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

2003; Elsevier BV; Volume: 278; Issue: 5 Linguagem: Inglês

10.1074/jbc.m204230200

1083-351X

Frank Petrat, Stanislaw Pindiur, Michael Kirsch, Herbert de Groot,

Sulfur Compounds in Biology

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 generating1O2 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. 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 generating1O2 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. reactive oxygen species singlet oxygen tetramethylrhodamine methyl ester 2′,4′,5′,7′-tetrabromorhodamine 123 bromide rhodamine 1,3-bis(chloroethyl)-1-nitrosourea glutathione (oxidized form) tert-butyl hydroperoxide Hanks' balanced salt solution mitochondrial permeability transition photodynamic therapy rate constant for single electron transfer 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 α-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 (1Kirsch M. de Groot H. FASEB J. 2001; 15: 1569-1574Google Scholar, 2Leopold J.A. Cap A. Scribner A.W. Stanton R. Loscalzo J. FASEB J. 2001; 15: 1771-1773Google Scholar). Consequently, NAD(P)H deficiencies are linked with an increased sensitivity to oxidative stress (2Leopold J.A. Cap A. Scribner A.W. Stanton R. Loscalzo J. FASEB J. 2001; 15: 1771-1773Google Scholar, 3Minard K.I. McAlister-Henn L. Free Radic. Biol. Med. 2001; 31: 832-843Google Scholar). 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 (4Schellenberg K.A. Hellerman L. J. Biol. Chem. 1958; 231: 547-556Google Scholar). However, we recently demonstrated that, in line with the Marcus theory of electron transfer (1Kirsch M. de Groot H. FASEB J. 2001; 15: 1569-1574Google Scholar, 5Kirsch M. de Groot H. J. Biol. Chem. 1999; 274: 24664-24670Google Scholar), the reaction constant of Reaction 1NADH+Rad⋅→NAD⋅+RHREACTION1correlated well with the reduction potential of the oxidizing radical (1Kirsch M. de Groot H. FASEB J. 2001; 15: 1569-1574Google Scholar). Consequently, putative harmful radicals (ROO⋅, RO⋅, CO3⋅−) react very fast with NADH (k r = 108-109m−1s−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.NAD⋅+O2→NAD++O2⨪REACTION2In chemical systems, O2⨪ spontaneously dismutates to H2O2 and 1O2 (6Corey E.J. Mehrotra M.M. Khan A.U. Biochem. Biophys. Res. Commun. 1987; 145: 842-846Google Scholar), shown in Reaction 3.2O2⨪+2H+→H2O2+1O2REACTION3In biological systems superoxide dismutase (SOD) catalyzes the dismutation of O2⨪, thereby preventing the formation of1O2, shown in Reaction 4.2O2⨪+2H+→SODH2O2+3O2REACTION4The H2O2-consuming enzymes catalase and glutathione peroxidase (GPx) strongly limit the noxious action of H2O2, shown in Reactions 5 and 6.REACTION5H2O2+2GSH→GPx2H2O+GSSGREACTION6Given 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 (1Kirsch M. de Groot H. FASEB J. 2001; 15: 1569-1574Google Scholar). Besides oxidizing radicals, the reactive intermediate1O2 also rapidly reacts with both NADH and NADPH (k r = 4.3 × 107m−1 s−1 and 8.4 × 107m−1 s−1) via single electron transfer (7Peters G. Rodgers M.A.J. Biochem. Biophys. Res. Commun. 1980; 96: 770-776Google Scholar, 8Peters G. Rodgers M.A.J. Biochim. Biophys. Acta. 1981; 637: 43-52Google Scholar), shown in Reactions 7 and 8.NAD(P)H+1O2→NAD(P)⋅+O2⨪+H+REACTION7NAD(P)⋅+O2→NAD(P)++O2⨪REACTION8In 1976 the thermodynamic capability of NAD(P)H to transfer only one electron to 1O2 was estimated by Koppenol (9Koppenol W.H. Nature. 1976; 262: 420-421Google Scholar). Experimental evidence of this reaction in cell-free systems was provided, and consequences of 1O2 generation in mitochondria were hypothesized (7Peters G. Rodgers M.A.J. Biochem. Biophys. Res. Commun. 1980; 96: 770-776Google Scholar, 8Peters G. Rodgers M.A.J. Biochim. Biophys. Acta. 1981; 637: 43-52Google Scholar, 10Bodaness R.S. Chan P.C. J. Biol. Chem. 1977; 252: 8554-8560Google Scholar, 11Bodaness R.S. Biochem. Biophys. Res. Commun. 1982; 108: 1709-1715Google Scholar) 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 (12Tyler D.D. The Mitochondrion in Health and Disease. VCH Publishers, Inc., New York1992Google Scholar). 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 (12Tyler D.D. The Mitochondrion in Health and Disease. VCH Publishers, Inc., New York1992Google Scholar, 13Hoek J.B. Rydström J. Biochem. J. 1988; 254: 1-10Google Scholar). To perform these studies, we established a system based on different rhodamine derivatives and on digital fluorescence microscopy to selectively generate1O2 in close proximity to this NAD(P)H pool and to record the effect on mitochondrial NAD(P)H fluorescence with high temporal resolution. 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), β-hydroxybutyric acid, acetoacetic acid, carbonyl cyanidem-chlorophenylhydrazone, β-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). 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. Hepatocytes were isolated from male Wistar rats as described previously (14de Groot H. Brecht M. Biol. Chem. Hoppe-Seyler. 1991; 372: 35-41Google Scholar). 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 mmKCl/1.0 mm CaCl2/0.5 mmMgCl2/0.4 mm KH2PO4/0.4 mm MgSO4/0.3 mmNa2HPO4/25.0 mm Hepes, pH 7.4) and supplied with fresh medium as reported previously (15Petrat F. Rauen U. de Groot H. Hepatology. 1999; 29: 1171-1179Google Scholar). The1O2 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 λexc. = 409 nm, λem. = 476 nm; 9,10-diphenylanthracene λexc. = 391 nm, λem. = 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 the1O2 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 λexc. = 535 ± 17.5 nm; Rho 123 λexc.= 488 ± 10 nm; TBRB λexc. = 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 the1O2 detector molecules was compared with that of the untreated controls. 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 (λexc. = 340 nm; λem. = 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 (λexc. = 340 nm; λem. = 460 nm) of the irradiated mixture indicating enzymatic re-reduction of NADP+ to NADPH was recorded spectrofluorometrically (11Bodaness R.S. Biochem. Biophys. Res. Commun. 1982; 108: 1709-1715Google Scholar). 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 (15Petrat F. Rauen U. de Groot H. Hepatology. 1999; 29: 1171-1179Google Scholar, 16Evans P.J. Halliwell B. Methods Enzymol. 1994; 233: 82-92Google Scholar) to minimize the transition metal contamination. 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 (17Petrat F. de Groot H. Rauen U. Biochem. J. 2001; 356: 61-69Google Scholar, 18Petrat F. Weisheit D. Lensen M. de Groot H. Sustmann R. Rauen U. Biochem. J. 2002; 362: 137-147Google Scholar). 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 (19Haugland R.P. Handbook of Fluorescent Probes and Research Products. 9th Ed. Molecular Probes, Inc., Eugene, OR1996: 479-487Google Scholar) to 514.5 nm within cells (20Shea C.R. Chen N. Wimberly J. Hasan T. Cancer Res. 1989; 49: 3961-3965Google Scholar, 21Shea C.R. Sherwood M.E. Flotte T.J. Chen N. Scholz M. Hasan T. Cancer Res. 1990; 50: 4167-4172Google Scholar). 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 (22Adamson G.M. Harman A.W. Biochem. Pharmacol. 1993; 45: 2289-2294Google Scholar, 23Harbrecht B.G., Di Silvio M. Chough V. Kim Y.M. Simmons R.L. Billiar T.R. Ann. Surg. 1997; 225: 76-87Google Scholar) 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. 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 (λexc. = 543 nm; λem. ≥ 560 nm), Rho 123 (λexc. = 488 nm; λem. ≥ 505 nm), and of TBRB (λexc. = 543 nm; λem. ≥ 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. 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 (λexc. = 535 ± 17.5 nm; λem. ≥ 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 (λexc. = 543 nm; λem. ≥ 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 (λexc. = 543 nm; λem.≥ 585 nm) and green fluorescence of Rho 123 (λexc. = 488 nm; λem. = 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 24Zahrebelski G. Nieminen A.-L., Al- Ghoul K. Qian T. Herman B. Lemasters J.J. Hepatology. 1995; 21: 1361-1372Google Scholar, 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 (24Zahrebelski G. Nieminen A.-L., Al- Ghoul K. Qian T. Herman B. Lemasters J.J. Hepatology. 1995; 21: 1361-1372Google Scholar). 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 (λexc. = 543 nm; λem. ≥ 585 nm) and green fluorescence of calcein (λexc. = 488 nm; λem. = 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 (24Zahrebelski G. Nieminen A.-L., Al- Ghoul K. Qian T. Herman B. Lemasters J.J. Hepatology. 1995; 21: 1361-1372Google Scholar, 25Nieminen A.-L. Saylor A.K. Tesfai S.A. Herman B. Lemasters J.J. Biochem. J. 1995; 307: 99-106Google Scholar, 26Nieminen A.-L. Byrne A.M. Herman B. Lemasters J.J. Am. J. Physiol. 1997; 272: C1286-C1294Google Scholar, 27Lemasters 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. Herman B. Biochim. Biophys. Acta. 1998; 1366: 177-196Google Scholar). 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 λexc. = 535 ± 17.5 nm and λem. ≥ 590 nm. 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. 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 (8Peters G. Rodgers M.A.J. Biochim. Biophys. Acta. 1981; 637: 43-52Google Scholar, 10Bodaness R.S. Chan P.C. J. Biol. Chem. 1977; 252: 8554-8560Google Scholar, 11Bodaness R.S. Biochem. Biophys. Res. Commun. 1982; 108: 1709-1715Google Scholar). When the known (20Shea C.R. Chen N. Wimberly J. Hasan T. Cancer Res. 1989; 49: 3961-3965Google Scholar, 21Shea C.R. Sherwood M.E. Flotte T.J. Chen N. Scholz M. Hasan T. Cancer Res. 1990; 50: 4167-4172Google Scholar, 28MacDonald I.J. Dougherty T.J. J. Porphyrins Phthalocyanines. 2001; 5: 105-129Google Scholar) 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 which1O2 generation has not yet been quantified, was even more effective than Rho 123, presumably because of the small1O2 quantum yield of the latter (20Shea C.R. Chen N. Wimberly J. Hasan T. Cancer Res. 1989; 49: 3961-3965Google Scholar, 28MacDonald I.J. Dougherty T.J. J. Porphyrins Phthalocyanines. 2001; 5: 105-129Google Scholar). 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(k r ≈ 1.0 × 109m−1 s−1, andk r ≈ 1.0 × 106m−1 s−1, respectively; (29Ross A.B. Mallard W.G. Helman W.P. Buxton G.V. Huie R.E. Neta P. NDRL/NIST Solution Kinetics Database 3.0. NDRL/NIST, Gaithersburg, MD1998Google Scholar)). In controls, in which the rhodamine derivatives were not photoactivated, or the samples were irradiated in the absence of the1O2 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 (30Ogilby P.R. Foote C.S. J. Am. Chem. Soc. 1982; 104: 2069-2070Google Scholar, 31Parker J.G. Stanbro W.D. J. Photochem. 1984; 25: 545-547Google Scholar). In line with our view, the fluorescence quenching of the1O2 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 of1O2. 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 a1O2 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 O2⨪, which may arise during 1O2generation (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 (11Bodaness R.S. Biochem. Biophys. Res. Commun. 1982; 108: 1709-1715Google Scholar), 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 by1O2 via Reactions 7 and 8 as suggested by Peters and Rodgers (7Peters G. Rodgers M.A.J. Biochem. Biophys. Res. Commun. 1980; 96: 770-776Google Scholar, 8Peters G. Rodgers M.A.J. Biochim. Biophys. Acta. 1981; 637: 43-52Google Scholar).Table IOxidation of NADPH to enzymatically active NADP + by 1 O2 generated during irradiation of different rhodamine derivativesRhodamine derivativeAdditions/treatmentsNADPH oxidizedRecovery of NADPH fluorescence% of control%Rhodamine 12311.2 ± 1.494.9 ± 3.4TMRM33.6 ± 5.784.4 ± 0.9TMRMSuperoxide dismutase (100 units/ml)35.1 ± 3.878.9 ± 4.4TMRMChelex31.2 ± 2.381.4 ± 3.4TBRB95.4 ± 3.791.6 ± 9.4NADPH (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 mmMgCl2 or MgCl2 (5 mm) plus superoxide dismutase (100 units/ml). NADPH fluorescence (λexc. = 340 nm; λem. = 460 nm) was recorded spectrofluorometrically at 120-s intervals before and after photoactivation (for 10 min) of the dyes (TMRM λexc. = 535 ± 17.5 nm; Rho 123 λexc. = 488 ± 10 nm; TBRB λexc. = 535 ± 17.5 nm). Enzymatically active NADP+ was determined from the increase in fluorescence (λexc. = 340 nm; λem. = 460 nm) of the irradiated mixture following the addition ofd-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. Open table in a new tab NADPH (20 μm) and the respective rhodamine derivatives (10 μm) Rho 123, TMRM, or TBRB were dissolved in HBSS (25 °C)