Institute for Environmental Medicine and Department of Biochemistry and Biophysics, School of Medicine, University of Pennsylvania, Philadelphia, Pennsylvania 19104
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ABSTRACT |
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Nitric oxide (· NO) can be produced
within the lung, and recently inhaled nitric oxide has been used as a
therapeutic agent. Peroxynitrite1
(ONOO), the product of
the nearly diffusion-limited reaction between · NO and
superoxide, may represent the proximal reactive species mediating
· NO injury to pulmonary cells. To investigate the
physiological and pathological reactivities of · NO and
ONOO
at the molecular and
cellular levels, bovine pulmonary artery endothelial cells (BPAEC) and
rat type II epithelial cells were exposed to · NO
(0.01-2.5 µM/min for 2 h) generated by spermine-NONOate and
papa-NONOate and to the same fluxes of
ONOO
generated by
1,3-morpholinosydnonimine (SIN-1). Exposure to SIN-1 resulted in
cellular injury and death in both cell types. Epithelial cells
displayed a concentration-dependent loss of cellular viability within 8 h of exposure. In contrast, BPAEC loss of cellular viability was
evident after 18 h postexposure. Events preceding cell death in BPAEC
include depolarization of the mitochondrial membrane, evident as early
as 6 h postexposure, loss of cellular redox activity at 16 h, and DNA
fragmentation detected by in situ staining at 18 h after exposure.
Exposure of BPAEC to · NO did not affect the cellular
viability, but type II cells were injured in a manner similar to
ONOO
exposure.
· NO-mediated cellular injury within type II cells was
reduced by preincubation with
N-acetylcysteine. The data imply that
the pathological and physiological effects of · NO may be regulated by its reactions with superoxide and reduced thiols.
apoptosis; superoxide; endothelium; type II epithelium
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INTRODUCTION |
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NITRIC OXIDE is an important physiological regulator controlling many functions within the pulmonary system (12). However, overproduction of nitric oxide and nitric oxide-derived oxidants such as peroxynitrite has been associated with pulmonary cellular injury (2, 11, 15, 19, 22, 27, 39). Nitric oxide is capable of interacting with a number of cellular targets, including heme and nonheme iron, thiols, oxygen, and superoxide anion (1, 12, 14, 32). Reaction with these targets can result in either physiological effects, such as the activation of guanylate cyclase, or pathological effects, as in the production of peroxynitrite. Peroxynitrite is formed by the nearly diffusion-limited reaction of nitric oxide with superoxide and is a stronger oxidant than either nitric oxide or superoxide (1). Based on the reported rate constants for the reactions of peroxynitrite with various biomolecules, it appears that zinc fingers, iron-sulfur centers, CO2, and tyrosine residues in proteins may be selective cellular targets of peroxynitrite in vivo (1, 7, 9, 16, 20).
Experimental evidence has indicated that both nitric oxide and peroxynitrite are capable of inducing oxidative stress and ultimately cell death (4, 5, 10, 13, 24). In these cell models, at relatively low concentration of nitric oxide or peroxynitrite, cell death was delayed and was associated with morphological and DNA changes indicative of apoptosis (4, 5). These observations raised interesting questions. 1) Are pulmonary cells similarly sensitive to nitric oxide and peroxynitrite exposure? 2) What are the initial events and molecular pathways that lead to pulmonary cell death?
In response to these questions, we exposed cultured bovine pulmonary artery endothelial cells (BPAEC) and primary cultured type II epithelial pneumocytes to peroxynitrite and nitric oxide in kinetically defined and controlled conditions. The evidence presented here shows that exposure to peroxynitrite resulted in cell death in both pulmonary endothelial and epithelial cells. However, the kinetics of cell death differed between the two cell types, occurring within 8 h of exposure within type II cells and only after 18-24 h in endothelial cells. In addition, cell death did result from nitric oxide exposure in type II cells but not in endothelial cells. Examination of the mechanism of peroxynitrite-mediated cell death in endothelial cells reveals that 1) an early event after exposure to peroxynitrite is the depolarization of mitochondrial membrane and 2) the collapse of the mitochondrial membrane potential results in a decline in energy production, loss of redox activity, and ultimately to cell death. The nitric oxide-mediated cell death seen in type II cells can be attenuated by pretreatment with N-acetylcysteine.
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MATERIALS AND METHODS |
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Methods for exposing cells to peroxynitrite and nitric oxide. For exposure to peroxynitrite, 3-morpholinosydnonimine (SIN-1) was used as described in detail previously (31). The formation of peroxynitrite by the autoxidation of SIN-1 was confirmed by the oxidation of dihydrorhodamine 123 (DHR-123). SIN-1, in a time- and concentration-dependent manner, oxidized DHR-123 to rhodamine 123 (R-123), and this oxidation was inhibited by superoxide dismutase (92% after 60 min at 37°C). The oxidation of DHR-123 is mediated by peroxynitrite and not by either nitric oxide or superoxide alone (31). For exposure to nitric oxide, we utilized spermine-NONOate and papa-NONOate. The concentration of nitric oxide from the decomposition of these compounds was measured by a nitric oxide-selective electrochemical sensor (World Precision Instruments, Sarasota, FL). The surface area of an average-size venule or arteriole (diameter of 50 µm) that encompasses 1 liter of blood is 1.96 × 105 cm2. Based on the reported rates of nitric oxide release by stimulated endothelial cells (1), the rate of nitric oxide and potentially of peroxynitrite released can be estimated to reach as much as 6.7 µM/min in a venuole or arteriole. Therefore, cells were exposed to nitric oxide and peroxynitrite generated at a rate of 0.01-2.5 µM/min. At the end of the exposure, the cells were extensively washed and returned to the normal culture media.
Cell culture. Starter cultures of BPAEC were originally a gift of Dr. E. J. Macarack (Connective Tissue Research Institute, University of Pennsylvania). Briefly, endothelial cells from calf pulmonary arteries were isolated by collagenase digestion and cultured in medium 199 supplemented with 16% fetal calf serum, 50 µg/ml gentamicin sulfate, and 2.5 µg/ml amphotericin B. Mitotic clones were obtained by isolating individual cells from the primary culture. The cells were subcultured as necessary on 35-cm2 plastic tissue culture plates. Studies were performed at 1-3 days postconfluence and at passages 3-6. For experimental use, confluent monolayers of BPAEC (0.8-1.0 × 106 cells/well from a 6-well plate) were used throughout these studies. The cellular protein content was determined by scraping the cells from the plates and sonication three times for 10 s at 75 W. The protein in cell lysates was determined by the Bio-Rad protein assay kit.
Type II cells were isolated from anesthetized pathogen-free male Sprague-Dawley rats weighing 200-250 g by the method of Chen et al. (8). The purity of the freshly isolated type II cell preparation was routinely >80% by modified Papanicolaou stain, and the viability was >98% by vital dye exclusion. Cells were plated at 5 × 106 type II cells per 35-mm plastic tissue culture dish. Cells were cultured in 10% fetal calf serum in Eagle's minimal essential medium at 37°C in a humidified incubator with 5% CO2 in air. After overnight culture and removal of nonadhered cells, the purity of the type II cells was >90%. Typically 2 × 106 type II cells were exposed to nitric oxide- and peroxynitrite-generating compounds.
Cellular viability. Viability was determined by the uptake of either ethidium homodimer-1 (EH-1) or YO-PRO (Molecular Probes). EH-1 is highly impermeant to intact cell membranes but readily crosses compromised membranes, and, upon binding to nucleic acids, undergoes a 40-fold enhancement of fluorescence. YO-PRO is also impermeant to cell membranes and undergoes a 100-fold enhancement of fluorescence upon binding nucleic acids. At the end of the exposures, cells were incubated with either 5 µM EH-1 or 2 µM YO-PRO for 1-24 h. Fluorescence was observed in an inverted microscope (Nikon Diaphot). The EH-1 emission fluorescent intensity in cell lysates was measured at 650 nm after excitation at 530 nm.
Rhodamine fluorescence. R-123 accumulates in the charged membrane compartments of living cells, and the loss of fluorescence indicates membrane depolarization (21). Cells were loaded with 10 µM R-123 for 1 h before different exposures. Rhodamine fluorescence was observed in an inverted microscope (Nikon Diaphot). The R-123 fluorescent intensity in cell lysates was measured at excitation of 500 nm and emission of 536 nm as described previously (31).
3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide reduction. At the end of the exposure, the cells were incubated with 0.5 mg/ml 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) for 30 min. The cells were then extensively washed, harvested, and lysed by sonication. The cell lysates were gently centrifuged, and the absorbance at 570 nm was recorded.
In situ DNA fragmentation. The
procedure utilizes the two enzymatic activities of unmodified T7 DNA
polymerase (a 5'3' DNA polymerase activity and a
3'
5' exonuclease activity) and has been shown to
measure DNA fragmentation in situ after exposure of PC-12 cells to
peroxynitrite (4). Cells were fixed by incubation for 10 min with 4%
formaldehyde in phosphate-buffered saline (PBS). The cells were
permeabilized by incubating for 5 min in 0.1% Triton X-100, washed in
PBS, and dehydrated by addition of ethanol solutions followed by
delipidation by addition of chloroform for 2 min. After
rehydration, the fixed cells were covered with a T7 reaction buffer
(Epicenter Technologies) for 10 min followed by a 10-min incubation
with 0.1 U/ml unmodified T7 DNA polymerase. The cells were then
incubated with 0.2 mM each dCTP, dGTP, and dTTP, 0.002 mM dATP, and 0.4 mM biotin-14-dATP. The reaction was stopped by the addition of 0.5 mM
EDTA. The single-stranded gaps generated by the exonuclease from any
DNA breaks present at the time of fixation were filled with newly
synthesized DNA that is tagged by the biotinylated nucleotides. The
biotinylated nucleotides are visualized by incubation with
streptavidin-peroxidase and a peroxidase substrate to produce a color
reaction.
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RESULTS |
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Exposure of vascular endothelial cells to peroxynitrite induces delayed cell death. Initially, the time course for the induction of cell death was determined by exposing BPAEC to 1-2 µM/min peroxynitrite generated by SIN-1 or to the same flux of nitric oxide generated by spermine-NONOate for 2 h. After exposure, the cells were extensively washed and returned to normal media, and cell viability was monitored by the uptake of EH-1. The cell membrane permeability and cell viability were not compromised in control cells or cells exposed to nitric oxide for up to 24 h after exposure (Fig. 1, A, C, D, and F). In contrast, starting at 16-18 h, the cell viability declined in peroxynitrite-exposed cells. Exposure to 2 µM/min peroxynitrite for 2 h was found to be the condition that resulted in >90% cell death after 18 h, and this exposure was used throughout this manuscript. Addition of superoxide dismutase (1 mg/ml, 3,250 U/mg) abolished SIN-1-mediated loss of cellular viability. Sixteen hours after exposure to nitric oxide, cell lysates contained 1.5 ± 0.5 arbitrary fluorescence units of EH-1 (AFU)/mg cell protein, and control cells contained 1.3 ± 0.3 AFU/mg, whereas peroxynitrite-exposed cells contained 5.6 ± 0.8 AFU/mg cell protein, indicating a significant increase in membrane permeability. By 24 h, all of the peroxynitrite-treated cells were nonviable (Fig. 1E). Completely decomposed donor compounds SIN-1C and spermine had no effect on cellular viability. Examination by phase-contrast microscopy 24 h after peroxynitrite exposure revealed cellular shrinkage as well as cellular rounding up (Fig. 1B). The induction of cell death several hours after the initial insult suggested that peroxynitrite was able to initiate cellular alterations that eventually lead to death.
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Exposure of type II pneumocytes to nitric oxide and peroxynitrite induces delayed cell death. The time course for type II cell injury upon exposure was examined by following the uptake of YO-PRO relative to the loss of R-123 fluorescence after a 2-h exposure to either 0.25-2.5 µM/min peroxynitrite generated by SIN-1 or 0.01-0.9 µM/min nitric oxide generated by papa-NONOate. After exposure, cells were extensively washed, and loss of fluorescence from R-123 and gain of fluorescence from YO-PRO were monitored by microscopy. R-123 is a lipophilic and positively charged molecule that partitions to mitochondria and other negatively charged intracellular compartments (21). Thus cells that maintain their mitochondrial membrane potential retain R-123 and can be visualized by epifluorescence imaging. Cells were loaded with R-123 1 h before exposure. Figure 2 shows fluorescence of both labels in cells 8 h after exposure to either nitric oxide or peroxynitrite. Both nitric oxide and peroxynitrite exposure reduced the number of cells with R-123 fluorescence relative to control, indicating depolarization of intracellular membranes, and increased the number of cells with YO-PRO fluorescence, indicating loss of membrane integrity. In addition, there was an increase in background fluorescence indicative of leakage of R-123 to the medium. Cell death was quantified by counting the number of cells per field displaying YO-PRO fluorescence (nonviable cells) relative to the number of cells displaying R-123 fluorescence (viable cells). Figure 3 shows the percentage of nonviable cells after exposure to either nitric oxide (Fig. 3A) or peroxynitrite (Fig. 3B). There was no evidence of loss of cellular viability when type II cells were exposed to the decomposed donor compounds SIN-1C and papa-NONOate. The degree of cellular injury was concentration dependent, and there was no increase in cell death between 8 and 18 h postexposure.
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Events preceeding peroxynitrite-mediated endothelial cell death. Changes in mitochondrial membrane potential within BPAEC were monitored by the ability of mitochondria to retain the fluorescent probe R-123. Figure 4, A-C, shows R-123 fluorescence at the end of 1 h of loading. Figure 4, D-F, shows R-123 fluorescence 6 h after exposure to either peroxynitrite or nitric oxide. The loss of R-123 fluorescence in peroxynitrite-treated cells indicates depolarization of the mitochondrial membrane potential. Cells were impermeable to EH-1 at this time (data not shown). The peroxynitrite-induced loss of mitochondrial potential was prevented by the addition of cyclosporin A at the end of the exposure. R-123 fluorescence in cell lysates after 2 h of exposure to 200 µM SIN-1 was 53.7 ± 11.7% of control (n = 5), whereas cells incubated with the same concentration of SIN-1 plus 30 µM cyclosporin A contained 127.8 ± 47.3% of control (n = 5). Cyclosporin A has been previously shown to prevent mitochondrial membrane depolarization and mitochondrial calcium efflux (28, 33).
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Loss of intracellular membrane potential, indicated by loss of R-123 fluorescence, predicted that the cells were energetically compromised some time after peroxynitrite exposure. Therefore, the redox activity and the pyridine nucleotide levels of BPAEC were measured by the intracellular reduction of MTT to give formazan, which has a 570-nm absorbance maximum. The reduction of MTT by the mitochondrial and cytosolic dehydrogenase(s) requires NADH and, therefore, reflects the availability of pyridine nucleotides and the cellular redox state. The ability of peroxynitrite-treated endothelial cells to reduce MTT declined over time after exposure. The ability of peroxynitrite-treated cells to form formazan 16 h after exposure was 20.5 ± 3.4% of control (control value, 0.169 ± 0.008 absorbance units at 570 nm/ 0.8 × 106 cells, n = 4). The loss in MTT reduction by peroxynitrite treatment was prevented by the addition of cyclosporin A at the end of the exposure (86 ± 10.2% of control, n = 4). A similar exposure of endothelial cells to nitric oxide did not influence the cellular redox activity.
DNA fragmentation in endothelial cells exposed to peroxynitrite but not in control or nitric oxide-treated cells was detected by in situ labeling of fragmented DNA. Figure 5 depicts the presence of fragmented DNA in peroxynitrite-treated cells 18 h after exposure and before induction of morphological changes. Apparently, not all cells showed fragmented DNA labeling, similar to the observation that not all cells showed the same morphological changes after 24 h.
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Nitric oxide-mediated epithelial cell death was attenuated by N-acetylcysteine. The intracellular concentration of reduced thiol may be critical in determining the outcome after nitric oxide exposure. Therefore, the experiments outlined in Fig. 3 were repeated after pretreating the type II cells with 2 mM N-acetylcysteine for 1 h to increase the intracellular concentration of reduced thiol. Figure 6 shows that pretreatment with N-acetylcysteine reduced the number of cells labeled with YO-PRO while increasing the number that retained R-123. These results indicate that N-acetylcysteine reduced the susceptibility of type II cells to nitric oxide-mediated injury.
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DISCUSSION |
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Nitric oxide is an important physiological regulator; however, even at low concentrations, it has been shown to alter respiration (3, 34) and to induce cell death (5). The balance between physiological regulation and pathological effect is dependent on the relative concentrations of nitric oxide and reactive biological targets. The interaction of nitric oxide with superoxide results in the formation of peroxynitrite, which may be a critical mediator capable of initiating events that lead to delayed cell death. Treatment with nitric oxide did not appear to result in cellular injury in cultured BPAEC. However, it did appear to induce cell death in type II epithelial cells. Interestingly, the cell death induced in type II cells by either nitric oxide or peroxynitrite did not appear to follow the same time course as endothelial cell death. Cell death was observed within 8 h of exposure, and there was no increase in the loss of cells in the next 10 h of incubation. Several factors, such as cellular antioxidant capacity, ability to sustain metabolic requirements by deriving energy from alternate pathways, efficiency to repair oxidatively modified biomolecules, and availability and utilization of trophic support, will influence the inherent ability of cells to withstand oxidative stress. Recently, some of these factors have been critically evaluated by Burney and co-workers (5) in cell lines exposed to nitric oxide. It appears that, in some cell types, reduced thiols may provide antioxidant defense against both nitric oxide and peroxynitrite (5, 30, 37). This may be true at least for the type II cells, since preincubation with N-acetylcysteine, which increases the intracellular reduced thiol concentration, provided protection against nitric oxide-mediated injury. Because endothelium continuously produces nitric oxide, it is reasonable to propose that an increased reduced thiol availability or a lower steady-state concentration of superoxide may prevent the pathological reactivities of nitric oxide. The observation that both epithelial and endothelial cells are susceptible to peroxynitrite-mediated cellular injury suggests that the cellular defenses that protected endothelial cells from nitric oxide-mediated injury are insufficient to protect against peroxynitrite. However, the time course of cell death within the two cell types was different, and it may be that endothelial defenses, such as reduced thiols, protected these cells from the immediate toxic effects of peroxynitrite but not against a delayed cellular injury mechanism. Delayed cellular dysfunction and death are observed after an initial exposure to oxidants in disease processes (36). The data presented indicated that mitochondrial depolarization and loss of redox capacity are early events in delayed cell death induced by peroxynitrite. Depletion of cellular energy can be derived by either alteration of mitochondrial function and inhibition of glycolytic pathways (5) or indirectly by activation of poly(ADP) ribosyl synthase, which appears to be activated in response to DNA breaks (35). Such alterations have been considered as early events in different models of apoptotic death (23, 40). Therefore, peroxynitrite may be a critical mediator capable of initiating events that lead to delayed vascular endothelial cell death during oxidative stress in sepsis, ischemia-reperfusion, and atherosclerosis (19, 38, 39).
Mitochondria appear to be a critical target for both nitric oxide and peroxynitrite-mediated cellular effects. Both nitric oxide and peroxynitrite may alter mitochondrial electron transport function, although by alternate mechanisms (3, 6, 18, 25, 28, 29, 33). Experimental evidence suggests that nitric oxide targets cytochrome oxidase and there is also a potential for interaction with iron sulfur proteins, although this is not thought to occur physiologically, whereas peroxynitrite appears to inhibit complexes I-III (3, 6, 25, 29) and to induce a cyclosporin A-sensitive calcium efflux (28). Peroxynitrite has also been shown to induce calcium efflux from isolated liver mitochondria by oxidation of critical thiols in a manner that induces pyridine nucleotide-linked calcium release, a pathway inhibited by cyclosporin A (33). Moreover, within neuronal cells, the intracellular level of glutathione, and hence reduced thiol, has been shown to be critical in determining the effectiveness of nitric oxide inhibition of mitochondrial function (3). The nitric oxide- or peroxynitrite-induced uncoupling of the mitochondrial electron transport chain will result in an increase in superoxide and hydrogen peroxide production (6, 29). The increase in superoxide and hydrogen peroxide can be further amplified by the inactivation of Mn superoxide dismutase that is strategically located inside the mitochondria to account for the production of superoxide (20, 26). Recently, Mn superoxide dismutase was found to be nitrated and inactivated in rejected human transplanted kidney tissues (26). Overall, it appears that mitochondria may represent a critical target for nitric oxide and peroxynitrite and collapse of the mitochondrial membrane potential, and decline in energy production and reduced equivalents represent early events in peroxynitrite-induced cell death.
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ACKNOWLEDGEMENTS |
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We are grateful to Dr. A. Al-Mehdi (University of Pennsylvania) for helpful discussions and reading of the manuscript, Dr. Q.-P. Chen for assistance in the preparation of type II pneumocytes, Drs. C. Richter and M. Schweizer (Eldgenössische Technische Hochschule, Zurich) for helpful discussions regarding the use of cyclosporin A, and June Nelson and Kathy Notarfransesco for expert technical assistance.
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FOOTNOTES |
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This work was supported by National Heart, Lung, and Blood Institute (NHLBI) Grant HL-54926 (to H. Ischiropoulos) and National Institute of Environmental Health Sciences Grant ES-05211 and by funds from the Council for Tobacco Research (to S. R. Thom). A. J. Gow was supported by a National Research Service Award from the NHLBI (HL-07748), and H. Ischiropoulos is an Established Investigator of the American Heart Association.
1
The IUPAC-recommended nomenclature for nitric
oxide is nitrogen monoxide and for peroxynitrite is oxoperoxonitrate
(1). Peroxynitrite indicates both the anion form and the
conjugated acid peroxynitrous acid (ONOOH), unless otherwise
indicated.
Address for reprint requests: H. Ischiropoulos, Univ. of Pennsylvania, Institute for Environmental Medicine, 1 John Morgan Bldg., 3620 Hamilton Walk, Philadelphia, PA 19104-6068.
Received 28 April 1997; accepted in final form 6 October 1997.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Beckman, J. S.,
and
W. H. Koppenol.
Nitric oxide, superoxide, and peroxynitrite: the good, the bad, and the ugly.
Am. J. Physiol.
271 (Cell Physiol. 40):
C1424-C1437,
1996
2.
Bernareggi, M.,
J. A. Mitchell,
P. J. Barnes,
and
M. G. Belvici.
Dual action of nitric oxide on airway plasma leakage.
Am. J. Respir. Crit. Care Med.
155:
869-874,
1997[Abstract].
3.
Bolanos, J. P.,
S. J. Heales,
S. Peuchen,
J. E. Barker,
J. M. Land,
and
J. B. Clark.
Nitric oxide mediated mitochondrial damage: a potential neuroprotective role for glutathione.
Free Radic. Biol. Med.
21:
995-1001,
1996[Medline].
4.
Bonfoco, E.,
D. Krainc,
M. Ankarcrona,
P. Nicotera,
and
S. A. Lipton.
Apoptosis and necrosis: two distinct events induced, respectively, by mild and intense insults with N-methyl-D-aspartate or nitric oxide/superoxide in cortical cell cultures.
Proc. Natl. Acad. Sci. USA
92:
7162-7166,
1995[Abstract].
5.
Burney, S.,
S. Tamir,
A. Gal,
and
S. R. Tannenbaum.
A mechanistic analysis of nitric oxide-induced cellular toxicity.
Nitric Oxide: Biol. Chem.
1:
130-144,
1997.[Medline]
6.
Cassina, A,
and
R. Radi.
Differential inhibitory action of nitric oxide and peroxynitrite on mitochondrial electron transport.
Arch. Biochem. Biophys.
328:
309-316,
1996[Medline].
7.
Castro, L.,
M. Rodriguez,
and
R. Radi.
Aconitase is readily inactivated by peroxynitrite, but not by its precursor, nitric oxide.
J. Biol. Chem.
269:
29409-29415,
1994
8.
Chen, Q.,
S. R. Bates,
and
A. B. Fisher.
Secretagogues increase the expression of SP-A receptors on lung type II cells.
J. Biol. Chem.
271:
25277-25283,
1996
9.
Crow, J. P.,
J. S. Beckman,
and
J. M. McCord.
Sensitivity of the essential zinc-thiolate moiety of yeast alcohol dehydrogenase to hypochlorite and peroxynitrite.
Biochemistry
34:
3544-3522,
1995[Medline].
10.
Estévez, A. G.,
R. Radi,
L. Barbeito,
J. T. Shin,
J. A. Thompson,
and
J. S. Beckman.
Peroxynitrite-induced cytotoxicity in PC12 cells: evidence for an apoptotic mechanism differentially modulated by neurotrophic factors.
J. Neurosci.
65:
1543-1550,
1995.
11.
Garat, C.,
C. Jayr,
S. Eddahibi,
M. Laffon,
M. Meigan,
and
S. Adnot.
Effects of inhaled nitric oxide or inhibition of endogenous nitric oxide formation on hyperoxic lung injury.
Am. J. Respir. Crit. Care Med.
155:
1957-1964,
1997[Abstract].
12.
Gaston, B.,
J. M. Drazen,
J. Loscalzo,
and
J. S. Stamler.
The biology of nitrogen oxides in the airways.
Am. J. Respir. Crit. Care Med.
149:
538-551,
1994[Abstract].
13.
Gergel, D.,
V. Misik,
K. Ondrias,
and
A. I. Cederbaum.
Increased cytotoxicity of 3-morpholinosydnonimine to HepG2 cells in the presence of superoxide dismutase. Role of hydrogen peroxide and iron.
J. Biol. Chem.
270:
20922-20929,
1996
14.
Gow, A. J.,
D. G. Buerk,
and
H. Ischiropoulos.
A novel reaction mechanism for the formation of S-nitrosothiol in vivo.
J. Biol. Chem.
272:
2841-2845,
1997
15.
Haddad, I. Y.,
G. Pataki,
P. Hu,
C. Galliani,
J. S. Beckman,
and
S. Matalon.
Quantitation of nitrotyrosine levels in lung sections of patients and animals with acute lung injury.
J. Clin. Invest.
94:
2407-2413,
1994[Medline].
16.
Hausladen, A.,
and
I. Fridovich.
Superoxide and peroxynitrite inactivate aconitases but nitric oxide does not.
J. Biol. Chem.
269:
29405-29408,
1994
17.
Henry, Y.,
and
R. Cassoly.
Chain non-equivalence in nitric oxide binding to hemoglobin.
Biochem. Biophys. Res. Commun.
51:
659-665,
1973[Medline].
18.
Hu, P.,
H. Ischiropoulos,
J. S. Beckman,
and
S. Matalon.
Peroxynitrite inhibition of oxygen consumption and sodium transport in alveolar type II cells.
Am. J. Physiol.
266 (Lung Cell. Mol. Physiol. 10):
L628-L634,
1994
19.
Ischiropoulos, H.,
A. B. Al-Mehdi,
and
A. B. Fisher.
Reactive species in rat lung injury: contribution of peroxynitrite.
Am. J. Physiol.
269 (Lung Cell. Mol. Physiol. 13):
L155-L164,
1995.
20.
Ischiropoulos, H.,
L. Zhu,
J. Chen,
J.-H. M. Tsai,
J. C. Martin,
C. D. Smith,
and
J. S. Beckman.
Peroxynitrite-mediated tyrosine nitration catalyzed by superoxide dismutase.
Arch. Biochem. Biophys.
298:
431-437,
1992[Medline].
21.
Johnson, L. V.,
M. L. Walsh,
B. J. Bocus,
and
L. B. Chen.
Monitoring of relative mitochondrial membrane potential in living cells by fluorescence microscopy.
J. Cell Biol.
88:
526-535,
1981[Abstract].
22.
Kooy, N. W.,
J. A. Royall,
Y.-Z. Ye,
D. R. Kelly,
and
J. S. Beckman.
Evidence for in vivo peroxynitrite production in human acute lung injury.
Am. J. Respir. Crit. Care Med.
151:
1250-1254,
1995[Abstract].
23.
Lemasters, J. J.,
J. DiGuiseppi,
A.-L. Nieminen,
and
B. Herman.
Blebbing, free calcium and mitochondrial membrane potential preceding cell death in hepatocytes.
Nature
325:
78-81,
1987[Medline].
24.
Lin, K.-T.,
J.-Y. Xue,
M. Nomen,
B. Spur,
and
P. Y.-K. Wong.
Peroxynitrite-induced apoptosis in HL-60 cells.
J. Biol. Chem.
270:
16487-16490,
1995
25.
Lizasoain, I.,
M. A. Moro,
D. G. Knowles,
V. Darley-Usmar,
and
S. Moncada.
Nitric oxide and peroxynitrite exert distinct effects on mitochondrial respiration which are differentially blocked by glutathione or glucose.
Biochem. J.
314:
877-880,
1996[Medline].
26.
MacMillan-Crow, L. A.,
J. P. Crow,
J. D. Kerby,
J. S. Beckman,
and
J. A. Thompson.
Nitration and inactivation of Mn superoxide dismutase in chronic rejection of human renal allografts.
Proc. Natl. Acad. Sci. USA
93:
11853-11858,
1996
27.
Mulligan, M. S.,
J. M. Hevel,
M. A. Marletta,
and
P. A. Ward.
Tissue injury caused by deposition of immune complexes is L-arginine dependent.
Proc. Natl. Acad. Sci. USA
88:
6338-6342,
1991[Abstract].
28.
Packer, M. A.,
and
M. P. Murphy.
Peroxynitrite formed by simultaneous nitric oxide and superoxide generation causes a cyclosporin A-sensitive mitochondrial calcium efflux and depolarisation.
Eur. J. Biochem.
234:
231-239,
1995[Abstract].
29.
Poderoso, J. J.,
M. C. Cerreras,
C. Lisdero,
N. Riobo,
F. Scöpfer,
and
A. Boveris.
Nitric oxide inhibits electron transfer and increases superoxide radical production in rat heart mitochondria and submitochondrial particles.
Arch. Biochem. Biophys.
328:
85-92,
1996[Medline].
30.
Radi, R.,
J. S. Beckman,
K. M. Bush,
and
B. A. Freeman.
Sulfhydryl oxidation by peroxynitrite: the cytotoxic potential of superoxide and nitric oxide.
J. Biol. Chem.
266:
4244-4250,
1991
31.
Royall, J. A.,
and
H. Ischiropoulos.
Evaluation of 2',7' dichlorofluorescin and dihydrorhodamine 123 as fluorescent probes for intracellular hydrogen peroxide in culture endothelial cells.
Arch. Biochem. Biophys.
302:
348-355,
1993[Medline].
32.
Rubbo, H.,
V. Darley-Usmar,
and
B. A. Freeman.
Nitric oxide regulation of tissue free radical injury.
Chem. Res. Toxicol.
9:
809-820,
1996[Medline].
33.
Schweizer, M.,
and
C. Richter.
Peroxynitrite stimulates the pyridine nucleotide-linked calcium release from intact rat liver mitochondria.
Biochemistry
35:
4524-4528,
1996[Medline].
34.
Shen, W.,
T. H. Hintze,
and
M. S. Wolin.
Nitric oxide: an important signaling mechanism between vascular endothelium and parenchymal cells in regulation of oxygen consumption.
Circulation
92:
3505-3512,
1995
35.
Szabo, C.,
B. Zingarelli,
and
A. Salzman.
Role of poly-ADP ribosyltransferase activation in the vascular contractile and energetic failure elicited by exogenous and endogenous nitric oxide and peroxynitrite.
Circ. Res.
78:
1051-1063,
1996
36.
Thompson, G. B.
Apoptosis in the pathogenesis and treatment of disease.
Science
267:
1456-1462,
1995[Medline].
37.
Walker, M. W.,
M. T. Kinter,
R. J. Roberts,
and
D. R. Spitz.
Nitric oxide-induced cytotoxicity: involvement of cellular resistance to oxidative stress and the role of glutathione in protection.
Pediatr. Res.
37:
41-49,
1995[Abstract].
38.
White, C. R.,
T. A. Brock,
L.-Y. Chang,
J. Crapo,
P. Brisco,
D. Ku,
W. A. Bradley,
S. H. Gianturco,
J. Gore,
B. A. Freeman,
and
M. M. Tarpey.
Superoxide and peroxynitrite in atherosclerosis.
Proc. Natl. Acad. Sci. USA
91:
1044-1048,
1994[Abstract].
39.
Wizemann, T. M.,
C. R. Gardner,
J. D. Laskin,
S. Quinones,
K. D. Durham,
N. L. Golle,
S. T. Ohnishi,
and
D. L. Laskin.
Production of nitric oxide and peroxynitrite in the lung during acute endotoxemia.
J. Leukoc. Biol.
56:
759-768,
1994[Abstract].
40.
Zamzami, N.,
P. Marchetti,
M. Castedo,
D. Decaudin,
A. Macho,
T. Hirsch,
S. A. Susin,
P. X. Petit,
B. Mignotte,
and
G. Kroemer.
Sequential reductions of mitochondrial transmembrane potential and generation of reactive oxygen species in early programmed cell death.
J. Exp. Med.
182:
367-377,
1995[Abstract].