From the Institut für Physiologische Chemie, Universitätsklinikum, Hufelandstrasse 55, D-45122 Essen, Germany
Received for publication, May 1, 2002, and in revised form, October 17, 2002
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Direct reaction of NAD(P)H with
oxidants like singlet oxygen (1O2) has
not yet been demonstrated in biological systems. We therefore chose
different rhodamine derivatives (tetramethylrhodamine methyl ester,
TMRM; 2',4',5',7'-tetrabromorhodamine 123 bromide; and rhodamine
123; Rho 123) to selectively generate singlet oxygen within the
NAD(P)H-rich mitochondrial matrix of cultured hepatocytes. In a
cell-free system, photoactivation of all of these dyes led to the
formation of 1O2, which readily oxidized
NAD(P)H to NAD(P)+. In hepatocytes loaded with the various
dyes only TMRM and Rho 123 proved suited to generating
1O2 within the mitochondrial matrix space.
Photoactivation of the intracellular dyes (TMRM for 5-10 s, Rho 123 for 60 s) led to a significant (29.6 ± 8.2 and 30.2 ± 5.2%) and rapid decrease in mitochondrial NAD(P)H fluorescence
followed by a slow reincrease. Prolonged photoactivation ( Pyridine nucleotides, i.e. NAD(H) and NADP(H), play a
central role in metabolism; they are the most important coenzymes
acting as hydride (hydrogen anion) donors of various cellular
dehydrogenases (e.g. glutathione reductase), functioning as
reducing/oxidizing equivalents in essential reactions such as energy
supply (aerobic or anaerobic) and photosynthesis, and are required for
DNA repair.
The ability of an organism to counteract reactive oxygen species
(ROS)1 or reactive nitrogen
species depends on its antioxidative capabilities, which involves
destroying of both pro-oxidants (e.g. ROOH,
H2O2, ONOOH) and oxidants (e.g.
radicals and reactive intermediates like singlet oxygen,
1O2). Whereas pro-oxidants are typically
degraded by enzymes (e.g. catalase, glutathione peroxidase,
and superoxide dismutase), oxidants are scavenged by relatively small
biomolecules (e.g. ascorbic acid, glutathione, and
The capability of NAD(P)H to additionally act as a directly operating
antioxidant, i.e. to donate only one electron, was sharply underestimated by various biochemical researchers, a fact that is
probably because of the observation that a biochemical standard one-electron oxidant, [Fe(CN)6]315 s) of
TMRM-loaded cells resulted in even stronger NAD(P)H oxidation, the
rapid onset of mitochondrial permeability transition, and apoptotic
cell death. These results demonstrate that NAD(P)H is the primary
target for 1O2 in hepatocyte mitochondria. Thus
NAD(P)H may operate directly as an intracellular antioxidant, as long
as it is regenerated. At cell-injurious concentrations of the oxidant,
however, NAD(P)H depletion may be the event that triggers cell death.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES
-tocopherol); these are termed directly operating antioxidants. In
this context, NAD(P)H is crucial to maintaining the cellular redox
state and/or antioxidative capacity, because of its essential role as a
coenzyme in the enzymatic re-reduction of directly operating
antioxidants (1, 2). Consequently, NAD(P)H deficiencies are linked with
an increased sensitivity to oxidative stress (2, 3).
, oxidizes
NADH only very slowly (4). However, we recently demonstrated that, in
line with the Marcus theory of electron transfer (1, 5), the reaction
constant of Reaction 1
correlated well with the reduction potential of the
oxidizing radical (1). Consequently, putative harmful radicals
(ROO·, RO·, CO3·
) react
very fast with NADH (kr = 108-109 M
1
s
1). The NAD· radical thus formed reacts with
molecular oxygen near to the diffusion-controlled limit, thereby
yielding NAD+ and superoxide, shown in Reaction 2.
In chemical systems, O
In biological systems superoxide dismutase (SOD) catalyzes
the dismutation of O
The H2O2-consuming enzymes catalase and
glutathione peroxidase (GPx) strongly limit the noxious action of
H2O2, shown in Reactions 5 and 6.
Given the high concentrations of NADH and NADPH and also the high
activity of both superoxide dismutase and glutathione peroxidase in
mitochondria, the reduced coenzymes are expected to act as directly
operating antioxidants in these organelles (1).
Besides oxidizing radicals, the reactive intermediate
1O2 also rapidly reacts with both NADH and
NADPH (kr = 4.3 × 107
M1 s
1 and 8.4 × 107 M
1 s
1) via
single electron transfer (7, 8), shown in Reactions 7 and 8.
![]() |
![]() |
![]() |
![]() |
In most cell types, the highest concentrations of reduced nicotinamides
are located within the matrix space of mitochondria (12). Taking this
into consideration, along with the kinetic data on reactions of
different ROS with NAD(P)H in comparison with other biomolecules,
1O2 can be expected to be most effective, and
most selective, in oxidizing mitochondrial NAD(P)H. We therefore
studied the effect of 1O2 on the NAD(P)H redox
state within the exceptional NAD(P)H-rich mitochondrial matrix space of
cultured hepatocytes (12, 13). To perform these studies, we established
a system based on different rhodamine derivatives and on digital
fluorescence microscopy to selectively generate
1O2 in close proximity to this NAD(P)H pool and
to record the effect on mitochondrial NAD(P)H fluorescence with high
temporal resolution.
![]() |
EXPERIMENTAL PROCEDURES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Chemicals
Leibovitz L-15 medium was obtained from Invitrogen;
collagenase, collagen (Type R), dexamethasone, and gentamicin were from Serva (Heidelberg, Germany); and KCN and Me2SO were
from Merck (Darmstadt, Germany). Bovine serum albumin came from
Behring Institute (Mannheim, Germany), and the following chemicals were
from Sigma: fetal calf serum, superoxide dismutase, NADP-linked
isocitric dehydrogenase, 1,3-bis(chloroethyl)-1-nitrosourea
(BCNU), -hydroxybutyric acid, acetoacetic acid, carbonyl cyanide
m-chlorophenylhydrazone,
-D-fructose,
glutathione (reduced) ethyl ester, dl-isocitric acid, NADH,
NADPH, tert-butyl hydroperoxide (t-BuOOH),
trifluoperazine, and propidium iodide. Chelex (chelating resin;
iminodiacetic acid), 1,3-diphenylisobenzofuran, and
9,10-diphenylanthracene were obtained from Sigma-Aldrich, and digitonin
was from Fluka. The fluorescent dyes tetramethylrhodamine methyl ester
(TMRM), 2',4',5',7'-tetrabromorhodamine 123 bromide (TBRB), rhodamine
(Rho) 123, and calcein-acetoxymethylester were purchased from
Molecular Probes Europe B.V. (Leiden, The Netherlands). Falcon 6-well
cell culture plates were obtained from BD Biosciences, and glass
coverslips were from Assistent (Sondheim/Röhn, Germany).
Animals
Male Wistar rats (200-350 g) were obtained from the Zentrales Tierlaboratorium (Universitätsklinikum Essen). Animals were kept under standard conditions with free access to food and water. All animals received humane care in compliance with the institutional guidelines.
Cell Culture
Hepatocytes were isolated from male Wistar rats as described previously (14). For the fluorescence measurements 1.7 × 105 cells/cm2 were seeded onto collagen-coated 6.2-cm2 glass coverslips in 6-well cell culture plates. Cells were cultured in L-15 medium supplemented with 5% fetal calf serum, L-glutamine (2.0 mM), glucose (8.3 mM), bovine serum albumin (0.1%), NaHCO3 (14.3 mM), gentamicin (50 mg/liter), and dexamethasone (1.0 µM) at 37 °C in a 100% humidified atmosphere of 5% CO2/21% O2/74% N2. Two h after seeding, adherent cells were washed three times with Hanks' balanced salt solution (HBSS; 137.0 mM NaCl/5.4 mM KCl/1.0 mM CaCl2/0.5 mM MgCl2/0.4 mM KH2PO4/0.4 mM MgSO4/0.3 mM Na2HPO4/25.0 mM Hepes, pH 7.4) and supplied with fresh medium as reported previously (15).
Experiments in a Cell-free System
Generation and Detection of 1O2--
The
1O2 detector molecules
1,3-diphenylisobenzofuran and 9,10-diphenylanthracene (each 5 µM; stock solutions 10 mM in
Me2SO) were added to HBSS (3.0 ml, 25 °C) and
transferred into the quartz cuvette of a spectrofluorometer (RF-1501;
Shimadzu, Kyoto, Japan). After recording the baseline fluorescence of
the detector molecules (1,3-diphenylisobenzofuran
exc. = 409 nm,
em. = 476 nm;
9,10-diphenylanthracene
exc. = 391 nm,
em. = 405 nm) for 5 min at 60-s intervals, TMRM (10 µM), Rho 123, (10 µM) or TBRB (10 µM) were added from concentrated stock solutions (10 mM in Me2SO), and the fluorescence of the 1O2 detector molecules was recorded for a
further 5 min. Afterward, the samples were transferred into a modified
Pentz chamber (diameter, 24 mm) placed on the microscope stage
(37 °C) of an inverted microscope; a second sample treated the same
way up to that point was kept in the dark and served as a control. To
photoactivate the different rhodamine derivatives (TMRM
exc. = 535 ± 17.5 nm; Rho 123
exc. = 488 ± 10 nm; TBRB
exc. = 535 ± 17.5 nm),
the 100-watt mercury short arc photo optic lamp (HBO 100; Osram,
Göttingen, Germany) of a digital fluorescence microscope
(Axiovert 135 TV; Zeiss, Oberkochen, Germany) equipped with the
Attofluor imaging system (Atto Instruments, Rockville, MD) was used. To
allow effective irradiation of the whole sample volume, the objective
(×63 numerical aperture 1.25 Plan-Neofluar; Zeiss,
Göttingen) of the microscope was removed, and the irradiation
period was set at 10 min; except for this modification, the same
conditions were used to photoactivate the dyes in the cell-free system
as those used in experiments with cells (see below). After this
treatment, the samples were again transferred to the cuvette of the
spectrofluorometer, and the fluorescence intensity of the
1O2 detector molecules was compared with that
of the untreated controls.
Determination of the Effect of 1O2 on
NAD(P)H and Scavenging of 1O2 by NADPH--
In
other experiments the rhodamine derivatives were photoactivated in the
presence of NADPH (20 µM; stock solution 2.0 mM in HBSS), or the 1O2 detector
molecules were replaced by NADH or NADPH (20 µM), and
MgCl2 (5.0 mM) was added to the reaction buffer
(HBSS, 25 °C). NADPH fluorescence intensity was detected
spectrofluorometrically (exc. = 340 nm;
em. = 460 nm) before and after photoactivation of the
rhodamine derivatives (see above). To determine the amount of
NADP+ formed, dl-isocitric acid (4.0 mM) and NADP-linked isocitric dehydrogenase (0.21 units/ml)
were added to the reaction buffer (HBSS, 37 °C) subsequent to the
irradiation procedures. The increase in fluorescence
(
exc. = 340 nm;
em. = 460 nm) of the
irradiated mixture indicating enzymatic re-reduction of
NADP+ to NADPH was recorded spectrofluorometrically (11).
Further experiments were performed in the presence of either
superoxide dismutase (100 units/ml) or various HBSS/D2O
ratios. Alternatively, experiments were performed with HBSS that had
been treated with chelex (15, 16) to minimize the transition metal contamination.
Experiments with Cultured Hepatocytes
Determination of Cellular NAD(P)H Fluorescence and Photoactivation of Intracellular Rhodamines-- Experiments with hepatocytes were started 20-24 h after isolation of the cells. The glass coverslips with adherent cells were transferred to a modified Pentz chamber, and cells were washed twice with warm (37 °C) HBSS. Hepatocytes were incubated with TMRM (0.5 µM), Rho 123 (0.5 or 10.0 µM), or TBRB (2.0 µM; stock solutions: 1.0 or 2.0 or 10.0 mM in Me2SO) for 20 min in L-15 cell culture medium (37 °C) and then washed three times with HBSS. Afterward, the hepatocytes thus loaded were incubated for another 15 min in dye-free L-15 medium; this incubation period has been found previously to strongly improve the selectivity of the mitochondrial loading with TMRM and Rho 123 (17, 18). The medium was then exchanged, and hepatocytes were covered again with complete L-15 cell culture medium (37 °C) to maintain optimal nutrition of the cells during the experiments. The presence of culture medium did not add significant background to the autofluorescence images at the setting used in this study.
A digital fluorescence microscope was used to measure cellular NAD(P)H fluorescence (see above). Measurements were performed at 37 °C using an excitation filter of 365 ± 12.5 nm and monitoring the emission at 450-490 nm using a bandpass filter. During the measurements cells were flushed with either 5% CO2/21% O2/74% N2 or 5% CO2/95% N2 (in air-tight chambers) to induce hypoxia. Cellular NAD(P)H fluorescence was recorded at 120-s intervals with an excitation period of 0.3 s and the intensity of the mercury lamp attenuated 99% using gray filters to minimize photochemical effects. Single cell fluorescence was determined by confining the regions of interest manually to individual cells. After establishing NAD(P)H baseline fluorescence (6-10 min), the intracellular rhodamine derivatives were photoactivated for 1-60 s at the wavelengths cited above, and NAD(P)H fluorescence measurements were continued without delaying the interval for data collection. Rho 123 was excited at either 488 ± 10 nm or 535 ± 17.5 nm as the excitation maximum of this dye has been reported to shift from 507 (19) to 514.5 nm within cells (20, 21).
In some experiments, cultured hepatocytes (in L-15 medium, 37 °C) were preincubated for 1 h with either 300 µM of the glutathione reductase inhibitor BCNU (22, 23) or an ethyl ester of reduced glutathione (4.0 mM) before fluorescence measurements were started (in the presence of these chemicals). All of the further chemicals were added from concentrated stock solutions during NAD(P)H fluorescence measurements at the respective concentrations detailed in the results. None of the chemicals/agents added in this study showed any detectable fluorescence under the conditions applied.
Determination of the Subcellular Distribution of the Different
Rhodamine Derivatives--
A laser scanning microscope (LSM 510;
Zeiss, Oberkochen, Germany) equipped with both argon and helium/neon
lasers was used to study the subcellular distribution of the different
rhodamine derivatives and their effect on mitochondrial integrity after photoactivation. Subcellular distribution of TMRM
(exc. = 543 nm;
em.
560 nm), Rho 123 (
exc. = 488 nm;
em.
505 nm), and of
TBRB (
exc. = 543 nm;
em.
560 nm)
was determined from the subcellular fluorescence of the probes at the
respective wavelengths. The objective lens was a ×63 numerical
aperture 1.40 Plan-Apochromat. The scanning parameters were as follows.
The pinhole was set at 130 µm, producing confocal optical slices of
less than 1.0 µm in thickness. Confocal images (scanning time
3.9 s, zoom factor 0.7 to 2.5) were collected at different
intervals and with different parameters. The power of the helium/neon
laser was set at 1.0%, and that of the argon laser was set at 0.1% to
minimize photochemical damage.
Similar to the experiments based on digital fluorescence microscopy, after establishing the baseline fluorescence (5-10 min), the rhodamine derivatives were photoactivated for 5-60 s using the 100-watt mercury short arc photo optic lamp of the LSM 510 system. In some experiments, hepatocellular autofluorescence was excited at 488 nm with the power of the argon laser set at 10%, collecting fluorescence emission through a 505-nm long pass filter. Image processing and evaluation were performed using the "physiology evaluation" software of the LSM 510 imaging system.
Recording of the Mitochondrial Membrane Potential and Detection
of Onset of Mitochondrial Permeability Transition--
Mitochondria
were identified, and their functional integrity was confirmed by
membrane potential-dependent staining with TMRM, using
either digital fluorescence microscopy or laser scanning microscopy.
Hepatocytes were incubated with TMRM (0.5 µM) as
described above. When digital fluorescence microscopy was used,
intracellular TMRM fluorescence (exc. = 535 ± 17.5 nm;
em.
590 nm) was recorded at 120-s intervals with
the intensity of the mercury lamp attenuated 40% using gray filters to
minimize photochemical effects; using laser scanning microscopy,
mitochondrial TMRM fluorescence (
exc. = 543 nm;
em.
560 nm) was scanned at different intervals as given above. In some experiments hepatocytes were incubated
simultaneously with TMRM (0.5 µM) and Rho 123 (0.5 µM). In experiments with double-stained mitochondria, red
fluorescence of TMRM (
exc. = 543 nm;
em.
585 nm) and green fluorescence of Rho 123 (
exc. = 488 nm;
em. = 505-530 nm) were optically isolated in
successive scans.
The onset of mitochondrial permeability transition (MPT) was detected
according to the procedure described in Ref 24, with slight
modifications. Briefly, cells were loaded simultaneously with
calcein-AM (1.0 µM) and TMRM (0.5 µM) as
described above for the loading with TMRM alone and then washed three
times with HBSS and covered again with L-15 cell culture medium (for 15 min) that contained propidium iodide (5 µg/ml) but not TMRM (100 nM) as originally reported (24). This incubation period and
the following experiments were performed in the absence of any TMRM within the supernatant to make sure that the probe was located exclusively within the mitochondrial matrix of the cells (see above).
Using laser scanning microscopy, red fluorescence of TMRM (exc. = 543 nm;
em.
585 nm) and green
fluorescence of calcein (
exc. = 488 nm;
em. = 505-530 nm) were recorded in successive scans.
Loss in mitochondrial TMRM fluorescence and redistribution of cytosolic
calcein fluorescence (into the mitochondrial matrix) were considered as
qualitative measures of a decrease in mitochondrial membrane potential
and an increased permeability of the inner mitochondrial membrane,
respectively, known to indicate the onset of MPT as high conductance
permeability transition pores are opened (24-27).
Cell Viability--
The uptake of the vital dye propidium iodide
(5 µg/ml) was routinely determined either during or at the end of the
experimental procedures to detect loss of cell viability. The red
fluorescence of propidium iodide excited at 543 nm was collected
through a 560-nm long pass filter when laser scanning microscopy was
used; using digital fluorescence microscopy, propidium iodide was
detected at exc. = 535 ± 17.5 nm and
em.
590 nm.
Statistics--
All experiments with hepatocytes were repeated
at least three times using cells from different animals, and
experiments in a cell-free system were repeated at least twice.
Cellular microfluorographs and traces shown in the figures are
representative of all the corresponding experiments performed. The
results are expressed as means ± S.D. or S.E.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Oxidation of NAD(P)H by 1O2 in a Cell-free System-- Before starting with the cellular measurements, we studied in a cell-free system whether photoactivation of the different rhodamine derivatives (TMRM, Rho 123, and TBRB) intended to be used for intramitochondrial generation of 1O2 did in fact generate sufficient 1O2. Additionally, we tested whether NAD(P)H, when reacting with this ROS, underwent significant oxidation to enzymatically active NAD(P)+ as reported previously (8, 10, 11).
When the known (20, 21, 28) 1O2 generators TBRB
and Rho 123 (10 µM) were photoactivated, the fluorescence
of both 1O2 detector molecules,
1,3-diphenylisobenzofuran (5 µM) and
9,10-diphenylanthracene (5 µM), was markedly quenched
(data not shown). Very surprisingly, TMRM, for which
1O2 generation has not yet been quantified, was
even more effective than Rho 123, presumably because of the small
1O2 quantum yield of the latter (20, 28). Using
TMRM, the fluorescence of 1,3-diphenylisobenzofuran was quenched more
strongly (54.5 ± 3.0%) than that of 9,10-diphenylanthracene
(14.1 ± 1.0%), in line with their rate constants for single
electron transfer to 1O2
(kr 1.0 × 109
M
1 s
1, and
kr
1.0 × 106
M
1 s
1, respectively; (29)). In
controls, in which the rhodamine derivatives were not photoactivated,
or the samples were irradiated in the absence of the
1O2 generators, no quenching of the detector
molecules became apparent. To confirm the conclusion that TMRM is
highly effective in generating 1O2, the
fluorescence quenching of 1,3-diphenylisobenzofuran and 9,10-diphenylanthracene was performed in the presence of
D2O, which is known to increase the lifetime and thus the
steady state level of 1O2 severalfold (30, 31).
In line with our view, the fluorescence quenching of the
1O2 detector molecules was enhanced 2-3-fold
in the presence of D2O (data not shown). In summary, the
data presented here clearly demonstrated that photoactivation of all
rhodamines resulted in the generation of
1O2.
When the 1O2 detector molecules were replaced
by NADPH (20 µM), its fluorescence significantly
decreased after photoactivation of the selected rhodamines (each 10 µM; see Table I). Similar to the experiments performed with 1,3-diphenylisobenzofuran and 9,10-diphenylanthracene, NAD(P)H fluorescence decreased more strongly (50-80%) in the presence of D2O (data not shown). Again,
the strongest decrease in fluorescence was observed with TBRB as a
1O2 generator. The decrease in NADPH
fluorescence was found to be independent of the presence of either
superoxide dismutase or contaminant transition metal ions (Table I).
The latter possibility was excluded by treating the reaction solution
with chelex. Thus, the fluorescence of NADPH was neither affected by
O
|
If NAD(P)H were a primary target of 1O2, this would prevent, or partially prevent, the oxidation of other molecules targeted by 1O2. In fact, when 1O2 was generated by photoactivation of TMRM, NADPH (20.0 µM), but not NADP+, significantly (22.4 ± 3.0%) and almost completely (98.6 ± 0.5%) diminished the decrease in fluorescence of both 1,3-diphenylisobenzofuran and 9,10-diphenylanthracene (5.0 µM), respectively. In summary, in the cell-free system, 1O2 as generated by photoactivation of Rho 123, TMRM, or TBRB, respectively, mainly oxidized NAD(P)H to enzymatically active NAD(P)+.
Oxidation of NAD(P)H by 1O2 in Mitochondria
of Hepatocytes--
When primary cultured hepatocytes were loaded with
the rhodamine derivatives, the intracellular fluorescence of TMRM
(exc. = 543 nm,
em.
560 nm; see below)
and Rho 123 (
exc. = 488 nm,
em.
505 nm), detected using laser scanning microscopy, was detectable entirely
within intact mitochondria, whereas TBRB (
exc. = 543 nm,
em. =
560 nm), which was hardly taken up by the cells even at higher concentrations (2.0 µM), was mainly
located within the lysosomes/endosomes and the cytosol of the cells
(data not shown). Under these conditions, none of the rhodamine
derivatives affected either cell viability (as detected by propidium
iodide uptake) or mitochondrial functionality as assessed by recording the mitochondrial membrane potential.
Hepatocellular autofluorescence as excited at exc. = 365 ± 12.5 nm and detected at
em. = 450-490 nm
using digital fluorescence microscopy has been considered to be almost
exclusively represented by the fluorescence of NAD(P)H (32, 33) and was
found to be well co-localized with TMRM and Rho 123 here (data not
shown). These results are in line with previous studies where reduced pyridine nucleotides, as well as TMRM and Rho 123, were found to be
almost exclusively located within the mitochondria of cultured rat
hepatocytes (17, 18, 24-26). As the intramitochondrial concentration
of NADH has been reported to be significantly smaller than that of
NADPH in hepatocytes (12), we considered the dominant fluorophore under
investigation here to be NADPH. The assumption that the hepatocellular
autofluorescence at these settings was largely represented by
mitochondrial NAD(P)H and not by other cellular fluorophores was
further supported by the observation that the addition of KCN (5 mM), an inhibitor of the respiratory chain, markedly
(14.5 ± 2.5%) and rapidly increased cellular autofluorescence, which, on the other hand, was decreased by 37.5 ± 5.3% when
oxidative phosphorylation was uncoupled from respiration with carbonyl
cyanide m-chlorophenylhydrazone (10 µM; data
not shown). These changes in cellular autofluorescence exhibited the
same tendencies as were observed in other studies with cultured
hepatocytes (26).
When hepatocytes loaded with 0.5 µM TMRM or Rho 123 were
continuously irradiated (TMRM exc. = 535 ± 17.5 nm; Rho 123
exc. = 488 ± 10 or 535 ± 17.5 nm) for 5-60 s using the 100-watt mercury short arc photo optic lamp
of the inverted microscope, a rapid decrease in NAD(P)H fluorescence
depending on the time of photoactivation was observed in TMRM-loaded
cells, whereas only a slight decrease in NAD(P)H fluorescence was
evident in cells loaded with Rho 123 (Fig.
1A). However, when the cells
were loaded with 10.0 µM Rho 123, i.e. with a
concentration as previously used in studies of photodynamic therapy
with different types of tumor cells and animal models (20, 21, 34-36),
photoactivation of intramitochondrial Rho 123 provided essentially the
same effect on cellular NAD(P)H fluorescence as TMRM. In cells loaded
with TBRB (2.0 µM), no decrease in fluorescence showed
through even after prolonged (60-s) photoactivation (
exc. = 535 ± 17.5 nm) of the dye. This result is
in apparent contrast to the strong oxidation of NAD(P)H after
photoactivation of TBRB in the cell-free system (Table I), but it is a
good reflection of the fact that TBRB is not co-localized with the
mitochondrial NAD(P)H pool. In line with the stronger oxidation of
NAD(P)H after photoactivation of TMRM in the cell-free system, TMRM
also decreased NAD(P)H fluorescence in mitochondria noticeably more
strongly than Rho 123 under comparable conditions (Fig. 1A).
In controls, the intensity of mitochondrial NAD(P)H fluorescence was
not affected by mitochondrial loading with either TMRM or Rho 123, and
in hepatocytes, which were not loaded with the dyes, no decrease in
NAD(P)H fluorescence was observed after photoactivation. In contrast to
the experiments performed in the cell-free system, the decrease in
cellular NAD(P)H fluorescence was not intensified when
intramitochondrial TMRM was excited in the presence of
D2O-enriched L-15 medium (data not shown). As
photoactivation of TMRM most effectively decreased mitochondrial
NAD(P)H fluorescence intensity (29.6 ± 8.2% after 10 s of
irradiation; Rho 123: 30.2 ± 5.2% after 60 s of
irradiation), most of the following experiments with hepatocytes were
performed using TMRM.
|
As known from studies of photodynamic therapy (PDT),
1O2 can lead to a marked oxidation of proteins
and membrane lipids, resulting in leakage of small biomolecules from
the damaged cells/cellular compartments. In line with this,
photoactivation of TMRM has been reported to result in generation of
free radicals (37, 38) leading to a gradual and reversible decline in
membrane potential of isolated individual rat heart mitochondria
because of repetitive opening and closing of the mitochondrial
transition pore (37). Therefore, to exclude the possibility that the
observed decrease in NAD(P)H fluorescence was a result of mitochondrial
NAD(P)H leakage, we studied the capability of rhodamines of inducing
MPT. When intramitochondrial TMRM or Rho 123 (after loading with 10 µM) were irradiated for 10 and
60 s, respectively, in
most experiments the initial decrease in NAD(P)H fluorescence was
followed by a fluorescence increase, suggesting regulatory re-reduction
of the intramitochondrial oxidized nicotinamides (Fig. 1). However,
such increases were not observed after prolonged photoactivation, which resulted in a rapid decrease of NAD(P)H and TMRM/Rho 123 fluorescence to almost background levels and subsequently in cell death as indicated
by the uptake of propidium iodide (Fig.
2; not shown for Rho 123). Interestingly,
when the deposited light dose was lethal, NAD(P)H fluorescence rapidly
increased in some of the cells immediately before mitochondrial
membrane potential completely dropped (Fig. 2A). In contrast
to the cells loaded with TMRM (0.5 µM), no cytotoxic
effects became apparent in experiments with Rho 123 (0.5 µM) and TBRB (2.0 µM), even after prolonged
(60-s) photoactivation. The reincrease in NAD(P)H fluorescence after short term photoactivation of TMRM and Rho 123 (Fig. 1B)
already strongly suggested that an opening of the mitochondrial
permeability transition pore was not responsible for the initial
decrease in NAD(P)H fluorescence. This conclusion was supported by
further findings. First, when the mitochondrial membrane potential of the hepatocytes was recorded with TMRM, and the probe was excited for
10 s, mitochondrial TMRM fluorescence decreased in the same manner as
the fluorescence of NAD(P)H but was rapidly restored when TMRM (0.5 µM) was added to the supernatant (data not shown). This
indicated that the mitochondrial membrane potential, i.e. the driving force for TMRM uptake, was still intact. The decrease in
mitochondrial TMRM fluorescence intensity was found to result primarily
from partial photodegradation of the dye (data not shown; see below).
Second, the onset of MPT was safely excluded using high resolution
laser scanning microscopy. After short term photoactivation the
cytosolic dye calcein did not diffuse into the mitochondrial compartment although TMRM fluorescence slightly decreased (Fig. 3). Third, neither TMRM nor NAD(P)H was
detectable within the cytosol, and the decrease in NAD(P)H fluorescence
was not prevented by trifluoperazine (5.0 µM)/fructose
(10 mM), known to effectively inhibit MPT (data not shown)
(26). The inability of trifluoperazine/fructose to inhibit the
initial decrease in NAD(P)H fluorescence further suggests that ROS
other than 1O2, large amounts of which may be
generated during the onset of MPT leading to NAD(P)H oxidation (26,
39), were not responsible for the decrease in NAD(P)H fluorescence
observed here. In summary, the decrease in mitochondrial NAD(P)H
fluorescence upon short term photoactivation of intramitochondrial TMRM
and Rho 123 did not result from mitochondrial damage but from NAD(P)H
oxidation. In contrast to the short term photoactivation, prolonged
(
15 s) photoactivation of TMRM caused a rapid decrease in membrane potential, onset of MPT within minutes, and subsequently apoptotic cell
death (Fig. 4).
|
|
|
A major protective effect of NADPH is associated with its role as a
coenzyme for the glutathione peroxidase/reductase system, which, for
purposes of the present study, ought to be the only enzymatic system
that could possibly be involved in 1O2-induced
NAD(P)H oxidation. To identify whether the mitochondrial NAD(P)H pool
was directly oxidized by 1O2 or whether its
oxidation resulted from the regeneration of glutathione, we studied the
role of the cytosolic/mitochondrial glutathione peroxidase/reductase
system in TMRM-induced NAD(P)H oxidation by inhibiting glutathione
reductase with BCNU (300 µM, 1 h preincubation). The
pretreatment of cultured hepatocytes with BCNU, however, had no
inhibiting effect on 1O2-mediated NAD(P)H
oxidation (Fig. 5). To verify whether the enzyme was indeed inhibited by BCNU, control experiments with t-BuOOH were performed, which rapidly leads to glutathione
reductase-catalyzed oxidation of NADPH and, via pyridine nucleotide
transhydrogenase, of NADH, too (13, 26, 27, 40). As expected, and in
contrast to the TMRM-mediated 1O2 generation,
t-BuOOH had no effect on mitochondrial NAD(P)H fluorescence
when the cells had been pretreated with BCNU (Fig. 5). Thus, the
applied BCNU (300 µM) did indeed completely inhibit glutathione reductase. The delayed (6-14-min) decrease in NAD(P)H fluorescence was attributed to a loss in mitochondrial membrane potential, onset of MPT, and cell death in line with the well known
fact that BCNU treatment sensitizes cells to oxidative stress (22, 41).
These results clearly demonstrated that glutathione reductase was not
involved in the 1O2-derived oxidation of
NAD(P)H. In line with this, an increase in the hepatocellular reduced
glutathione concentration, achieved experimentally by incubating the
cells with an ethyl ester of reduced glutathione (4 mM, for
1 h), had no effect on the mitochondrial NAD(P)H oxidation after
photoactivation of TMRM (data not shown), clearly demonstrating that
reduced glutathione, even at higher concentrations, cannot compete with
NAD(P)H for 1O2. Therefore, and also in view of
the rapidity of the decrease in NAD(P)H fluorescence, it is most
unlikely that NAD(P)H was enzymatically oxidized in our experiments.
This is in line with a study by Kessel and Luo (42) in which
photodamaging effects of intramitochondrial porphycenes were found to
be independent of the (low) ambient temperature and thus of enzymatic
processes.
|
To confirm that the mitochondrial NAD(P)H was oxidized by
1O2 and not directly by the photoactivated
rhodamine derivatives, we studied the influence of the environmental
pO2 on intramitochondrial NAD(P)H oxidation. When the cells
were flushed with 95% N2/5% CO2 for 20 min,
hypoxia, as indicated by a slight increase in NAD(P)H fluorescence,
completely prevented the decrease in NAD(P)H fluorescence after
photoactivation of TMRM (Fig. 6). This
strongly suggested that NAD(P)H was oxidized under normoxia by (most
likely) 1O2, the main ROS generated in
photochemical processes, rather than by products (radicals, radical
ions) of the photochemically activated process or any photochemical
activation of TMRM itself in a type-1 photoreaction. As expected,
hypoxic cells were found to resist photoactivation of TMRM. In contrast
to normoxic conditions no loss in cell viability was observed even
after prolonged photoactivation (data not shown). Rather than
completely preventing NAD(P)H oxidation, hypoxia only partly prevented
the decrease in intramitochondrial TMRM fluorescence (Fig. 6) (see
above); it follows that this probably resulted from both uncoupled,
i.e. 1O2-independent, and coupled
photobleaching of the dye. The fact that the decrease in TMRM
fluorescence showed a relatively weak dependence on the environmental
pO2 further suggested that the indicator molecule itself
did not react with 1O2 very well, which
possibly explains its high oxidizing effect on NAD(P)H.
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
NAD(P)H, the Primary and Restorable Target of
1O2 in Mitochondria of Viable
Cells--
During the past 25 years, a good deal of thermodynamically
and experimentally based data have been reported concerning the rapid
reaction of 1O2 with NAD(P)H (see Reactions 7 and 8). However, the resulting consequences for intracellular
conditions have hardly been considered. The rate constants for single
electron transfer (kr) from NADH or NADPH to
1O2 are significantly higher (4.3 × 107 and 8.4 × 107
M1 s
1) than the
kr values for the well known directly operating
antioxidants ascorbate (8.3 × 106
M
1 s
1), glutathione (2.4 × 106 M
1 s
1), and
-tocopherol (5.0 × 106
M
1 s
1) (8)). When the
respective intramitochondrial concentrations of these biomolecules are
taken into consideration for hepatocytes (NADH: 4.0 mM,
NADPH: 6.0 mM, glutathione: 10.0 mM (43);
ascorbate: 0.1-0.5 mM (44);
-tocopherol: 0.05-2.28
nmol/mg protein (45, 46) (
10-450 µM)), NADH can be
expected, and NADPH even more so, to be the primary targets of
1O2 within the mitochondrial matrix of this
cell; the concentrations given were calculated in part from the
mitochondrial content of each compound, assuming that about 7.2 × 109 rat liver mitochondria contain 1 mg of protein, and the
volume of a single mitochondrion is 0.71 µm3 (12).
In line with this assumption, selective generation of moderate amounts
of 1O2 within the mitochondrial matrix space of
cultured hepatocytes by local photoactivation of TMRM and Rho 123 led
to a rapid oxidation of mitochondrial NAD(P)H followed by obviously
enzymatic re-reduction of NAD(P)+ (Fig. 1, A and
B). Prolonged photoactivation of TMRM further increased
NAD(P)H oxidation and resulted in a rapid decrease in mitochondrial
membrane potential (37, 38), the onset of MPT, loss of mitochondrial
NAD(P)H, and finally apoptotic cell death (see Figs. 2 and 4). In
controls, NAD(P)H oxidation in the cell-free system was found to be
independent of O
The strong oxidation of NAD(P)H after photoactivation of TMRM and Rho 123 indicates that 1O2 was generated very close to the mitochondrial NAD(P)H molecules; 1O2 can diffuse only 10-100 nm during its lifetime in the cell, which is much shorter (0.01-0.04 µs) than in simple aqueous solutions (2-4 µs) (8, 11, 28, 47-49). This short lifetime is consistent with the lack of an intracellular D2O effect as found here and in many other photosensitized processes in which 1O2 was very likely to be involved (48).
Whether the generation of 1O2 predominantly
leads to (NAD(P))2 or NAD(P)+ ought mainly to
depend on the local oxygen tension (7, 8, 10, 11). In the present
study, experiments in the cell-free system demonstrated that
photoactivation of the different rhodamine derivatives yielded 80%
enzymatically active NADP+ (Table I), indicating that
dimerization of NAD(P)· is too slow under normoxic conditions
and that oxidation mainly occurs at the C-4 position of the
nicotinamide ring as reported previously (7, 8, 11). Under decreased
oxygen tensions, however, the
1O2-dependent yield of
enzymatically active NADP+ was reported to be only 40% in
a cell-free system (10). In line with these considerations, after
1O2 generation the cells were unable to fully
restore their NAD(P)H levels (Fig. 1), most likely because the very
high concentration of NAD(P)H and the low pO2 present
within this compartment enhances the yield of
(NAD(P))2.
The role of NAD(P)H as a Directly Operating Antioxidant-- In the cell-free system NADPH significantly diminished the reaction of 1O2 with both 1,3-diphenylisobenzofuran and 9,10-diphenylanthracene. This is in line with a previous study where NADPH protected NADP-linked isocitric dehydrogenase against photochemically generated 1O2, thus allowing the enzyme to regenerate the NADPH responsible for its own protection (11). These results obtained from experiments in cell-free systems clearly indicate that NAD(P)H has the potential to directly protect targets against attack by 1O2 and would therefore be expected to play a role as a directly operating antioxidant in living cells, as well.
It also follows that the ability of a mitochondrion to resist
1O2 ought to depend on its NAD(P)H
concentration, as well as on its capability to re-reduce oxidized
nicotinamides. However, it is very problematic to experimentally
manipulate the well regulated mitochondrial NAD(P)H levels in living
cells without affecting the basic cell metabolism, metabolic
compartmentation, or cell viability. Because of this experimental
limitation it is not possible to differentiate unequivocally between
the indirect and the direct antioxidative and protective effect of
reduced pyridine nucleotides. For instance, in preliminary studies, it
was only possible to slightly (5.2-7.8%) increase the mitochondrial
NAD(P)H concentration using -hydroxybuturic acid (10 mM), whereas hardly any decrease in NAD(P)H fluorescence
was detectable when acetoacetic acid (10 mM) was added to
the supernatant. Consequently, none of these substrates provided either
significant protection of cultured hepatocytes or diminished
mitochondrial integrity/cell viability when 1O2
was generated during photoactivation of intracellular TMRM. Despite the
limitation that any protective effect offered by mitochondrial NAD(P)H
against 1O2 is very difficult to demonstrate
experimentally, the fact that mitochondrial NAD(P)H is the primary and
restorable target of 1O2 (see above) leaves
almost no doubt that NAD(P)H acts as a directly operating antioxidant
in this compartment. As a directly operating antioxidant, NAD(P)H is
likely to act collectively and on a concerted basis with the cellular
enzymes superoxide dismutase, catalase, and glutathione peroxidase,
which can degrade the O
Mitochondrial NAD(P)H Depletion as a Decisive Trigger of Apoptotic
Cell Death--
Besides being a major site of intracellular generation
of reactive oxygen species (O
In the present study, TMRM was found to rapidly induce onset of MPT in cultured rat hepatocytes followed by apoptotic cell death in less than 30 min when the photoactivation periods of the probe were prolonged (see Figs. 2 and 4). Given the rapidity of apoptotic cell death observed here and in other studies (42, 49, 52, 53), it seems rather unlikely that any intermediate steps of biosynthesis and signal transduction pathways were required (49).
In this context, the finding that NAD(P)H is a primary target of
1O2 in living cells is likely to be of major
importance for the general understanding of the photochemotherapeutic
potential of photosensitizing molecules. When small amounts of
1O2 are generated, NAD(P)H should act as a
directly operating antioxidant thereby terminating the attacking
1O2 molecules (see above). However, when the
amount of 1O2 generated exceeds the capacity of
this antioxidative system, excess oxidation of NAD(P)H probably
actually becomes a trigger for cell damage (see Figs. 2 and 4). Large
amounts of the pro-oxidant O
TMRM is a potentiometric fluorescent probe that is widely used for
several tasks in cell biology and physiology. It serves as a marker for
identifying mitochondria (17) and for recording their membrane
potential (37, 38, 60), for example, and is used in an assay for
detecting onset of MPT (24-27). In the light of the present results,
however, one should keep in mind that TMRM, even when excited only for
a short period, most likely affects cellular NAD(P)H homeostasis and
consequently weakens the antioxidative capacity of the cells. This will
be of relevance especially when TMRM, combined with the cytosolic
marker calcein, is used to study the MPT-inducing potential of ROS and
reactive nitrogen species (24-27).
![]() |
CONCLUSIONS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The results presented, obtained from experiments both in the cell-free system and in primary cultured rat hepatocytes, strongly suggest that NAD(P)H is the primary and enzymatically restorable target of 1O2 within mitochondria of viable cells. It follows that mitochondrial NAD(P)H is likely to act as a directly operating antioxidant and thus provides protection when 1O2 is generated within this organelle. However, when the amount of 1O2 generated exceeds the capacity of the NAD(P)H-regenerating systems, one-electron oxidation of NAD(P)H by 1O2 might even be an as-yet unnoticed pathogenetic event responsible for effects (including photodynamic ones) like inhibition of respiration and electron transport, disruption of the mitochondrial electrochemical gradient, oxidation of NAD(P)H-dependent compounds in mitochondria, onset of MPT, and finally apoptosis.
Having regard to the susceptibility of NAD(P)H to one-electron
oxidations when reacting with oxygen-centered species (1) and to the
ubiquitous distribution of NAD(P)H within the cell, it is most likely
that both roles of NAD(P)H, i.e. as a directly operating
antioxidant and as a decisive trigger of cell injury, are also of
relevance in connection with other ROS. One of the most likely
candidates is the carbonate radical
(CO3·), donated from peroxynitrite.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank the following experts in the field of photodynamic therapy for helpful discussions pertaining to the existence of "mitochondrial" photosensitizers: Dr. Sol Kimel, Dr. Roger Ackroyd, Dr. Thomas Dougherty, Dr. David Kessel, Dr. Johan Moan, Dr. Nancy L. Oleinick, Dr. David I. Vernon, Dr. Stan Brown, and Dr. Petras Juzenas. The present investigation would have been impossible without the technical assistance of A. Wensing and E. Heimeshoff.
![]() |
FOOTNOTES |
---|
* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
To whom correspondence should be addressed. Tel.: 49-201-723-4101;
Fax: 49-201-723-5943; E-mail: h.de.groot@uni-essen.de.
Published, JBC Papers in Press, November 13, 2002, DOI 10.1074/jbc.M204230200
2 F. Petrat, S. Pindiur, M. Kirsch, and H. de Groot, submitted for publication.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: ROS, reactive oxygen species; 1O2, singlet oxygen; TMRM, tetramethylrhodamine methyl ester; TBRB, 2',4',5',7'-tetrabromorhodamine 123 bromide; Rho, rhodamine; BCNU, 1,3-bis(chloroethyl)-1-nitrosourea; GSSG, glutathione (oxidized form); t-BuOOH, tert-butyl hydroperoxide; HBSS, Hanks' balanced salt solution; MPT, mitochondrial permeability transition; PDT, photodynamic therapy; kr, rate constant for single electron transfer.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. |
Kirsch, M.,
and de Groot, H.
(2001)
FASEB J.
15,
1569-1574 |
2. |
Leopold, J. A.,
Cap, A.,
Scribner, A. W.,
Stanton, R.,
and Loscalzo, J.
(2001)
FASEB J.
15,
1771-1773 |
3. | Minard, K. I., and McAlister-Henn, L. (2001) Free Radic. Biol. Med. 31, 832-843[CrossRef][Medline] [Order article via Infotrieve] |
4. |
Schellenberg, K. A.,
and Hellerman, L.
(1958)
J. Biol. Chem.
231,
547-556 |
5. |
Kirsch, M.,
and de Groot, H.
(1999)
J. Biol. Chem.
274,
24664-24670 |
6. | Corey, E. J., Mehrotra, M. M., and Khan, A. U. (1987) Biochem. Biophys. Res. Commun. 145, 842-846[Medline] [Order article via Infotrieve] |
7. | Peters, G., and Rodgers, M. A. J. (1980) Biochem. Biophys. Res. Commun. 96, 770-776[Medline] [Order article via Infotrieve] |
8. | Peters, G., and Rodgers, M. A. J. (1981) Biochim. Biophys. Acta 637, 43-52[Medline] [Order article via Infotrieve] |
9. | Koppenol, W. H. (1976) Nature 262, 420-421[Medline] [Order article via Infotrieve] |
10. | Bodaness, R. S., and Chan, P. C. (1977) J. Biol. Chem. 252, 8554-8560[Abstract] |
11. | Bodaness, R. S. (1982) Biochem. Biophys. Res. Commun. 108, 1709-1715[Medline] [Order article via Infotrieve] |
12. | Tyler, D. D. (1992) The Mitochondrion in Health and Disease , VCH Publishers, Inc., New York |
13. | Hoek, J. B., and Rydström, J. (1988) Biochem. J. 254, 1-10[Medline] [Order article via Infotrieve] |
14. | de Groot, H., and Brecht, M. (1991) Biol. Chem. Hoppe-Seyler 372, 35-41[Medline] [Order article via Infotrieve] |
15. | Petrat, F., Rauen, U., and de Groot, H. (1999) Hepatology 29, 1171-1179[Medline] [Order article via Infotrieve] |
16. | Evans, P. J., and Halliwell, B. (1994) Methods Enzymol. 233, 82-92[CrossRef][Medline] [Order article via Infotrieve] |
17. | Petrat, F., de Groot, H., and Rauen, U. (2001) Biochem. J. 356, 61-69[CrossRef][Medline] [Order article via Infotrieve] |
18. | Petrat, F., Weisheit, D., Lensen, M., de Groot, H., Sustmann, R., and Rauen, U. (2002) Biochem. J. 362, 137-147[CrossRef][Medline] [Order article via Infotrieve] |
19. | Haugland, R. P. (1996) Handbook of Fluorescent Probes and Research Products , 9th Ed. , pp. 479-487, Molecular Probes, Inc., Eugene, OR |
20. | Shea, C. R., Chen, N., Wimberly, J., and Hasan, T. (1989) Cancer Res. 49, 3961-3965[Abstract] |
21. | Shea, C. R., Sherwood, M. E., Flotte, T. J., Chen, N., Scholz, M., and Hasan, T. (1990) Cancer Res. 50, 4167-4172[Abstract] |
22. | Adamson, G. M., and Harman, A. W. (1993) Biochem. Pharmacol. 45, 2289-2294[CrossRef][Medline] [Order article via Infotrieve] |
23. | Harbrecht, B. G., Di, Silvio, M., Chough, V., Kim, Y. M., Simmons, R. L., and Billiar, T. R. (1997) Ann. Surg. 225, 76-87[CrossRef][Medline] [Order article via Infotrieve] |
24. | Zahrebelski, G., Nieminen, A.-L., Al-, Ghoul, K., Qian, T., Herman, B., and Lemasters, J. J. (1995) Hepatology 21, 1361-1372[Medline] [Order article via Infotrieve] |
25. | Nieminen, A.-L., Saylor, A. K., Tesfai, S. A., Herman, B., and Lemasters, J. J. (1995) Biochem. J. 307, 99-106[Medline] [Order article via Infotrieve] |
26. |
Nieminen, A.-L.,
Byrne, A. M.,
Herman, B.,
and Lemasters, J. J.
(1997)
Am. J. Physiol.
272,
C1286-C1294 |
27. | Lemasters, J. J., Nieminen, A.-L., Qian, T., Trost, L. C., Elmore, S. P., Nishimura, Y., Crowe, R. A., Cascio, W. E., Bradham, C. A., Brenner, D. A., and Herman, B. (1998) Biochim. Biophys. Acta 1366, 177-196[Medline] [Order article via Infotrieve] |
28. | MacDonald, I. J., and Dougherty, T. J. (2001) J. Porphyrins Phthalocyanines 5, 105-129[CrossRef] |
29. | Ross, A. B., Mallard, W. G., Helman, W. P., Buxton, G. V., Huie, R. E., and Neta, P. (1998) NDRL/NIST Solution Kinetics Database 3.0 , NDRL/NIST, Gaithersburg, MD |
30. | Ogilby, P. R., and Foote, C. S. (1982) J. Am. Chem. Soc. 104, 2069-2070 |
31. | Parker, J. G., and Stanbro, W. D. (1984) J. Photochem. 25, 545-547 |
32. | Richards-Kortum, R. (1996) Annu. Rev. Phys. Chem. 47, 555-606[CrossRef][Medline] [Order article via Infotrieve] |
33. | Pogue, B. W., Pitts, J. D., Mycek, M.-A., Sloboda, R. D., Wilmot, C. M., Brandsema, J. F., and O'Hara, J. A. (2001) Photochem. Photobiol. 74, 817-824[Medline] [Order article via Infotrieve] |
34. | Modica-Napolitano, J. S., and Aprille, J. R. (1987) Cancer Res. 47, 4361-4365[Abstract] |
35. | Nadakavukaren, K. K., Nadakavukaren, J. J., and Chen, L. B. (1985) Cancer Res. 45, 6093-6099[Abstract] |
36. | Shea, C. R., Chen, N., and Hasan, T. (1989) Lasers Surg. Med. 9, 83-89[Medline] [Order article via Infotrieve] |
37. | Hüser, J., and Blatter, L. A. (1999) Biochem. J. 343, 311-317[CrossRef][Medline] [Order article via Infotrieve] |
38. | Diaz, G., Falchi, A. M., Gremo, F., Isola, R., and Diana, A. (2000) FEBS Lett. 475, 218-224[CrossRef][Medline] [Order article via Infotrieve] |
39. | Zamzami, N., and Kroemer, G. (2001) Nature Rev. Mol. Cell. Biol. 2, 67-71[CrossRef][Medline] [Order article via Infotrieve] |
40. | Lee, C. P., and Ernster, L. (1964) Biochim. Biophys. Acta 81, 187-190 |
41. | Snyder, J. W., Alexander, G. M., Ferraro, T. N., Grothusen, J. R., and Farber, J. L. (1993) Chem. Biol. Interact. 88, 209-223[Medline] [Order article via Infotrieve] |
42. | Kessel, D., and Luo, Y. (1999) Cell Death Differ. 6, 28-35[CrossRef][Medline] [Order article via Infotrieve] |
43. |
Costantini, P.,
Chernyak, B. V.,
Petronilli, V.,
and Bernardi, P.
(1996)
J. Biol. Chem.
271,
6746-6751 |
44. | Li, X., Cobb, C. E., Hill, K. E., Burk, R. F., and May, J. M. (2001) Arch. Biochem. Biophys. 387, 143-153[CrossRef][Medline] [Order article via Infotrieve] |
45. | Ferreira, F. M. L., Palmeira, C. M., Matos, M. J., Seica, R., and Santos, M. S. (1999) Life Sci. 65, 1013-1025[CrossRef][Medline] [Order article via Infotrieve] |
46. |
Lass, A.,
and Sohal, R. S.
(2000)
FASEB J.
14,
87-94 |
47. | Moan, J. (1990) J. Photochem. Photobiol. B Biol. 6, 343-344[CrossRef] |
48. | Moan, J., and Berg, K. (1991) Photochem. Photobiol. 53, 549-553[Medline] [Order article via Infotrieve] |
49. |
Dougherty, T. J.,
Gomer, C. J.,
Henderson, B. W.,
Jori, G.,
Kessel, D.,
Korbelik, M.,
Moan, J.,
and Peng, Q.
(1998)
J. Natl. Cancer Inst.
90,
889-905 |
50. |
Atamna, H.,
Robinson, C.,
Ingersoll, R.,
Elliott, H.,
and Ames, B. N.
(2001)
FASEB J.
15,
2196-2204 |
51. | Benzi, G., and Moretti, A. (1995) Neurobiol. Aging 16, 661-674[CrossRef][Medline] [Order article via Infotrieve] |
52. | Kessel, D., Luo, Y., Deng, Y., and Chang, C. K. (1997) Photochem. Photobiol. 65, 422-426[Medline] [Order article via Infotrieve] |
53. | Kessel, D., and Luo, Y. (1998) J. Photochem. Photobiol. B Biol. 42, 89-95[CrossRef][Medline] [Order article via Infotrieve] |
54. | Ackroyd, R., Kelty, C., Brown, N., and Reed, M. (2001) Photochem. Photobiol. 74, 656-669[CrossRef][Medline] [Order article via Infotrieve] |
55. | Henderson, B. W., and Dougherty, T. J. (1992) Photochem. Photobiol. 55, 145-157[Medline] [Order article via Infotrieve] |
56. | Peng, Q., Moan, J., and Nesland, J. M. (1996) Ultrastruct. Pathol. 20, 109-129[Medline] [Order article via Infotrieve] |
57. | Oleinick, N. L., and Evans, H. H. (1998) Radiat. Res. 150, S146-S156[Medline] [Order article via Infotrieve] |
58. | Moor, A. C. E. (2000) J. Photochem. Photobiol. B Biol. 57, 1-13[CrossRef][Medline] [Order article via Infotrieve] |
59. |
Pierce, R. H.,
Campbell, J. S.,
Stephenson, A. B.,
Franklin, C. C.,
Chaisson, M.,
Poot, M.,
Kavanagh, T. J.,
Rabinovitch, P. S.,
and Fausto, N.
(2000)
Am. J. Pathol.
157,
221-236 |
60. |
Hüser, J.,
Rechenmacher, C. E.,
and Blatter, L. A.
(1998)
Biophys. J.
74,
2129-2137 |