grown on coverslips was removed, and a modified medium containing
10 µg/ml JC-1 in prewarmed MEM was added. The cells were placed back
into the incubator (37°C, 5% CO2, 100% humidity)
for 10 min, and the cells were then washed twice with HBSS to remove
unbound dye. The coverslips were then placed into a thermostatically
controlled cell chamber (Fine Science Tools Ltd.) into which warmed MEM
was circulated. Thermoregulation of the chamber was achieved by passing
a current through an enclosed nichrome wire, which was controlled by
feedback from a thermocoupler immersed in the media to maintain media
temperature at 37 ± 0.1°C.
The JC-1-treated coverslips were excited at 488 nm, and emission
was recorded simultaneously at 527 and 590 nm into independent detectors. Images were scanned using an oil immersion, 100×, 1.3 NA
objective at 256 × 256 or 512 × 512 × 8 bits per
pixel resolution, background offset of 0, and averaged 16-32 times in
a bidirectional scan mode. The confocal pinhole aperture was set to 50, and the voltage to the photomultiplier tubes of each channel was
maintained at equal values. Illumination was limited to periods of
image acquisition. In this way, the two images were exactly in phase and represented the amount of monomeric and J-aggregate JC-1
fluorescence (see Fig. 7 for images generated by combining the 527 and
590 nm images into a single image). For each time point, 10-20
mitochondria were measured from 50 to 70 cells from each of three to
six coverslips taken from two duplicate experiments.
Each image pair was analyzed in TIFF format using Image Pro Plus for
Windows software (Media Cybernetics) by conversion from 8 bit format to
floating point format to avoid division errors. The 527:590 nm ratio
image was calculated by dividing the pixel values of the 527 nm image
with its corresponding pixel value of the 590 nm image and then scaling
the result by 50. Using the ratio image, individual mitochondria were
digitally traced from the corresponding 527 nm image, and the average
intensity was calculated by integrating all of the intensity values
within the bound area and dividing by the number of pixels. The
mitochondrial data from all cells within a single treatment group were
pooled and plotted as an aggregate frequency distribution
histogram.
Dihydro-rhodamine-2-acetylmethyl ester estimation of
mitochondrial free calcium. Dihydro-rhodamine-2-acetylmethyl ester
(Rhod-2AM, Molecular Probes) is structurally related to Rh123 and
undergoes a 3.4-fold enhancement of fluorescence on binding to Ca
2+ (Minta et al., 1989). The rhodamine portion of the
molecule causes the Rhod-2AM to be concentrated in mitochondria and
thereby serves as an indicator of mitochondrial Ca2+
levels. Cells were incubated at 37°C for 20 min in media containing 5 mM Rhod-2AM in MEM. The cells were washed with two changes
of HBSS and transferred to a temperature-controlled chamber (see above). Confocal images of the cellular fluorescence were obtained using a 100×, 1.3 NA oil immersion objective, excitation wavelength of
488 nm, emission wavelength of 575 nm, pinhole setting of 90, and
averaging of 16 images (see Fig. 8). Individual mitochondrial-like organelles within each cell were digitally traced, and the mean fluorescence was determined using Image Pro Plus for Windows software similar to the procedures above for CMTMR. Data for each treatment group were plotted as a frequency distribution histogram.
2,7-Dihydrodichlorofluorescein estimation of cytoplasmic
peroxide radicals. 2,7-Dihydrodichlorofluorescein
(DCFH2) is a sensitive fluorometric probe for
peroxide radicals. The addition of the acetate moieties gives the
molecule lipophilic properties that enables the use of
dihydrodichlorofluorescein diacetate (DCFH2-DA) as a
probe for H2O2 production in living cells
(LeBel et al., 1992). DCFH2-DA permeates the cell membrane
and is hydrolyzed into DCFH2, a nonfluorescent compound
that remains trapped within the cell. Oxidation of DCFH2 is
induced by H2O2 in a stoichiometric reaction
catalyzed by peroxidases, yielding the fluorescent product 2,7-dichlorofluorescein.
Stock DCFH2-DA solutions were prepared by dissolving
the solid in ethanol to a concentration of 1 mg/ml. Aliquots were
stored in the dark at 80°C. At 24 hr after washing, the media were
replaced with 100 µM DCFH2-DA in MEM. The
cells were incubated at 37°C, 5% CO2, and 100%
humidity in the dark for 30 min and then washed three times in HBSS
before placement into a temperature-controlled cell chamber. Confocal
images of the cellular fluorescence were obtained using a 100×, 1.3 NA
oil immersion objective, excitation wavelength of 488 nm, emission
wavelength of 530 nm, pinhole setting of 90, and line averaging of 16 images.
Cellular fluorescence was quantitated by measuring the average
intensity within the non-nuclear cytoplasm of each cell using Image Pro
Plus for Windows software and calculating the average intensity of the
cell body cytoplasmic area for each cell.
Statistical evaluation. To evaluate the data, the individual
measurements from different treatment groups were first analyzed using
Statistica software (StatSoft) to perform two-tailed independent sample
t testing. Levene's testing for homogeneity of variances showed that most pairs of samples were not homogeneous, and
2 evaluation of the distributions showed that most did
not fit a normal distribution (see Siegel, 1956). If the data were
nonhomogeneous and did not fit to a normal distribution, analysis with
parametric methods such as the t test might not provide
valid results. The data were therefore rank-ordered and compared in a
pairwise manner using Statistica software to perform nonparametric
Mann-Whitney U testing (Siegel, 1956). The method depends
on permutations to calculate significance values and therefore does not
require homogeneity of variances, that the underlying distributions for
the data be known, or that the values are linearly related.
| |
RESULTS |
Cell death, nuclear chromatin condensation, and DNA strand
breakage increases at 6 hr
As illustrated in Figure
1A2, cells that
underwent NGF and serum withdrawal and survived beyond 6 hr showed a
marked reduction in process number and length compared with cells
replaced into M/S+N (Fig. 1A1). Counts of intact
nuclei and intact methylene blue-stained cells (Fig.
2A) revealed
statistically insignificant losses of between 5 and 10% at 3 hr for
cells in M/O (p > 0.05 compared with M/S+N).
The losses increased to 10-15% at 6 hr (p < 0.05 compared with M/S+N), 60-70% at 12 hr (p < 0.0001), and 65-80% at 24 hr (p < 0.0001).
Hence about two-thirds of the total cell death found over 24 hr
occurred between 6 and 12 hr after trophic withdrawal. The addition of
M/d did not alter the nuclear or cell loss at 6 hr
(p > 0.05 compared with M/O) but increased both
the percentages of intact cells (Fig. 2A) and nuclei
(Fig. 2B) at both 12 and 24 hr
(p < 0.001 compared with M/O).
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Figure 1.
Estimation of cell survival, nuclear chromatin
condensation, nuclear DNA fragmentation, and mitochondrial membrane
potential. A1, Typical interference contrast micrograph
of methylene blue-stained PC12 cells that were partially neuronally
differentiated in M/S+N for 6 d and were then washed to remove
trophic proteins and immediately replaced in M/S+N to reestablish
trophic support at 12 hr before fixation for histology.
A2, Similar micrograph for cells that were washed and
trophically withdrawn by placement in M/O at 12 hr before fixation.
B, Fluorescence photomicrographs of in
situ nuclei stained with Hoechst 33258 at 12 hr after washing
and placement into M/O. Normal Hoechst 33258-stained nuclei showed a
diffuse, granular substructure with division by fine septate-like
structures (inset 1). Apoptotic nuclei showed dense
staining characterized by the formation of shrunken, intensely
fluorescent lobular structures, those being apoptotic bodies
(inset 2) or ring-like structures (inset
3). C, Interference contrast micrograph of cells
reacted for in situ detection of nuclear DNA
fragmentation using the Apop Tag method at 12 hr after washing and
placement into M/O. The method revealed nuclei that were shrunken in
comparison to those without evidence of DNA fragmentation. Nuclear DNA
fragmentation is evident in three of the nuclei, the external plasma
membranes of which appear intact using interference contrast
microscopy. D, Typical epifluorescence micrograph of
CMTMR fluorescence for a partially neuronally differentiated PC12 cell
grown in M/S+N and fixed 24 hr after washing and replacement into
M/S+N. CMTMR fluorescence from individual mitochondria was discernible
using epifluorescence microscopy with 1000×, 1.3 NA objective.
However, the thickness of the optical sections resulted in the
superimposition of the fluorescence signals from adjacent mitochondria.
E, Confocal micrograph of a cell in M/S+N at 24 hr after
washing. A pinhole value of 50 produced optical sections that were
sufficiently thin to allow for fluorescence measurements from
individual mitochondria and the immediately adjacent cytoplasm without
superimposition of fluorescence from nearby mitochondria.
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Figure 2.
Estimates of the time course of cell death and the
appearance of the nuclear stigmata (chromatin condensation and DNA
fragmentation) of apoptosis. A, Values at four time
points after washing for the two methods that we used to estimate cell
survival: counts of intact nuclei after plasma membrane lysing using a
hemocytometer, and counts of intact cells from coverslips after
fixation and methylene blue staining. B, Percentages of
cells with nuclear chromatin condensation and nuclear DNA fragmentation
at the same four time points.
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|
A small baseline percentage (1-2%) of nuclei with chromatin
condensation or nuclear DNA fragmentation were found in cells in M/S+N
at all four time points (Fig. 2B). Neither percentage was increased above the baseline for cells in M/O at 3 hr
(p > 0.05 compared with M/S+N), but both
chromatin condensation (~4%, p < 0.05 compared with
M/S+N)) and nuclear DNA fragmentation (~3.5%, p < 0.05 compared with M/S+N) showed small increases at 6 hr. Both
increased to >20% at the 12 hr time point (p < 0.001 compared with M/S+N) and exceeded 30% at the 24 hr time point
(p < 0.001 compared with M/S+N). Examples of
chromatin condensation (Fig. 1B) and DNA
fragmentation (Fig. 1C) in M/O cells at 9 hr after trophic
withdrawal are illustrated. Accordingly, the major increase in cells
showing the nuclear stigmata of apoptosis occurred after the 6 hr time
point. Our previous use of DNA electrophoresis gels to determine the
time course of DNA fragmentation in the partially neuronally
differentiated PC12 cells after trophic withdrawal (Tatton et al.,
1994b) revealed a time course similar to that found with the ISEL
method (also see BODIPY-dUTP below). In those studies, DNA ladders were
not detectable at 3 and 6 hr but were easily detectable at 9 and 12 hr
after placement in M/O.
Figure 2B illustrates that cells in M/d did not
differ in the percentage of nuclei with chromatin condensation or DNA
fragmentation from those in M/O at either 3 or 6 hr
(p > 0.05 compared with M/O). The percentage of
nuclei with either chromatin condensation or DNA fragmentation for
cells in M/d at 12 and 24 hr after washing exceeded those in M/S+N
(p < 0.05 compared with M/S+N) but was markedly
reduced compared with those in M/O (p < 0.0001 compared with M/O).
Changes In average cellular CMTMR fluorescence are present by
3 hr
Superimposition of fluorescence from adjacent mitochondria
made it impossible to measure CMTMR fluorescence reliably from individual mitochondria with epifluorescence microscopy (Fig. 1D). Accordingly, we compared cell body cytoplasmic
fluorescence with nuclear fluorescence (cytoplasmic/nuclear ratio) as
an estimate of average M in each cell (Ubl et al.,
1996). The left column of CMTMR cytoplasmic/nuclear ratio
distributions in Figure 3 for cells in
M/S+N at the four time points shows that the ratios ranged from 1.0 to
~5.0 with means of 2.2-2.4. The values did not differ significantly
for cells in M/S+N among any of the four time points (all
p < 0.05; see Table 1
for results of pairwise comparisons using both parametric and
nonparametric statistical tests). In contrast to those for M/S+N, the
ratio distribution for cells in M/O at 3 hr showed an overall shift to
lower values when compared with M/S+N at the same time point (Fig. 3,
Table 1, first row of distributions). The extent of the
shift to lower ratios is illustrated by the graph labeled
M/O in Figure 6A, which shows that more than 25% of the cells had average ratios of <1.6 in M/O at
3 hr, whereas only 11% of cells in M/S+N had ratios smaller than that
value (see Fig. 6A, graph labeled
M/S+N).
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Figure 3.
Estimation of the time course of changes in
average cellular M using epifluorescence imaging of
the rhodamine derivative CMTMR. Distributions for cell body cytoplasmic
CMTMR fluorescence divided by CMTMR nuclear fluorescence
(cytoplasmic/nuclear ratio) as an estimate of changes in average
M after washing in each cell for the three treatments
at four time points.
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Table 1.
Statistical testing of data for cytoplasm/nuclear ratio of
CMTMR fluorescence measurements taken using epifluorescence microscopy
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By 6 hr after washing and placement in M/O, there was a marked shift in
CMTMR cytoplasmic/nuclear ratios to lower values (probabilities ranging
from 1.51 × 102 to 1.07 × 104 depending on the statistical test used; see
Table 1). The distributions for the CMTMR cytoplasmic/nuclear ratios
for cells in M/O progressively shifted to lower and more highly
statistically significant values compared with those in M/S+N at 12 and
24 hr (Fig. 3, Table 1). The extent of the shift to lower ratios is
illustrated in Figure 6A, showing that ~47, 72, and
70% of cells had average ratios of <1.6 at 6, 12, and 24 hr,
respectively.
Treatment with 109 M ()-deprenyl
reduced the shift in the CMTMR cytoplasmic/nuclear ratio distributions
(Fig. 3, right column of distributions; see Fig. 6A)
so that ratios were significantly different for M/d relative to M/O
but not for M/d relative to M/S+N (Table 1).
CMTMR fluorescence for individual mitochondria have shifted
markedly by 6 hr
To address the possibility that changes in nuclear CMTMR binding
accounted for the above findings and to determine how changes in
average M within a cell relate to shifts in the
values for individual mitochondria within a single cell, we measured
mitochondrial CMTMR fluorescence relative to the fluorescence in the
immediately adjacent cytoplasm (mitochondrial/cytoplasmic ratio) using
confocal laser microscopy (Fig. 1E).
The distributions for cells in M/S+N (Fig.
4, top row) did not differ at
6 and 24 hr (p > 0.05), whereas those for cells
in M/O (Fig. 4, middle row) were progressively and
significantly shifted to lower values at the same time points (see Fig.
6, Table 2). The mean values for the
inset plots (each column of points in the insets present 15 mitochondrial/cytoplasmic ratios measured from mitochondria in a single
cell) show variability for different cells in M/S+N that is similar to
that found for the cytoplasmic/nuclear ratios presented above. The
insets show a surprisingly high variability in CMTMR fluorescence for
different mitochondria within a single cell in M/S+N. Relative CMTMR
fluorescence for individual mitochondria could vary as much as sixfold
within a single cell. Mitochondria in M/O showed a progressive fall and
narrowing of the CMTMR ratios when 6 and 24 hr were compared (Fig. 4).
()-Deprenyl (Fig. 4, bottom row of distributions) shifted
the mitochondrial/cytoplasmic distributions toward those for cells in
M/S+N. The shift was not complete because the Mann-Whitney
U test revealed a statistically significant difference
between the treatments at 6 and 24 hr (Table 2).
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Figure 4.
Estimation of the changes in M
for individual mitochondria using confocal microscopic imaging of CMTMR
fluorescence. Distributions of the CMTMR mitochondrial/cytoplasmic
ratios for the three treatments at the 6 and 24 hr time points together
with corresponding examples (inset) of the CMTMR
mitochondrial/cytoplasmic ratios for 20 mitochondria from 10 randomly
chosen cells.
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Table 2.
Statistical testing of data for mitochondrial/cytoplasmic
ratio of CMTMR fluorescence measurements taken using laser confocal microscopy
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Cells with chromatin condensation or nuclear DNA fragmentation have
reduced or absent M
The BODIPY-dUTP ISEL method for detecting nuclear DNA
fragmentation provided complementary results to those for the ApopTag method. For cells in M/O, the method revealed that 1.4 ± 0.2% of
nuclei showed evidence of DNA fragmentation at 3 hr and 3.1 ± 0.5, 26.2 ± 3.1, and 34.5 ± 5.2% at 6, 12 (Fig.
5A1,A2), and 24 hr,
respectively.
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Figure 5.
Nuclear DNA fragmentation demonstrated with
BODIPY-dUTP simultaneously with CMTMR staining to estimate
M. A1, A2, Low-power epifluorescence
micrographs of the same image fields of cells after 12 hr in M/O that
were immunoreacted for tubulin (A1) and also reacted
using the BODIPY-dUTP method for detecting nuclear DNA fragmentation
(A2). Confocal laser images of identical fields for
tissue dually reacted with an anti-histone antibody are shown in
B1 and C1, and BODIPY-dUTP is shown in
B2 and C2. D1-D3, E1-E3, Identical confocal image fields for serial image planes (separated by
~1.0 µm) through a group of partially neuronally differentiated PC12 cells in M/O at 6 hr after washing. Cultures are stained for
BODIPY-dUTP nuclear staining and CMTMR mitochondrial fluorescence, respectively.
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Immunoreactive nuclear histones were distributed in a diffuse,
lattice-like pattern in nuclei that did not show evidence of DNA
fragmentation in corresponding images for BODIPY-dUTP
(Fig. 5B1,B2, respectively). In nuclei demonstrating DNA
fragmentation, the histone immunoreaction was found to be marginalized
to the outer portion of areas of BODIPY fluorescence, as shown in
Figure 5, B1, B2, C1, and C2.
Figure 5, D1-D3 and E1-E3, shows identical
confocal image fields for serial image planes (separated by ~1.0
µm) through a group of cells in M/O at 6 hr. Figure
5D1-D3 shows the fluorescence for the B-dUTP reaction and
demonstrates a nucleus that has undergone DNA fragmentation and has
broken into three or more pieces. The CMTMR fluorescence in the
corresponding images (Fig. 5E1-E3) shows well defined
mitochondrial CMTMR fluorescence in all cells that do not show evidence
of nuclear DNA fragmentation, whereas the cell with nuclear DNA
fragmentation shows shrunken cytoplasm with markedly reduced CMTMR
fluorescence.
Cells in M/O at 6 hr after washing that showed evidence of nuclear DNA
fragmentation showed average mitochondrial/cytoplasmic ratios of 1.15 (Fig. 6D, left). Ratios
of 1.0-1.4 would fall among the lowest values in the 6 hr M/O
distribution in Figure 4 and would appear to represent ~20% of the
mitochondria in comparison to <2% of the mitochondria for cells in
M/S+N (Fig. 6B). Similarly, cells without evidence of
chromatin condensation had average CMTMR cytoplasmic/nuclear ratios
(Fig. 6D, right) that were not significantly different (p > 0.05) from those previously
found for cells in M/S+N at the same time point (Fig. 3). Those with
well defined chromatin condensation had significantly reduced CMTMR
cytoplasmic/nuclear ratios (p < 0.001) that
were in the lower range of those previously found for cells in M/O
(Figs. 3, 6A).
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Figure 6.
Time course of changes in the levels of CMTMR and
JC-1 fluorescence and differences in CMTMR fluorescence for cells with
or without nuclear DNA fragmentation or chromatin condensation. Each plot in A-C shows percentage of cells or mitochondria
with CMTMR fluorescence ratios or JC-1 527:590 nm fluorescence ratio
values that are less than values shown to correspond with specific
M levels in other cellular models. D,
Bar graphs of the CMTMR mitochondrial/cytoplasmic ratios and CMTMR
cytoplasmic/nuclear ratios for cells in M/O at 6 hr after washing with
and without evidence of nuclear DNA fragmentation or chromatin
condensation shown by staining with Hoechst 33258.
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Decreases in mitochondrial JC-1 fluorescence in living cells
parallel those found with CMTMR
To address the possibilities that changes in mitochondrial
size accounted for some or all of the changes in fluorescence or that
fixation altered the cytoplasmic/nuclear or mitochondrial/cytoplasmic ratios to differing extents for the three treatments, we maintained living cells in a temperature-controlled bath and used dual
simultaneous detection confocal laser microscopy to measure the 527 and
590 nm fluorescence of the JC-1 monomer and J-aggregate, respectively. Pixel-for-pixel algebraic addition of the 527 nm emission images colored green and 590 nm emission images colored red allowed us to
visualize mitochondria with low (green motochondria), medium (yellow
mitochondria), and high (red mitochondria) M levels in single images (Fig. 7,
insets). A mixture of green, yellow, and orange-red
mitochondria within single cells in M/S+N or M/d (Fig.
7A1,A2,C1,C2, insets) seemed to show the same wide range of
M for different mitochondria in single cells
indicated by CMTMR mitochondrial/cytoplasmic ratios (Fig. 4,
insets for M/S+N and M/d). A progressive reduction in
orange-red and then yellow mitochondria in cells in M/O at 6 and 24 hr
(Fig. 7B1,B2) revealed the same fall and narrowing of
M for the mitochondrial population in a single cell
as found with the CMTMR mitochondrial/cytoplasmic ratios.
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Figure 7.
Dual-emission images for JC-1 fluorescence. Each
inset image shows the effect of superimposition of the
recolored 527 and 590 nm images by the algebraic addition of
corresponding pixels: regions of mitochondria with predominant 527 nm
emission color green, those with overlapping 527 and 590 nm emission color yellow, and those with higher 590 nm
emission color orange-red in the superimposed images.
Each panel shows typical examples of superimposed JC-1 images and
527:590 nm emission ratio distributions for the pixel domains of
individual mitochondria for the three treatment conditions at 6 and 24 hr after washing. A1, A2, C1, C2, For cells in M/S+N and
M/d, respectively, show a mixture of green,
yellow, and orange-red mitochondria in a
single cell and appear to indicate a wide range of M
values for mitochondria within a single cell. The corresponding
distributions for these panels show that the majority of mitochondria
have 527:590 nm emission ratios that are <1.0, indicating high levels
of M. B1, Cells in M/O at 6 hr and
relative decrease in orange-red mitochondria and a
preponderance of yellow and green
mitochondria found in those cells. As shown in the accompanying
distribution, those changes reflect a shift in the mitochondrial
527:590 nm emission ratios to higher values, with a majority of the
mitochondria showing ratios of >1.0. B2, Cells in M/O
at 24 hr and little, if any, yellow or
orange-red. The accompanying distribution shows a
further shift of the emission ratios to values of >1.0.
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It should be noted that shifts of distributions of CMTMR
cytoplasmic/nuclear (Fig. 3) or mitochondrial/cytoplasmic (Fig. 4) ratios to lower values indicate a decrease in M. In
contrast, shifts of the distributions (Fig. 7) of the 527:590 nm ratios to lower values indicate an increase in M,
whereas increased ratios indicate decreased M values
(Di Lisa et al., 1995). Mitochondria in cells in M/O showed a
significant shift in the 527:590 ratio to higher values compared with
those in M/S+N at both 6 and 24 hr (Figs. 6C, 7, Table
3). The distributions for cells in M/d revealed a maintenance of the ratio for a majority of the mitochondria but similar to that found for the CMTMR mitochondrial/cytoplasmic ratios (Figs. 4, 6B); a distinct subpopulation of
mitochondria with ratios of >2.0 did not appear to respond to
()-deprenyl treatment at either 6 or 24 hr (Fig.
7C1,C2).
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Table 3.
Statistical testing of data for 527:590 nm ratio of JC-1
fluorescence measurements taken using laser confocal microscopy
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CMTMR cytoplasmic/nuclear ratio measured with epifluorescence
microscopy or whole cell cytofluorescence measurements with a
single-wavelength fluorescence emitter offer values that depend on both
M and the cytoplasmic density of mitochondria. Cell shrinkage would increase the number of mitochondria per unit of cross-sectional area and therefore the measurements. An increase in the
number of mitochondria per unit of volume or an increase in
mitochondrial size would increase whole cell cytofluorometric measurements independently of M when using a
single-wavelength emitter dye (Castedo et al., 1996). In contrast, the
use of CMTMR mitochondrial/cytoplasmic ratios or JC-1 with confocal
images provides measures of M that should be
uninfluenced by changes in mitochondrial mass.
Increased calcium levels in mitochondrial-like organelles parallel
preapoptotic decreases in M
Increases in intramitochondrial Ca2+ secondary
to high cytoplasmic Ca2+ levels and increases in ROS
(Richter et al., 1995) have been considered as primary factors in
causing reductions of M. To estimate mitochondrial
Ca2+ levels and cytoplasmic ROS levels in PC12 cells
that survived for 6 and 24 hr after NGF and serum withdrawal, we used
the fluorescent dye Rhod-2AM. The Rhod-2AM images showed localized
fluorescence in a subcellular component that appeared to correspond to
mitochondria. Both the intensity of fluorescence in individual
components and the number of visibly fluorescent components increased
greatly for cells in M/O (Fig.
8A2,B2) compared with
those in M/S+N (Fig. 8A1,B1). Cells in M/O often
showed a localized region of nuclear fluorescence that appeared to
correspond to the nucleolus, and high-resolution confocal images
revealed increased fluorescence in the background areas surrounding the
subcellular components.
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Figure 8.
Rhod-2AM as an indicator of mitochondrial
Ca2+. A1-A3, B1-B3, insets, Typical
confocal microscopic images of living cells stained with Rhod-2AM for
the three treatments at 6 and 24 hr after washing. The accompanying
graphs show the distributions for the Rhod-2AM fluorescence for subcellular structures likely to correspond to mitochondria at the two time points. Values above each distribution present the mean ± SD and the numbers of mitochondria examined in
each group. The distributions show that Ca2+ levels
in mitochondrial-like organelles were markedly elevated at both 6 and
24 hr after washing and placement in M/O and were only partially
reduced by treatment with ()-deprenyl.
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Distributions for the fluorescence in the subcellular component
revealed a significant shift to higher values for cells in M/O (Fig.
8A2,B2) compared with those in M/S+N (Fig.
8A1,B1) with an 80% average increase in fluorescence
at 6 hr and 92% average increase at 24 hr (p < 0.001 for the t test and Mann-Whitney statistical comparisons). Addition of ()-deprenyl to the MEM (Fig.
8A3,B3) shifted the distribution back toward the
lower values found for cells in M/S+N, but the mean values for the
distributions were increased by 62 and 53% above M/S+N values at 6 and
24 hr, respectively (p > 0.01).
Cytoplasmic peroxyl radical levels increase after decreases
in M
Cells in M/S+N showed low levels of
DCFH2 fluorescence (Fig.
9A1,B1). DCFH2
cytoplasmic fluorescence was not significantly increased at 6 hr in
cells in M/O (Fig. 9A2; p > 0.05) but was increased in cells surviving to 24 hr (Fig. 9B2,
p > 0.05). As illustrated by the image and the
distribution in Figure 9B2, nearby cells could show markedly
different levels of cytoplasmic DCFH2 fluorescence after
trophic withdrawal. ()-Deprenyl treatment at 24 hr returned the
average cytoplasmic DCFH2 fluorescence levels (Fig.
9B3) to within the range of those found in cells in M/S+N at
the same time points (p > 0.05).
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Figure 9.
DCFH fluorescence as an indicator of cytoplasmic
peroxyl radical levels. A1-A3, B1-B3, insets, Typical
examples of DCFH fluorescence of living cells in M/S+N, M/O, and M/d,
respectively, at 6 and 24 hr after washing. The accompanying
graphs show the distributions for the DCFH fluorescence
in the cellular cytoplasm. Values above each distribution present the
mean ± SD and the number of cells examined in each group. The
distributions show that cytoplasmic peroxyl radical levels were not
elevated at 6 hr when M was already markedly
decreased but were elevated in cells surviving to 24 hr.
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DISCUSSION |
Decreased mitochondrial membrane potential precedes nuclear
DNA cleavage, nuclear chromatin condensation, and cell death
One third of the average decrease in our estimators
of M found at 24 hr had occurred by 3 hr, and more
than half had occurred by 6 hr (Fig.
10A). The percentages
of cells with detectable nuclear chromatin condensation or DNA
fragmentation were not increased above baseline at 3 hr and
were increased by only 2-3% above baseline at 6 hr. Therefore a
decrease in M led the nuclear changes in an average
cell by at least 3 hr. The relative timing of cell death,
nuclear changes, and a decrease in the fluorescence of a
mitochondrially accumulated dye in our study using microscopic measurements seems similar to that found using whole-cell
cytofluorescence measurements in rodent embryo cells (Vayssiere et al.,
1994), thymocytes (Castedo et al., 1995; Petit et al., 1995), and
lymphocytes (Zamzami et al., 1995a) entering apoptosis.
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Figure 10.
Timing of cell death, the appearance of the
nuclear stigmata of apoptosis, Rhod-2AM levels in mitochondrial-like
organelles, cytoplasmic peroxyl radical levels, and decreases in
M in trophically withdrawn cells. A,
Time course for cells in MEM only; B, time course for
those in MEM with 109 M ()-deprenyl.
The number of intact nuclei (open triangles) determined
by cell lysing and direct nuclear counting and number of intact cells
(inverted open triangles) determined by methylene blue
staining yielded a decrease to 30-38% of MS+N values by 12 hr and
~20% by 24 hr. Average values for the CMTMR cytoplasmic/nuclear ratio (open circles) were used to estimate
M at 3 hr. CMTMR cytoplasmic/nuclear ratio, CMTMR
mitochondrial/cytoplasmic ratio (gray circles),
and the reciprocal of the JC-1 527:590 nm ratio (black
circles) are presented for cells in M/O at 6 hr and show decreases of 20, 39, and 53%, respectively. By 24 hr, surviving cells
showed average decreases in M of 60-70% depending
on the method used. Chromatin condensation (open
diamonds) and DNA cleavage (open squares) were
shown to increase above baseline only after 6 hr after trophic
withdrawal. Calcium accumulation into mitochondrial-like organelles
(gray squares) increased to 80% of that measured
in cells in MS+N, and cytoplasmic peroxyl radical levels (black
diamonds) increased linearly to 65% at 24 hr. Similar
measurements were performed on cells treated with M/d. All
measurements remained similar to MS+N values with the exception of
calcium accumulation into mitochondrial-like organelles, in which
levels increased to 60% beyond MS+N values at 6 hr and stabilized to
50% above control at 24 hr.
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Different timetables for apoptosis in partially neuronally
differentiated PC12 cells and fully neuronally differentiated PC12
cells
In a previous study of PC12 cell death after
trophic withdrawal (Mesner et al., 1995), cells were maintained in RPMI
1640 media with 1% horse serum and 50 ng/ml NGF. The horse serum was withdrawn after 12 d, and the NGF was withdrawn at 15 d. The
major period of PC12 cell death was delayed by ~3 hr compared with
the present study, and ~10% of nuclei in control cells were positive for DNA strand breaks, compared with 2% in our study. Changes in the
expression of a variety of genes in their trophically withdrawn, fully
differentiated PC12 cells appeared to occur at later times than those
found in the partially neuronally differentiated PC12 cells (Tatton et
al., 1996). The accelerated apoptosis in our cells may relate to the
rapidity of trophic withdrawal or to the constitutive presence of
proteins necessary for the progression of apoptosis. Undifferentiated
PC12 cells (Rukenstein et al., 1991) and partially neuronally
differentiated PC12 cells (Tatton et al., 1994b) do not require new
protein synthesis for apoptosis caused by trophic withdrawal. Fully
differentiated PC12 cells (Mesner et al., 1992) do require newly
synthesized protein for apoptosis, which may also account for the
slower progression of apoptosis.
Increased mitochondrial Ca2+ may be responsible
for the decrease in M after serum and NGF
withdrawal
Several factors can account for a decrease in
M, including increased cytosolic
Ca2+ levels with Ca2+ uptake into
mitochondria (Richter, 1993), elevated levels of ROS (Skulachev, 1996),
decreased electron transport (Krippner et al., 1996), or decreased
proton pumping by the mitochondrial respiratory chain (Wolvetang et
al., 1994). Our cells in M/O at 6 hr had decreased and relatively
uniform M levels but showed unchanged cytoplasmic
levels of peroxyl radicals according to DCFH2 fluorescence
(Fig. 10A). Hence increased cytoplasmic ROS levels
did not seem essential for the early decreases in
M.
Alternately, Rhod-2AM fluorescence suggested an inverse relationship
between intramitochondrial Ca2+ levels and
M in the cells at 6 and 24 hr (Fig.
10A). M should vary inversely with
intramitochondrial Ca2+ levels according to a
Nernstian relationship (Richter, 1993). Increases in cytosolic
Ca2+ are induced by trophic withdrawal and can lead
to apoptosis in PC12 cells (Mariggio et al., 1994; Fulle et al., 1997).
NGF, basic fibroblast growth factor (bFGF), and insulin-like growth
factor II (IGF-II) prevented increases in cytosolic
Ca2+ levels and concomitant decreases in
M in cultured hippocampal neurons deprived of glucose
(Mattson et al., 1993). bFGF and IGF-II reduce the death of PC12 cells
caused by serum and NGF withdrawal (Rydel and Greene, 1987; Rukenstein
et al., 1991) and are likely constituents of the serum used in the
present study. Hence the capacity of NGF- and serum-borne trophic
agents to maintain normal cytosolic Ca2+ levels
could explain the maintenance of M in our cells in
M/S+N and the decrease caused by their withdrawal.
Decreases in M induce the opening of a PTP in the
mitochondrial membranes (Zoratti and Szabo, 1995) and the consequent
rupture of the outer mitochondrial membrane, which may allow the
release of heat labile AIFs from the intermembrane mitochondrial space (Marchetti et
Top
Abstract
Introduction
