NAD(P)H, a Primary Target of 1O2 in Mitochondria of Intact Cells*

Frank Petrat, Stanislaw Pindiur, Michael Kirsch, and Herbert de GrootDagger

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
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES

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 (>= 15 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

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 alpha -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).

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]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 
<UP>NADH</UP>+<UP>Rad</UP><SUP>⋅</SUP>→<UP>NAD</UP><SUP>⋅</SUP>+<UP>RH</UP>

<UP><SC>Reaction</SC> 1</UP>
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. 
<UP>NAD</UP><SUP>⋅</SUP>+<UP>O<SUB>2</SUB></UP>→<UP>NAD<SUP>+</SUP></UP>+<UP>O</UP><SUB>2</SUB><SUP>&cjs1138;</SUP>

<UP><SC>Reaction</SC> 2</UP>
In chemical systems, O<UP><SUB>2</SUB><SUP>&cjs1138;</SUP></UP> spontaneously dismutates to H2O2 and 1O2 (6), shown in Reaction 3. 
<UP>2O</UP><SUB>2</SUB><SUP>&cjs1138;</SUP>+2<UP>H</UP><SUP>+</SUP>→<UP>H<SUB>2</SUB>O<SUB>2</SUB></UP>+<SUP>1</SUP><UP>O</UP><SUB>2</SUB>

<UP><SC>Reaction</SC> 3</UP>
In biological systems superoxide dismutase (SOD) catalyzes the dismutation of O<UP><SUB>2</SUB><SUP>&cjs1138;</SUP></UP>, thereby preventing the formation of 1O2, shown in Reaction 4. 
<UP>2O</UP><SUB>2</SUB><SUP>&cjs1138;</SUP>+<UP>2H</UP><SUP>+</SUP> <LIM><OP><ARROW>→</ARROW></OP><UL><UP>SOD</UP></UL></LIM><UP> H</UP><SUB>2</SUB><UP>O</UP><SUB>2</SUB>+<SUP>3</SUP><UP>O</UP><SUB>2</SUB>

<UP><SC>Reaction</SC> 4</UP>
The H2O2-consuming enzymes catalase and glutathione peroxidase (GPx) strongly limit the noxious action of H2O2, shown in Reactions 5 and 6. 

<UP><SC>Reaction</SC> 5</UP>

<UP>H</UP><SUB>2</SUB><UP>O</UP><SUB>2</SUB>+2<UP>GSH</UP> <LIM><OP><ARROW>→</ARROW></OP><UL><UP>GPx</UP></UL></LIM><UP> 2H</UP><SUB>2</SUB><UP>O</UP>+<UP>GSSG</UP>

<UP><SC>Reaction</SC> 6</UP>
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 M-1 s-1 and 8.4 × 107 M-1 s-1) via single electron transfer (7, 8), shown in Reactions 7 and 8. 
<UP>NAD</UP>(<UP>P</UP>)<UP>H</UP>+<SUP>1</SUP><UP>O</UP><SUB>2</SUB> → <UP>NAD</UP>(<UP>P</UP>)<SUP>⋅</SUP>+<UP>O</UP><SUB>2</SUB><SUP>&cjs1138;</SUP>+<UP>H</UP><SUP>+</SUP>

<UP><SC>Reaction</SC> 7</UP>

<UP>NAD</UP>(<UP>P</UP>)<SUP>⋅</SUP>+<UP>O</UP><SUB>2</SUB>→<UP>NAD</UP>(<UP>P</UP>)<SUP>+</SUP>+<UP>O</UP><SUB>2</SUB><SUP>&cjs1138;</SUP>

<UP><SC>Reaction</SC> 8</UP>
In 1976 the thermodynamic capability of NAD(P)H to transfer only one electron to 1O2 was estimated by Koppenol (9). Experimental evidence of this reaction in cell-free systems was provided, and consequences of 1O2 generation in mitochondria were hypothesized (7, 8, 10, 11) two decades ago. In biological systems, however, direct, i.e. non-enzymatic, oxidation of NAD(P)H by 1O2 or by any other oxidant has not yet been demonstrated.

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
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES

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), beta -hydroxybutyric acid, acetoacetic acid, carbonyl cyanide m-chlorophenylhydrazone, beta -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 lambda exc. = 409 nm, lambda em. = 476 nm; 9,10-diphenylanthracene lambda exc. = 391 nm, lambda 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 lambda exc. = 535 ± 17.5 nm; Rho 123 lambda exc. = 488 ± 10 nm; TBRB lambda 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 (lambda exc. = 340 nm; lambda 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 (lambda exc. = 340 nm; lambda 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 (lambda exc. = 543 nm; lambda em. >=  560 nm), Rho 123 (lambda exc. = 488 nm; lambda em. >=  505 nm), and of TBRB (lambda exc. = 543 nm; lambda 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 (lambda exc. = 535 ± 17.5 nm; lambda 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 (lambda exc. = 543 nm; lambda 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 (lambda exc. = 543 nm; lambda em. >=  585 nm) and green fluorescence of Rho 123 (lambda exc. = 488 nm; lambda 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 (lambda exc. = 543 nm; lambda em. >=  585 nm) and green fluorescence of calcein (lambda exc. = 488 nm; lambda 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 lambda exc. = 535 ± 17.5 nm and lambda 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
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES

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 approx  1.0 × 109 M-1 s-1, and kr approx  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<UP><SUB>2</SUB><SUP>&cjs1138;</SUP></UP>, which may arise during 1O2 generation (see Reactions 7 and 8), nor by ·OH, resulting from transition metal-dependent Fenton reactions. In the absence of the 1O2 generators the NAD(P)H fluorescence hardly decreased (2%/h) via autoxidation (data not shown). To verify that NADPH was actually oxidized to its enzymatically active non-fluorescent form (NADP+), we tested whether re-reduction was possible, using the procedure described by Bodaness (11), with slight modifications. When dl-isocitric acid and NADP-linked isocitric dehydrogenase were added to the incubation buffer after photoactivation of the selected rhodamine derivatives, NADPH fluorescence was largely restored within minutes (Table I). These results strongly indicated that NAD(P)H was oxidized by 1O2 via Reactions 7 and 8 as suggested by Peters and Rodgers (7, 8).

                              
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Table I
Oxidation of NADPH to enzymatically active NADP+ by 1O2 generated during irradiation of different rhodamine derivatives
NADPH (20 µM) and the respective rhodamine derivatives (10 µM) Rho 123, TMRM, or TBRB were dissolved in HBSS (25 °C), which additionally contained 5.0 mM MgCl2 or MgCl2 (5 mM) plus superoxide dismutase (100 units/ml). NADPH fluorescence (lambda exc. = 340 nm; lambda em. = 460 nm) was recorded spectrofluorometrically at 120-s intervals before and after photoactivation (for 10 min) of the dyes (TMRM lambda exc. = 535 ± 17.5 nm; Rho 123 lambda exc. = 488 ± 10 nm; TBRB lambda exc. = 535 ± 17.5 nm). Enzymatically active NADP+ was determined from the increase in fluorescence (lambda exc. = 340 nm; lambda em. = 460 nm) of the irradiated mixture following the addition of D-L-isocitric acid (4.0 mM) and NADP-linked isocitric dehydrogenase (0.21 units/ml). The data shown are expressed in percent of NADPH fluorescence of untreated controls (set at 100%) and were obtained after complete equilibration and corrected for NADPH autoxidation. Zero fluorescence is equal to fluorescence of HBSS without NADPH. Values shown represent means ± SD of three experiments; compare with Fig. 1.

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 (lambda exc. = 543 nm, lambda em. >=  560 nm; see below) and Rho 123 (lambda exc. = 488 nm, lambda em. >=  505 nm), detected using laser scanning microscopy, was detectable entirely within intact mitochondria, whereas TBRB (lambda exc. = 543 nm, lambda 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 lambda exc. = 365 ± 12.5 nm and detected at lambda 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 lambda exc. = 535 ± 17.5 nm; Rho 123 lambda 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 (lambda 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.


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Fig. 1.   Effect of short term photoactivation of different rhodamine derivatives on mitochondrial NAD(P)H fluorescence of cultured hepatocytes. Cells were cultured on glass coverslips and loaded with Rho 123 (0.5 or 10.0 µM), TMRM (0.5 µM), or TBRB (2.0 µM) for 20 min in L-15 cell culture medium (37 °C). Hepatocytes were then washed three times with HBSS and incubated for another 15 min in dye-free L-15 medium. Cellular NAD(P)H fluorescence was recorded at 120-s intervals using digital fluorescence microscopy (lambda exc. = 365 ± 12.5 nm; lambda em. = 450-490 nm) and is given in arbitrary units (a.u.). After establishing the baseline fluorescence (10 min), the rhodamine derivatives were photoactivated (TMRM lambda exc. = 535 ± 17.5 nm; Rho 123 lambda exc. = 488 ± 10 nm; TBRB lambda exc. = 535 ± 17.5 nm; open arrows) for the periods indicated, and NAD(P)H fluorescence measurements were continued. Rho 123 was additionally irradiated at lambda exc. = 535 ± 17.5 nm (filled arrow). Each trace shown in A is the average of 15-25 cells. Data are representative of at least three experiments using hepatocytes from different animals. In B the effect of TMRM irradiation (10-s) on mitochondrial NAD(P)H fluorescence is shown for 28 single cells.

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).


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Fig. 2.   Effect of prolonged photoactivation of mitochondrial TMRM on mitochondrial NAD(P)H fluorescence, mitochondrial membrane potential, and viability of cultured hepatocytes. Hepatocytes were cultured on glass coverslips and loaded with TMRM (0.5 µM) for 20 min in L-15 cell culture medium (37 °C). Cells were then washed three times with HBSS and incubated for another 15 min in dye-free L-15 medium that contained propidium iodide (5 µg/ml). Cellular NAD(P)H fluorescence (lambda exc. = 365 ± 12.5 nm; lambda em. = 450-490 nm) (A), as well as mitochondrial TMRM and nuclear propidium iodide fluorescence (lambda exc. = 535 ± 17.5 nm; lambda em. >=  590 nm) (B), were recorded at 120-s intervals using digital fluorescence microscopy and are given in arbitrary units (a.u.). After establishing the baseline fluorescence (10-min), intramitochondrial TMRM was photoactivated (lambda exc. = 535 ± 17.5 nm) for 20 s (arrow), and fluorescence measurements were continued. The mitochondrial TMRM fluorescence was used as a measure for the mitochondrial membrane potential; the uptake of the vital dye propidium iodide was determined to detect loss of cell viability. Mitochondrial NAD(P)H fluorescence (A) is shown for 33 single cells as not evident from the figure. Each trace shown in B is the average of 15-30 cells. Data are representative of at least three experiments using hepatocytes from different animals; compare with Figs. 1B, 3, and 4.


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Fig. 3.   Effect of short term photoactivation of TMRM on mitochondrial integrity of cultured hepatocytes. Cells were cultured on glass coverslips and loaded simultaneously with TMRM (0.5 µM) and calcein-AM (1.0 µM) for 20 min in L-15 cell culture medium (37 °C). Cells were then washed three times with HBSS and incubated for another 15 min in L-15 medium that contained propidium iodide (5 µg/ml). The intracellular fluorescence of TMRM and propidium iodide (lambda exc. = 543 nm; lambda em. >=  585 nm) (B and D), as well as of calcein (lambda exc. = 488 nm; lambda em. = 505-530 nm) (A and C) was imaged using laser scanning microscopy. After establishing the baseline fluorescence, images were collected before (A and B) and 4 min after (C and D) photoactivation of TMRM for 10 s. The mitochondrial TMRM fluorescence was used as a measure for the mitochondrial membrane potential and cytosolic calcein to assess the permeability of the inner mitochondrial membrane. The uptake of the vital dye propidium iodide was determined to detect loss of cell viability. Note that mitochondria continued to exclude calcein after photoactivation, and TMRM fluorescence was only slightly decreased (C and D). Bar indicates 10 µm; compare with Figs. 1B, 2, and 4.


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Fig. 4.   Effect of prolonged photoactivation of TMRM on mitochondrial integrity of cultured hepatocytes. Cells were cultured on glass coverslips and loaded simultaneously with TMRM (0.5 µM) and calcein-AM (1.0 µM) for 20 min in L-15 cell culture medium (37 °C). Cells were then washed three times with HBSS and incubated for another 15 min in dye-free L-15 medium that contained propidium iodide (5 µg/ml). The intracellular fluorescence of TMRM and propidium iodide (lambda exc. = 543 nm; lambda em. >=  585 nm) (B, D, and F), as well as of calcein (lambda exc. = 488 nm; lambda em. = 505-530 nm) (A, C, and E) was imaged using laser scanning microscopy. After establishing the baseline fluorescence, images were collected before (A and B) and 4 min (C and D), as well as 22 min (E and F) after photoactivation of TMRM for 20 s. The mitochondrial TMRM fluorescence was used as a measure for the mitochondrial membrane potential and cytosolic calcein to assess the permeability of the inner mitochondrial membrane. The uptake of the vital dye propidium iodide was determined to detect loss of cell viability. Note that already 4 min after photoactivation (C and D) calcein fluorescence filled all mitochondria that lost TMRM fluorescence within the irradiated area of the cells. These cells went on to lose virtually all intracellular calcein fluorescence and finally died (E and F) as indicated by the uptake of propidium iodide. Note the nuclear shrinking and condensation of chromatin. Bar indicates 10 µm; compare with Figs. 1B, 2, and 3.

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.


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Fig. 5.   Effect of BCNU on the photoactivation-induced decrease in mitochondrial NAD(P)H fluorescence of cultured hepatocytes. Hepatocytes were cultured on glass coverslips and incubated for 1 h in L-15 cell culture medium (37 °C) with or without the glutathione reductase inhibitor BCNU (300 µM; Control). Cells were then loaded with TMRM (0.5 µM), and cellular NAD(P)H fluorescence was recorded in the presence of BCNU at 120-s intervals as described in the legend for Fig. 1. After establishing the baseline fluorescence (10-min), either intramitochondrial TMRM was photoactivated (lambda exc. = 535 ± 17.5 nm) for 10 s (arrows) or t-BuOOH (100 µM), known to lead to NAD(P)H oxidation via the glutathione peroxidase/reductase system, was added to the supernatant, and fluorescence measurements were continued. Note that BCNU completely inhibited the effect of t-BuOOH but not that of TMRM photoactivation on NAD(P)H fluorescence. Each trace shown is the average of 20-30 cells and is representative of at least three experiments using hepatocytes from different animals.

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.


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Fig. 6.   Effect of the environmental pO2 on the decrease in mitochondrial NAD(P)H fluorescence of cultured hepatocytes after short term photoactivation of TMRM. Hepatocytes were cultured on glass coverslips and loaded with TMRM (0.5 µM) as described in the legend for Fig. 1. Cellular NAD(P)H fluorescence (lambda exc. = 365 ± 12.5 nm; lambda em. = 450-490 nm) and mitochondrial TMRM fluorescence (lambda exc. = 535 ± 17.5 nm; lambda em. >=  590 nm) were recorded at 120-s intervals using digital fluorescence microscopy and are given in arbitrary units (a.u.). After establishing mitochondrial baseline fluorescence of NAD(P)H and TMRM, respectively, flushing the cells (in air-tight chambers) with 5% CO2/21% O2/74% N2 was continued, or hypoxia was induced by flushing with 95% N2/5% CO2 (open arrows). Twenty min later, intramitochondrial TMRM was photoactivated (lambda exc. = 535 ± 17.5 nm) for 10 s (filled arrows), and fluorescence measurements were continued. Note the slight increase in NAD(P)H fluorescence and the lack of any effect of TMRM photoactivation on NAD(P)H of hypoxic cells. Each trace shown is the average of 20-30 cells and is representative of at least three experiments using hepatocytes from different animals.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES

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 M-1 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 alpha -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); alpha -tocopherol: 0.05-2.28 nmol/mg protein (45, 46) (approx 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<UP><SUB>2</SUB><SUP>&cjs1138;</SUP></UP> and contaminating heavy metal ions (Table I); this is in line with previous studies that reported that 1O2 quenching other than by electron transfer from NADPH would be most unlikely (7, 8, 10). It is true that the generation of intracellular reactive species other than 1O2 such as lipid peroxyl, lipid alkoxyl radicals, or ·OH cannot be completely excluded. However, it is very unlikely that they contribute to the decrease in NAD(P)H fluorescence, as their formation would surely result in enzymatic NAD(P)H oxidation via the glutathione peroxidase/reductase system that was found to be unlikely here (see Figs. 3, 5, and 6).

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 beta -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<UP><SUB>2</SUB><SUP>&cjs1138;</SUP></UP> generated during the 1O2-mediated NAD(P)H oxidation (see Reactions 7 and 8).

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<UP><SUB>2</SUB><SUP>&cjs1138;</SUP></UP>, H2O2) (50), mitochondria are also very vulnerable to attacks by these species (50, 51). Accordingly, mitochondrial photosensitizers have been reported to induce apoptosis very rapidly (42, 49, 52, 53). However, the initial events leading to the several apoptotic cascades and pathways involved in cell death are often unknown (54) and have almost invariably been attributed to oxidation of proteins and membrane lipids, leading to disruption of the inner mitochondrial membrane (28, 38, 42, 49, 52, 55-58). In this context, it appears doubtful whether, among the photochemical drugs applicable for PDT in clinical trials, real mitochondrial photosensitizers (like TMRM and Rho 123) actually exist. Irradiation of cultured hepatocytes loaded with the photochemical drugs photofrin, Al(III) phthalocyanine chloride tetrasulfonic acid (AlPcS4), meso-tetra(4-sulfonatophenyl)porphine dihydrochloride (TPPS4), or 5-aminolevulinic acid-(5-ALA-) generated protoporphyrin IX, in no case led to a direct oxidation of mitochondrial NAD(P)H, most likely because the required co-localization was not given.2 The observed decreases in NAD(P)H fluorescence always resulted from loss in cell viability/mitochondrial integrity (most likely mediated by extramitochondrial 1O2 generation) and were thus in contrast to the effects of TMRM and Rho 123 described here.

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<UP><SUB>2</SUB><SUP>&cjs1138;</SUP></UP>, which may, for example, promote formation of hydroxyl radicals, are generated within the mitochondrial matrix space (see Reactions 7 and 8), and under these conditions of oxidative stress both the direct antioxidative function of NAD(P)H and the whole antioxidative network will be impaired, the latter because of the central role of NADPH as an indirectly operating antioxidant in the regeneration of others that operate directly (1, 2). Thus, NAD(P)H-dependent enzymatic reductions will be abruptly terminated. Consequently, mitochondrial energy status/ATP levels and membrane potential will be disturbed by deprivation of electrons from metabolic processes (7, 8, 10, 11). Such an imbalance in mitochondrial energy and redox status is known to be involved in modulating the mitochondrial permeability transition pore, thus promoting further ROS generation and onset of MPT, a well known trigger of apoptosis (26, 39, 43, 59). This presumption is supported by the fact that after photoactivation of low concentrations of Rho 123 or TBRB, which had no significant effect on NAD(P)H redox state, almost no toxic effects became apparent within 60 min. The lack of a NAD(P)H oxidizing effect of both dyes under these conditions is likely to reflect either the low 1O2 quantum yield of Rho 123 (20, 28) or the weak cellular uptake and predominantly cytosolic/lysosomal localization of TBRB. Although the photochemical effects observed here will inevitably differ with cell type, for instance high phototoxicity of Rho 123 and especially TBRB has been convincingly demonstrated for MGH-U1 bladder carcinoma cells (20, 21), the data presented here should have implications for photochemotherapy. The effective generation of 1O2 in close proximity to the main pool of cellular NAD(P)H by using a photosensitizer with a high 1O2 quantum yield should improve the photochemotherapeutic potential of cancer treatment and diminish side effects because of the effective destruction of the antioxidative network of the targeted cells. In this context, recording of NAD(P)H fluorescence intensity was found to be a reasonable dosimetric measure of cell damage induced by TMRM here and during PDT (33). Especially mitochondrial photosensitizers should be advantageous for PDT as they (i) are effective inducers of apoptosis, which, in contrast to necrosis is normally not accompanied with an inflammatory response, and (ii) do not lead to sublethal nuclear DNA damages and thus genome aberrations, a risk accompanied with the application of nuclear photosensitizers.

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
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES

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.

Dagger 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
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES

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