Journal of Leukocyte Biology eBioscience full spectrum cell analysis
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published online as doi:10.1189/jlb.0103028 on August 11, 2003

Published online before print August 11, 2003
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow EPO induces dynamic membrane blebbing (MOV, 747 KB)
Right arrow All Versions of this Article:
jlb.0103028v1
74/5/897    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by McElhinney, B.
Right arrow Articles by Janssen-Heininger, Y. M. W.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by McElhinney, B.
Right arrow Articles by Janssen-Heininger, Y. M. W.
(Journal of Leukocyte Biology. 2003;74:897-907.)
© 2003 by Society for Leukocyte Biology

Eosinophil peroxidase catalyzes JNK-mediated membrane blebbing in a Rho kinase-dependent manner

Brian McElhinney*, Matthew E. Poynter*, Punya Shrivastava*, Stanley L. Hazen{dagger},{ddagger} and Yvonne M. W. Janssen-Heininger*,§,1

* Department of Pathology, University of Vermont, Burlington; Departments of
{dagger} Cell Biology and
{ddagger} Cardiology and the Center for Cardiovascular Diagnostics and Prevention, Cleveland Clinic Foundation, Ohio; and
§ Department of Pulmonology, Maastricht University, The Netherlands

1 Correspondence: Dept. of Pathology, University of Vermont, 216A Health Sciences Research Facility, Burlington, VT 05405. E-mail: yvonne.janssen{at}uvm.edu


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Eosinophilic influx is characteristic of numerous inflammatory conditions. Eosinophil peroxidase (EPO) is a major enzyme present in eosinophils and upon degranulation, becomes released into the airways of asthmatics. As a result of its cationic nature and its ability to catalyze the formation of highly toxic oxidants, EPO has significant potential to induce cellular injury. The focus of the present study was to determine the cell-signaling events important in EPO-induced death of lung epithelial cells. In the presence of hydrogen peroxide and nitrite (NO2-; hereafter called EPO with substrates), EPO catalyzes the formation of nitrogen dioxide. EPO with substrates induced rapid and sustained activation of c-Jun-NH2-terminal kinase (JNK) and led to cell death, as was evidenced by enhanced mitochondrial depolarization, cytochrome c release, cleavage of caspases 9 and 3, poly-adenosine 5'-diphosphate ribosylation of proteins, the formation of single-stranded DNA, and membrane permeability. Moreover, EPO with substrates caused Rho-associated coiled coil-containing kinase-1-dependent dynamic membrane blebbing. Inhibition of JNK activity in cells expressing a dominant-negative JNK-1 construct (JNK-APF) prevented mitochondrial membrane depolarization and substantially decreased the number of cells blebbing compared with vector controls. The cellular responses to EPO with substrates were independent of whether NO2-, bromide, or thiocyanide was used as substrates. Our findings demonstrate that catalytically active EPO is capable of causing significant damage to lung epithelial cells in vitro and that this involves the activation of JNK.

Key Words: nitrotyrosine • asthma • epithelium • ROCK • cationic protein


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The recruitment and subsequent accumulation of eosinophils in the lung occur in a number of inflammatory disorders, including asthma [1 2 3 4 ]. Asthma is a disease characterized by reversible airway obstruction, bronchial hyper-responsiveness, inflammation, epithelial cell sloughing, and lung remodeling [5 ]. It has been suggested that eosinophils may be key effecter cells contributing to these effects [6 ]. The production of mediators by activated eosinophils is thought to be an important event in the late-phase allergic response [7 ]. Eosinophils contain numerous cytotoxic components, such as the cationic proteins, major basic protein, and eosinophil peroxidase (EPO), which can be released into the airway upon eosinophil degranulation [8 ]. EPO, a cationic, heme-containing protein, which is produced solely in eosinophils, is the most abundant of the secondary granule proteins [9 ]. Much attention has previously focused on the antimicrobial and antiparasitic effects of EPO and of leukocyte peroxidases in general [10 , 11 ], yet little is known about the effects of EPO on resident lung epithelial cells. Activated eosinophils generate hydrogen peroxide (H2O2) as a result of superoxide production from the reduced nicotinamide adenine dinucleotide phosphate oxidase system. Intraphagosomal concentrations of superoxide, the precursor of H2O2, have been estimated to exceed molar amounts [12 ], and local concentrations of H2O2 produced by granulocytes can approximate 100 µM [13 ] for protracted times [14 ], constituting a significant potential for eosinophil-derived oxidants to damage lung epithelial cells. EPO is capable of catalyzing the reaction of H2O2 with a number of ions, including bromide (Br)-, chloride (Cl-), iodide (I-), thiocyanide (SCN)-, and nitrite (NO2-; a major end-product of nitric oxide metabolism) to generate reactive species, such as hypobromous acid (HOBr) and hypochlorous acid (HOCl), capable of killing invading microbes [15 ]. The reaction of H2O2 and NO2-, catalyzed by EPO, was recently shown to generate nitrogen dioxide (·NO2) as an end product at concentrations greater than 1 ppm [16 ]. A series of elegant experiments examining the biochemistry of EPO with substrates have shown that the oxidation of resident lung proteins results in the formation of bromotyrosine and nitrotyrosine moieties [17 , 18 ]. The accumulation of bromotyrosine residues is currently viewed as a specific marker of eosinophilic inflammation, and bromotyrosine levels in severe asthmatics can be increased 84-fold as compared with nonasthmatic patients [15 ]. It is important that the use of EPO knockout (KO) mice has shown that nitrotyrosine formation is severely impaired in these mice compared with strain-matched controls after immunization and subsequent challenge with ovalbumin (OVA), illustrating that the catalytic activity of EPO plays a predominant role in tyrosine nitration observed in mice [9 ] and perhaps humans with asthma [19 ].

Previous work has focused on whether EPO is capable of causing cell death [6 ], yet the underlying mechanisms responsible for this outcome have not been elucidated. Oxidant stresses have been reported to induce apoptotic [20 , 21 ] and necrotic [22 , 23 ] forms of cell death via mechanisms that involve the mitochondria [24 25 26 27 ]. The activation of stress-activated protein kinase-1, c-Jun NH2-terminal kinase (JNK), has been demonstrated to play a crucial role in apoptotic cell death by promoting the release of cytochrome c from the mitochondria [28 ], activation of the apoptosome [29 ], and consequently, the activation of caspases 9 and 3 [30 ].

In the present study, we chose to examine the molecular effects of EPO with and without various substrates on lung epithelial cells in vitro. We demonstrate that reactive species generated by EPO with substrates induced cell death, evidenced by mitochondrial membrane depolarization, cytochrome c release, cleavage of caspases 9 and 3, poly-adenosine 5'-diphosphate (ADP)-ribosylation of proteins, increases in single-stranded DNA content, and Rho-associated coiled coil-containing kinase (ROCK)-dependent dynamic membrane blebbing. In contrast to a published report demonstrating that the toxic effects of EPO are dependent on the substrate used by EPO [31 ], our findings demonstrate that death of airway epithelial cells induced by EPO occurred independently of which substrate was present and was dependent on oxidant production by EPO. Lastly, mitochondrial depolarization and membrane blebbing, caused by EPO with substrates, were induced as a result of rapid and sustained activation of JNK.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Cell culture and exposure to agents
Spontaneously transformed alveolar type II cells (C10) were kindly provided by Dr. Alvin Malkinson (University of Colorado Health Sciences, Denver) [32 ]. The C10 cells were propagated in CRML-1066 medium containing 50 U/ml penicillin-50 µg/ml streptomycin (P/S), 2 mM L-glutamine, and 10% fetal bovine serum (FBS), all from (Gibco-BRL, Grand Island, NY). Alternatively, a line of Simian virus-40-transformed mouse Clara cells (MTCC) was obtained from Dr. Francesco DeMayo (Baylor College of Medicine, House, TX) and was propagated in Dulbecco’s modified Eagle’s medium (DMEM) high glucose-containing 50 U/ml P/S, 10% FBS, and 2 mM L-glutamine [33 ]. For the experiments, cells were plated onto 60 mm dishes and were grown to 70–90% confluency. At least 1 h before adding the test agents, the cells were switched to phenol red-free DMEM/F12 (Gibco-BRL) containing P/S and 0.5% FBS. In control experiments, glucose was supplemented to confirm that the cellular responses were not a result of glucose deprivation or pH changes as a result of glucose oxidase activity.

Reagents and plasmids
Sodium NO2, potassium Br, and potassium SCN were all obtained from Sigma Chemical Co. (St. Louis, MO) and were used at concentrations of 100 µM. Glucose oxidase (GOx), grade 2, was obtained from Boehringer Mannheim (Indianapolis, IN), reconstituted in phosphate buffer (pH 7.4) containing 50% glycerol and stored at -20°C. GOx was used at a concentration of 15 mU/ml, the minimal concentration required to ensure maximal catalytic activity of EPO. To initiate H2O2 production, GOx was incubated in medium for 5 min before addition to cell cultures. Porcine EPO was purified as described elsewhere [15 ], was stored at 4ºC, and was used at concentrations of 50 nM. H2O2 concentrations were determined by formation of the ferric-SCN complex and visualized at 450 nm [34 ]. The JNK-APF plasmid was kindly obtained from Dr. Roger Davis (Howard Hughes Medical Institute, University of Massachusetts Medical School, Worcester) and was subcloned into pcDNA3 (Invitrogen, Carlsbad, CA). Purified bovine catalase was obtained from Worthington Chemical (Lakewood, NJ) and was used at a concentration of 2 mU/ml.

Immunoprecipitation and kinase assays
C10 cells were exposed to the test agents individually or simultaneously. At times ranging from 30 min to 5 h, cells were transferred to ice, washed once with cold phosphate-buffered saline (PBS), and lysed in buffer [50 mM HEPES, pH 6.5, 150 mM NaCl, 1 mM EDTA, 2 mM MgCl2, 10 mM Na3VO4, 1 mM phenylmethylsulfonyl fluoride, 0.1% Nonidet P-40, 10 µg/ml leupeptin, 1% aprotenin, 0.5 mM dithiothreitol (DTT), 100 µM NaF]. Lysates were cleared by centrifugation at 14,000 rpm, 4°C, for 10 min. Protein concentrations were determined with the Lowry method (Dc protein assay, BioRad Laboratories, Hercules, CA). ROCK-1 or JNK-1 was immunoprecipitated from 100 µg protein with a ROCK-1 (H-85, Santa Cruz Biotechnology, Santa Cruz, CA) or JNK-1 (C-17, Santa Cruz Biotechnology) antibody at 4°C for 1.5 h using protein G agarose beads (Gibco-BRL). Precipitates were washed twice with lysis buffer and once with kinase buffer containing 20 mM HEPES, pH 7.4, 20 mM ß-glycerolphosphate, 5 mM MgCl2, 2 mM NaF, 1 mM DTT for the ROCK-1 assay and 20 mM HEPES, pH 7.4, 20 mM ß-glycerolphosphate, 20 mM MgCL2, 2 mM DTT, and 0.1 mM Na3VO4 for the JNK assay. The kinase reaction was performed using 5 µg glutathione (GSH)-S-transferase (GST)-c-Jun or 1 µg purified recombinant human histone H1 (Upstate Biotechnology, Saranac Lake, NY) and 5 µCi {gamma}32P-adenosine 5'-triphosphate at 30°C for 30 min. Reactions were stopped by the addition of 2x Laemmli sample buffer (2% sodium dodecyl sulfate, 10% glycerol, 0.1 M DTT, and 0.01% bromophenol blue). Samples were boiled 5 min and stored at -20ºC. Proteins were separated on 15% polyacrylamide gels, and gels were dried and examined by autoradiography. Results were quantitated by phosphoimaging on a Molecular Imager FX (BioRad).

Western blotting
At selected times following exposure to test agents, cells were washed once with cold PBS and lysed and cleared as described previously. Protein (10 µg) was loaded onto polyacrylamide gels, electrophoresed, and transferred to nitrocellulose (Schleicher & Schuell, Keene, NH). Membranes were blocked in 5% milk in Tris-buffered saline. JNK-1 (FL, Santa Cruz Biotechnology), caspase 3 (Becton Dickinson, San Jose, CA), caspase 9 (H-170, Santa Cruz Biotechnology), and poly-ADP-ribose (Trevigen, Gaithersburg, MD) were detected as described elsewhere [35 ].

Cell death assay
Cell death was assessed by trypan blue exclusion by evaluating adherent and floating cells using a hemocytometer to count a minimum of 200 cells [36 ].

Immunofluorescence
Cells grown on glass coverslips were fixed in 2% paraformaldehyde (PFA) in PBS and permeabilized using 0.1% Triton X-100 in PBS for 15 min at room temperature (RT). Glass coverslips were washed in PBS, blocked three times for 20 min in 1% bovine serum albumin (BSA; Sigma-Aldrich, St. Louis, MO) in PBS, and then incubated overnight at 4°C with 500 µg/ml anticytochrome c (Becton Dickinson), anti-O2–tyrosine (Cayman Chemical, Ann Arbor, MI), or antiphospho-JNK (BioSource International, Camarillo, CA) in PBS/BSA. Coverslips were washed, blocked, and then incubated for 30 min at RT with 10 µg/ml Cy-3-conjugated anti-mouse immunoglobulin G (IgG; Jackson Immunoresearch, West Grove, PA). Nuclei were counterstained with 100 nM Sytox Green (Molecular Probes, Junction City, OR) for 15 min, and staining was evaluated using a BioRad MRC 1024ES confocal microscope and Lasersharp 2000 software (BioRad). Control experiments included incubation with secondary antibody only or with an isotype-control IgG1 and revealed minimal fluorescent staining (data not shown).

Mitochondrial respiration
Cells were grown on glass coverslips and treated as indicated. At various times, media was removed and replaced with media containing 150 nM freshly prepared Mitotracker Red and 500 nM Mitotracker Green (Molecular Probes) for 15 min at 37ºC in the dark. Hoechst 33342 (5 µg/ml; Sigma Chemical Co.) was added, followed by an additional 15-min incubation. Coverslips were washed 2x PBS, and images were collected using Metamorph software (Universal Imaging, Downingtown, PA) connected to an Olympus PM30 camera attached to an Olympus BX50 immunofluorescence microscope. Alternatively, cells were stained with the potentiometric dye JC-1 (2 µg/ml) for 10 min and visualized on a BioRad MRC 1240 confocal microscope to assess mitochondrial respiration.

GSH assay
The levels of reduced GSH were determined as described previously [37 ]. Briefly, cells were lysed, protein was precipitated, and supernatant was reacted with o-phtaldehyde. Fluorescence was measured using {lambda}ex = 320 nm and {lambda}em = 420 nm using a Hitachi F-4500 fluorescence spectrophotometer.

Visualization of membrane blebs
Membrane blebbing was assessed by phase-contrast microscopy using an Olympus BX50 microscope at 20x magnification.

Measurement of single-stranded DNA
Cells were fixed overnight in anhydrous methanol and assessed for DNA fragmentation. Cells were subsequently rinsed and boiled in MgCl2, blocked with FBS, and incubated with primary antibody (monoclonal antibody F7-26, Alexis Biochemicals, San Diego, CA). Cells were washed, and binding was detected by probing with horseradish peroxidase-conjugated secondary antibody and immunoperoxidase staining.

Generation of stable cell lines
C10 cells were plated in 60 mm dishes (1.5x105) and transfected with JNK-1-APF or pcDNA3.0 using lipofectamine 2000 (Gibco-BRL). Cultures were incubated in media containing 500 µg/ml G418, and resistant colonies were propagated as pools and tested for inhibition of JNK activation. Cultures were subsequently used for up to 3 weeks and then discarded.

Statistical analysis
Mean values were calculated by ANOVA using the Student-Newman-Kuels test to adjust for multiple pair-wise comparisons. An arscin square-root transformation of the percentages was used to stabilize the variance.

On-line supplemental material
After treatment (3.5 h) with 50 nM EPO, 15 mU/ml GOx, and 100 µM NO2-, time-lapse video microscopy was used to create Video 1 (Fig. 7video1.mov), which represents dynamic membrane blebbing induced by catalytically active EPO. Images were captured at 30-s intervals for 15 min. Phase-contrast images were collected with an Olympus PM30 digital camera using Metamorph software (Universal Imaging). Images were merged using Metamorph software and exported in QuickTime format.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Characterization of EPO catalytic activity
It was recently shown that the product of the EPO-catalyzed reaction of H2O2 and NO2- is ·NO2 [16 ]. Previous work in our laboratory has demonstrated that 5 ppm ·NO2 induces death selectively in log-phase epithelial cells [38 ]. We therefore sought to determine the mode of cell death in response to catalytically active EPO. Initial studies characterized the enzymatic activity of purified porcine EPO by measuring the levels of H2O2 in the media of cultures treated with EPO with substrates (Fig. 1 ). Glucose plus glucose oxidase (15 mU/ml) was used to generate a stable, catalase-inhibitable flux of H2O2, as was determined spectrophotometrically (Fig. 1A) . At physiological concentrations of 50 nM EPO and 100 µM of the substrate NO2-, EPO was able to completely consume the H2O2 in the media ranging from 15 min to 5 h (Fig. 1A) , and the H2O2 concentration never exceeded 20 µM, ensuring that H2O2 was not directly affecting the cells. Similar results were obtained when Br- and SCN- were added as substrate (data not shown). As a control, 2 mU/ml catalase was added to glucose oxidase-containing media, which mimicked the effects of EPO with regard to the consumption of H2O2. To confirm that EPO, in the presence of H2O2 and NO2-, catalyzes the production of ·NO2, we assessed tyrosine nitration, the stable end product of the reaction of ·NO2 with L-tyrosine [39 ], by immunofluorescence. Marked tyrosine nitration occurred in cells exposed to EPO/GOx/NO2- (Fig. 1B) . It is interesting that the majority of immunoreactivity occurred in the peripheral, membrane-proximal regions of the cells. Omission of NO2-, GOx, or EPO from the reaction mix did not result in tyrosine nitration, illustrating that the complete system was necessary to produce ·NO2.



View larger version (29K):
[in this window]
[in a new window]
  		Figure 1. Substrate use by EPO-GOx-NO2- system. (A) Consumption of H2O2 by EPO with substrates. C10 cells were exposed to the indicated conditions after which the media was collected, and the concentration of H2O2 was measured as described in Experimental Procedures. The concentrations used were EPO, 50 nM; GOx, 15 mU/ml; NO2-, 100 µM; catalase (Cat), 2 mU/ml. Results are representative of seven independent experiments. Values represent mean ± SD. -5 represents the addition of GOx to medium 5 min before addition to cells. (B) EPO catalyzes the formation of 3-nitrotyrosine. C10 cells were exposed to the indicated conditions for 2 h, and nitrotyrosine formation (white) was assessed by immunofluorescence.

 
EPO with substrates induces death in lung epithelial cells
To investigate whether the production of ·NO2 by EPO was associated with cell death, we examined the plasma membrane permeability of cells exposed to the EPO with substrates (Fig. 2 ). Exposure of cells to EPO with substrates resulted in ~90% cell death within 6 h. As all of the H2O2 was consumed from the media (Fig. 1A) , the cellular responses observed are not a result of H2O2 but likely of the products of EPO activity. Futhermore, the substrates NO2-, Br-, or SCN- alone did not cause cell death. The concern also existed that cell death may be the consequence of glucose depletion from the medium as a result of catalytic activity of GOx. The addition of catalase consumed all of the H2O2 produced (Fig. 1A) and prevented EPO/GOx-associated cell death (Fig. 2) . Furthermore, the pH remained constant during the course of the experiments, and the addition of glucose did not affect cell death caused by EPO with substrates (data not shown). We therefore can conclude that the toxicity of EPO is a result of the production of reactive oxidants. Previous investigations have suggested that supplementation of SCN- may provide some protection against the toxic effects of catalytically active EPO by driving product formation away from ·NO2 and HOBr and toward the formation of hypothiocyanite [31 ]. However, our results demonstrate that the toxicity of EPO was equivalent regardless of which substrate was present (Fig. 2) .



View larger version (10K):
[in this window]
[in a new window]
  		Figure 2. EPO with substrates causes cell death. C10 cells were treated with the indicated conditions, and cell death was measured with the trypan blue exclusion assay at 6 h post-treatment. The concentrations used were EPO, 50 nM; GOx, 15 mU/ml; NO2-, Br, or SCN-, 100 µM; catalase (Cat), 2 mU/ml. Results are the average of three separate experiments. Values represent mean ± SD.

 
Catalytically active EPO depletes cellular stores of reduced GSH
A common cellular target of reactive oxidant species, such as those generated by EPO, is the sulfhydryl group. To identify a common oxidant effect that could explain the lack of substrate specificity in the causation of cell death, we examined the effects of EPO in the presence of the substrates H2O2 + NO2-, Br-, or SCN- on levels of reduced GSH (Fig. 3 ). Following 2 h of exposure to EPO/GOx/NO2-, cells contained approximately half the concentration of GSH as compared with sham-treated cells. Similar effects were seen when Br- was used as cosubstrate. However, use of SCN- instead of NO2- or Br- was not associated with a marked decrease in cellular GSH by 2 h (Fig. 3) . Examination of GSH levels in response to EPO/GOx/SCN- after 4 h of exposure demonstrated that GSH levels were reduced to ~1.5 ng/µg cellular protein, illustrating that GSH depletion was delayed when SCN- was used as substrate, compared with the other substrates examined.



View larger version (13K):
[in this window]
[in a new window]
  		Figure 3. Cellular GSH content is decreased in cells exposed to EPO with substrates. C10 cells were exposed to 50 nM EPO, 15 mU/ml GOx, and 100 µM Br, NO2-, or SCN- for 2 h, and the levels of reduced GSH were measured fluorometrically using o-phthaldehyde (1 mg/ml) as the substrate. Fluorescence was measured at {lambda}ex = 320 nm and {lambda}em = 420 nm and was expressed per µg total cellular protein. Results represent mean ± SD.

 
Activation of JNK in response to catalytically active EPO
Oxidative stress is known to activate numerous stress-response pathways, including the stress-activated protein kinase (SAPK) pathway. We therefore examined the effects of EPO with and without substrates on the SAPK family member, JNK (Fig. 4 ). Addition of EPO/GOx/NO2- caused a rapid activation of JNK within 15 min that persisted up to 4 h (Fig. 4A) . It is important that this increase in JNK activity also occurred to a similar extent when Br- or SCN- was substituted for NO2- as substrates for EPO (Fig. 4B) , indicating that this response was not specific to ·NO2. In addition, EPO alone did not cause a significant increase in JNK activity, and the addition of catalase abolished JNK activity induced by EPO with substrates. The addition of 20 µM H2O2 failed to induce JNK activation. Collectively, these findings indicate that JNK activation was mediated by the oxidants generated by EPO and was not a result of its cationic charge or substrates alone. To further verify the activation of JNK, C10 cells grown on glass coverslips were exposed to EPO with substrates for 4 h; fixed and phosphorylated JNK was assessed by confocal microscopy. Results in Figure 4C demonstrate a marked increase in phospho-JNK content, depicting punctate membrane and nuclear localization in cells exposed to EPO with substrates. In contrast, EPO alone resulted in occasional cells containing phosphorylated JNK, consistent with the marginal JNK activity observed in Figure 4A .



View larger version (44K):
[in this window]
[in a new window]
  		Figure 4. Activation of JNK by the EPO-GOx-NO2- system. (A) Time course of JNK activation by EPO with substrates. C10 cells were treated with 50 nM EPO, 15 mU/ml GOx, and 100 µM NO2- for the indicated times, and JNK activity was assessed via immunoprecipitation of the JNK-1 complex from 100 µg protein and a kinase assay using the substrate GST-c-Jun. Results are representative of five experiments. (B) Activation of JNK by EPO with or without substrates. C10 cells were exposed to 50 nM EPO, 15 mU/ml GOx, and 100 µM Br, NO2-, or SCN- for 2 h, after which cells were lysed. Catalase (Cat) was used at 2 mU/ml, and H2O2 was used at 20 µM. Equal loading was ensured by immunoblotting 10 µg cell lysate for total JNK. (C) Subcellular localization of activated JNK. C10 cells were treated with 50 nM EPO, alone or in combination with 15 mU/ml GOx and 100 µM NO2- for 4 h, and were fixed in 1% PFA. JNK activity was assessed using an antibody specific to the dually phosphorylated form of JNK (upper), and nuclei were counterstained with propidium iodide (lower). (D) Thiol supplementation prevents the activation of stress responses induced by EPO with substrates. Cells were pretreated with N-acetyl-cysteine (NAc) for 16 h and washed, and JNK activity was assessed as described in A.

 
Thiol oxidation is one form of stress that has been shown to activate the SAPK pathway [25 ]. Pretreatment of cells with NAc for 16 h, which increases cellular reduced thiol content (ref. [40 ], and data not shown), resulted in a dose-dependent decrease in JNK activity caused by EPO with substrates (Fig. 4D) , illustrating that thiol oxidation events caused by EPO with substrates are essential to initiate signaling to JNK.

EPO with substrates causes mitochondrial perturbations
Activation of JNK has been linked to initiation of the mitochondrial apoptosis pathway in many cells [28 , 41 ]. We next examined the mitochondrial membrane potential of cells exposed to EPO/GOx/NO2- by incubating cells with Mitotracker green, which visualizes mitochondria, and Mitotracker red, which visualizes respiring mitochondria (Fig. 5A ). Mitochondrial membrane potential was lost within 1 h of exposure to EPO with substrates (Fig. 5A) and did not recover up to 6 h, indicating rapid and sustained loss of mitochondrial respiration (Fig. 5) . Following 6 h of treatment, mitochondrial membranes were disrupted, apparent from transmission electron micrographs (data not shown), and the nuclei of the cells were condensed (Fig. 5A , lower), evidenced by the Hoechst staining pattern. After establishing that treatment with EPO/GOx/NO2- resulted in mitochondrial membrane depolarization and disruption, we next determined if this effect was associated with the release of proapoptotic factors from the mitochondria. The release of cytochrome c is a well-known feature of some forms of apoptosis. Sham-treated cells exhibited bright, perinuclear cytochrome c staining, indicating that cytochrome c was retained in the mitochondria (Fig. 5B) . Cells treated with EPO/GOx/NO2- for 4 h exhibited little to no cytochrome c staining, demonstrating that cytochrome c was released from the mitochondria. The presence of catalase caused a partial rescue of the cytochrome c release caused by EPO with substrates (Fig. 5B) , illustrating that the catalytic activity of EPO is responsible for cytochrome c release.



View larger version (62K):
[in this window]
[in a new window]
  		Figure 5. EPO-dependent oxidative chemistry causes mitochondrial alterations in lung epithelial cells. (A) Assessment of mitochondrial depolarization. C10 cells were treated with 50 nM EPO, 15 mU/ml GOx, and 100 µM NO2- (EGN) or were sham-treated. Mitochondrial membrane potential was measured using the Mitotracker dyes (Molecular Probes). Mitotracker green stains all mitochondria regardless of potential, whereas Mitotracker red stains only actively respiring mitochondria. Nuclei were counterstained with Hoechst 33342 (lower), and live images were collected at 1 and 6 h post-treatment. (B) EPO with substrates (EGN) cause release of cytochrome c from mitochondria. C10 cells were treated 4 h, fixed in 4% PFA, permeabilized, and treated with anti-cytochrome c antibody, followed by Cy-3-conjugated secondary antibody and Sytox Green. Images were collected on a BioRad MRC 1240 confocal microscope. Red, Cytochrome c; green, nuclei.

 
Caspase cleavage and DNA damage following EPO exposure
Following the release of cytochrome c from mitochondria and formation of the apoptosome, caspase 9 becomes activated [42 ], which in turn activates caspase 3 via cleavage of procaspase 3 into the active 17 and 12 kDa fragments. Results in Figure 6A demonstrate that the levels of procaspase 9 were reduced in cells exposed to EPO/GOx/NO2- (or SCN-) at 5 h yet were not changed upon exposure to EPO, indicating cleavage to the active form by EPO with substrates. Furthermore, minor cleavage of procaspase 3 to the active p17 form was detected following 5 h of exposure to EPO + GOx in the presence of NO2- or SCN-. As these changes are minimal and occur at time points after other features of EPO-induced cell death occurred (Fig. 5) , it suggests that EPO/GOx/NO2- (or SCN-)-induced death may not involve the activity of executioner caspases and that caspase activation may be secondary to death induced by other mediators.



View larger version (21K):
[in this window]
[in a new window]
  		Figure 6. EPO with substrates induces caspase cleavage and DNA damage. (A) EPO with substrates induces cleavage of caspases 3 and 9. MTCC cells were treated with 50 nM EPO, 15 mU/ml GOx, and 100 µM NO2- or SCN- and were lysed upon the onset of membrane blebbing. Western blots were performed and probed with antibodies to caspases 3 and 9. (B) Protein ADP ribosylation by EPO with substrates. Poly-ADP-ribose polymerase (PARP) activity was assessed by immunoblotting against poly-ADP-ribosylated proteins following 1 h of treatment with 50 nM EPO, 15 mU/ml GOx, and 100 µM NO2- (EGN) or 100 µM H2O2. As a positive control (+Con), known ADP-ribosylated proteins from Escherichia coli were run in parallel with samples. (C) EPO causes DNA damage. C10 cells were treated with EPO and EPO plus substrates in the presence or absence of 2 mU/ml catalase (Cat) for 4 h, and DNA damage was assessed using an antibody to single-stranded DNA (apostain). *, P< 0.05, versus sham-treated cells; #, P< 0.05, versus EPO and EPO/GOx/Cat. Results are expressed as % of cells staining positive for single-stranded DNA ± SD.

 
To further define parameters of cell death induced by EPO with substrates, we assessed activation of PARP, which is induced by DNA damage and causes ADP-ribosylation of cellular proteins. PARP activity was restricted to constitutive auto-ADP-ribosylation (Fig. 6B) in control cells. Treatment of cells with EPO plus substrates or 100 µM H2O2, previously shown to activate PARP, caused enhanced ADP ribosylation of multiple proteins, indicating that oxidative stress induced by EPO with substrates causes DNA damage. To further assess DNA damage, the single-stranded DNA content of cells was assessed using an antibody to single-stranded DNA. C10 cells were treated for 4 h with EPO, EPO plus substrates, or EPO + GOx + catalase (Cat; Fig. 6C ) and were stained for single-stranded DNA. In response to EPO + GOx + NO2-, ~85% of cells contained single-standed DNA. It is interesting that EPO alone or EPO + GOx + catalase also resulted in significant single-stranded DNA reactivity, and ~40% of the cells labeled positive. These results demonstrate that DNA damage is an endpoint of exposure to EPO with substrates but is not causally linked to JNK activation, cytochrome c release, or membrane permeability, as these events are not observed in the absence of substrates.

Morphological changes induced by EPO with substrates
The preceding findings suggested that EPO with substrates was capable of initiating a death-inducing signaling cascade within lung epithelial cells. To investigate this in further detail, we examined the morphology of cells exposed to catalytically active EPO. Exposure of MTCC cells to EPO with substrates resulted in extensive and dynamic plasma membrane blebbing, initiating at approximately 3.5 h of treatment (Fig. 7 , and supplemental data). The blebs remained dynamic up to 5 h, at which time they no longer retracted and formed very large blisters on the cell surface that frequently pinched off from the plasma membrane and floated into the media. This phenomenon was not simply a response to oxidative stress induced by GOx alone (data not shown) and was not caused by the cationic charge of EPO, as it was not observed in cells exposed to EPO + GOx + catalase. We also observed that EPO + GOx, in combination with any of the three substrates tested (NO2-, Br-, SCN-), induced the identical phenotype, albeit with slightly different kinetics (data not shown).



View larger version (36K):
[in this window]
[in a new window]
  		Figure 7. EPO with substrates causes dynamic membrane blebbing. (A) MTCC cells were treated with the indicated conditions for 4 h, and live cells were imaged using phase-contrast microscopy. Conditions were 50 nM EPO; 15 mU/ml GOx; 100 µM NO2-; 2 mU/ml catalase (Cat). (B) Images from A were digitally captured, and the percentage of cells blebbing was quantified by counting at least 200 cells/dish. % Cells blebbing represents the number of cells blebbing divided by the total number of cells ± SD.

 
Involvement of ROCK-1 in EPO-dependent plasma membrane blebbing
Apoptotic membrane blebbing is mediated by the activity of ROCK-1. Constitutive ROCK-1 activity is thought to result from caspase 3-dependent cleavage of the inhibitory domain from p160 ROCK, resulting in the active p130 fragment. ROCK-1 induces membrane blebbing via phosphorylation of myosin light-chain kinase and myosin light-chain phosphatase, leading to acto-myosin-based contractility of the cortical actin cytoskeleton [43 , 44 ]. We used the specific ROCK-1 inhibitor Y-27632 [45 ] to investigate the role of ROCK-1 in EPO-induced membrane blebbing. Preincubation with Y-27632 resulted in a complete block of plasma membrane blebbing induced by EPO with substrates (EGN, Fig. 8A ). It is important that addition of Y-27632 did not block the catalytic activity of EPO (Fig. 8B) , indicating that the inhibition of blebbing is not a result of nonspecific inhibition of EPO. The kinase activity of ROCK-1 was assessed to determine the extent to which ROCK-1 activity was affected by EPO with substrates (Fig. 8C) . ROCK-1 kinase activity was significantly increased at 2 h (lane 2), and this activity remained elevated up to 5 h (lane 4) following treatment with EPO plus substrates. In addition, the response was equivalent at 5 h, regardless of whether NO2- (lane 4) or SCN- (lane 6) was used as the substrate. Treatment with Y-27632 completely abolished ROCK-1 activity in cells exposed to EPO with substrates, confirming its potency as an inhibitor. It is interesting that Y-27632 reduced ROCK-1 activity to levels below baseline, which may explain the abnormal phenotype seen in cells treated with Y-27632 alone (Fig. 8A) .



View larger version (39K):
[in this window]
[in a new window]
  		Figure 8. Involvement of ROCK-1 in EPO-dependent plasma membrane blebbing. (A) Inhibition of ROCK prevents membrane blebbing. MTCC cells were treated as indicated for 4 h, and membrane blebbing was imaged using phase-contrast microscopy. Y-27632 (10 µM) was added to cells 30 min before exposure with EPO plus substrates. (B) Y-27632 does not inhibit EPO. As a marker of EPO activity, H2O2 levels were assessed following 4 h of treatment as in Figure 1A . (C) EPO plus substrates induces sustained increases in ROCK-1 kinase activity. C10 cells were exposed for 2 h (lanes 1–3) or 5 h (lanes 4–6), after which cells were lysed, and ROCK-1 activity was assessed by phosphorylation of histone H1. (D) Y-27632 does not prevent cell death caused by EPO with substrates. Cells were treated for 5 h, after which permeability to trypan blue was determined. Values represent the mean ± SD. (E) Y-27632 does not prevent mitochondrial depolarization. C10 cells were treated for 3 h, after which mitochondrial membrane potential was determined as in Figure 5 . (F) Y-27632 does not inhibit JNK. JNK activity was determined following 1 h of treatment by in vitro kinase assay and quantified by phosphorimager analysis. Values represent the mean ± SD.

 
We next determined whether ROCK-1 kinase activity was important in cell death caused by EPO with substrates. Inhibition of ROCK-1 activity with Y-27632 was not able to suppress cell death induced by EPO with substrates (Fig. 8D) . Furthermore, treatment with Y-27632 did not rescue the loss of mitochondrial respiration in response to EPO with substrates (Fig. 8E) . Lastly, the use of Y-27632 did not interfere with the activity of JNK induced by EPO with substrates (Fig. 8F) . Collectively, these results indicate that ROCK-1 activity is not responsible for mediating mitochondrial membrane depolarization or overt membrane permeability and that its activation may be the consequence of JNK activity through mitochondrial depolarization and cytochrome c release.

Role of JNK in mediating cellular responses to EPO
Mitochondrial perturbations and apoptosis have been linked to JNK activity in various systems [30 ]. To investigate the importance of JNK activation in mediating the response to EPO/GOx/NO2-, we generated a cell line stably expressing a dominant-negative version of JNK (JNK-APF). We identified a pool of cells (lanes 5 and 6) that was refractory to JNK activation above baseline in response to EPO with substrates, as measured by phosphorylation of the substrate GST-c-Jun (Fig. 9A ). We then exposed these cells to EPO with substrates and evaluated the effects on plasma membrane blebbing (Fig. 9C and 9D ). JNK-APF-expressing pools exhibited significantly less (4.8±1.0%) blebbing cells than vector-transfected controls (45.4±4.0%) in response to EPO with substrates. The residual blebbing observed in JNK-APF-expressing cells exposed to EPO with substrates was abolished by the addition of Y-27632 (Fig. 9D) , consistent with its ability to completely inhibit ROCK kinase activity in cells exposed to EPO plus substrates (Fig. 8C) . These studies demonstrate that JNK activation is mediating signaling leading to plasma membrane blebbing associated with ROCK-1 activity.



View larger version (42K):
[in this window]
[in a new window]
  		Figure 9. Dominant-negative JNK suppresses membrane blebbing and prevents mitochondrial depolarization. (A) Assessment of JNK activity in parental cells, pcDNA3-transfected cells, or JNK-APF-transfected cells. C10 cells expressing the indicated constructs were left untreated or treated with 50 nM EPO, 15 mU/ml GOx, and 100 µM NO2-, and JNK activity was measured as in Figure 4 . (B) Assessment of mitochondrial membrane potential in pcDNA3 or JNK-APF-transfected cells. Cells were treated with EPO/GOx/NO2- for 1 h, and mitochondrial membrane potential was measured using the potentiometric dye JC-1, as described in Figure 5A . (C) Visualization of membrane blebbing in vector or JNK-APF-transfected cells. Cells were treated with EPO/GOx/NO2- (as in A) for 4.5 h and imaged live using dark-field microscopy. (D) Images from C were quantified by counting at least 200 cells/treatment and are represented as the mean ± SD.

 
We next investigated the role of JNK in mediating mitochondrial membrane depolarization, which is known to occur upstream of activation of caspase 3 and as we have shown, the activation of ROCK-1 [43 , 44 ]. Cells were exposed to EPO with substrates for 1 h, and the mitochondrial membrane potential was measured with the potentiometric dye JC-1 (Fig. 9B) . A marked decrease in mitochondrial membrane potential can be seen in vector-transfected cells when exposed to EPO with substrates, and this effect was virtually abrogated in cells expressing JNK-APF. These results demonstrate that JNK activation is required for depolarization of mitochondria in response to EPO with substrates and subsequent signaling cascades that culminate in dynamic membrane blebbing associated with ROCK-1 activation.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The recent observation that EPO with substrates is capable of forming high levels of ·NO2 stresses the potential involvement of this highly reactive, free-radical gas in the pathobiology of inflammatory diseases and also suggests that ·NO2 formation may be an important mediator of eosinophil effecter function in vivo [16 ]. In fact, in response to immunization and challenge with OVA, EPO–KO mice exhibit significantly decreased levels of nitrotyrosine, the reaction product of ·NO2 with L-tyrosine, compared with identically treated wild-type mice [9 ], illustrating that EPO catalyzes the formation of this reactive oxidant in the lung during physiological stress. In addition, inflammatory airway disease is associated with marked increases in levels of bromotyrosine, the stable end product of L-tyrosine attack by HOBr. As HOBr is formed specifically by EPO and not other peroxidases, in vivo bromotyrosine is considered a sensitive marker of EPO catalytic activity [18 ]. The results presented here demonstrate that at physiological concentrations of EPO and its required substrates, reactive oxidants generated by EPO are capable of inducing signaling cascades and extensive damage to lung epithelial cells in vitro.

The effect of ·NO2 on lung epithelial cells was first characterized by directly exposing cells to gas-phase ·NO2, which revealed the occurrence of death selectively in log-phase cells [38 ], whereas quiescent cells were highly resistant. In this study, we illustrate that the patterns of cell death in response to a more physiological source of ·NO2, generated by EPO with substrates, were distinct from cell death induced by pure ·NO2 gas. Whereas extensive membrane blebbing is observed in response to ·NO2 generated by EPO with substrates, membrane blebbing is not observed in response to pure ·NO2 gas [38 ]. The reason for the observed discrepancies between ·NO2 generated by EPO versus pure ·NO2 gas may be the cationic charge of EPO, which could contribute to the toxicity of ·NO2 by specific targeting of ·NO2 to the cell surface or by activating additional signaling pathways. Furthermore, it is possible that the generation of other oxidants by active EPO may also account for the different phenotype.

The substitution of SCN- for NO2- or Br- has been proposed as a mechanism to alleviate the injurious effects of EPO [31 ] by preventing the formation of the highly toxic ·NO2 or HOBr by instead promoting generation of the less toxic gases hypothiocyanite and cyanate. The results of our experiments do not support these earlier findings. The use of SCN- in place of NO2- induced identical responses in numerous experiments. It is worth noting, however, that the toxicity of SCN- may appear decreased at early times, as the onset of overt cytotoxic effects and membrane blebbing generally occurred along a more protracted time frame, compared with Br- or NO2-. A plausible explanation for our findings showing the lack of substrate specificity in the cellular responses to active EPO may be the observed loss of GSH content, likely a result of thiol oxidation [46 ], which we demonstrated to be causally linked to JNK activation.

The activation of JNK in response to oxidative stress [28 , 47 ] has been causally linked to the induction of apoptosis [25 , 28 , 48 , 49 ], yet the exact mechanism of how this occurs has not been elucidated. JNK activity has been linked to opening of the mitochondrial permeability pore [50 ] and the subsequent release of cytochrome c [25 ]. Our results show that EPO with substrates causes mitochondrial membrane depolarization and the release of cytochrome c. In fact, mitochondrial membrane depolarization induced by active EPO is dependent on active JNK, as the inhibition of JNK by the stable expression of a dominant-negative construct blocked this event.

The activation of a signaling cascade in lung epithelial cells exposed to EPO with substrates, which culminates in features reminiscent of apoptosis, such as caspase cleavage and dynamic membrane blebbing, constitutes an intrinsic, protective response against certain forms of oxidative stress, where cells can sense a toxic level of oxidative stress and delay necrosis. The activation of ROCK-1, shown in the present study, which occurs during apoptosis, is required for the formation of membrane-bound autophagic vesicles [43 , 44 ], which can be recognized and internalized by phagocytic cells [51 ]. This process is associated with a reduction of inflammation by preventing the release of cellular contents, which are known to cause a marked inflammatory response [52 ]. However, the mode of cell death triggered by EPO with substrates in the lung epithelial cells examined here is not strictly apoptotic, as caspase cleavage was modest and membrane permeability, a feature of necrosis, occurred to a significant extent.

In summary, we have demonstrated that EPO, in the presence of physiological concentrations of its required substrates, triggers a signaling cascade in lung epithelial cells that culminates in cell death. This cascade, which depends on the activation of JNK and consequently, mitochondrial perturbations, culminates in dynamic membrane blebbing and is observed independently of which substrate is present. Whether these findings have ramifications for the pathobiology of inflammatory lung diseases that are associated with airway injury and remodeling remains to be determined through the evaluation of EPO–KO mice.


   ACKNOWLEDGEMENTS
 
National Institutes of Health RO1 HL60014 and PO1 HL67004 funded this work (Y. M. W. J-H.). NIH Grant RR P20 RL15557 from the Centers of Biomedical Research Excellence (COBRE) program of the National Center for Research Resources also made this publication possible. Its contents are solely the responsibility of the authors and do not necessarily represent the official views of NIH. The authors thank Dr. Pamela Vacek for her excellent assistance with statistical analyses, Dr. Charles Irvin for his scientific advice, and Marleen Harink for technical assistance.

Received January 17, 2003; accepted July 18, 2003.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Wu, W., Samoszuk, M. K., Comhair, S. A., Thomassen, M. J., Farver, C. F., Dweik, R. A., Kavuru, M. S., Erzurum, S. C., Hazen, S. L. (2000) Eosinophils generate brominating oxidants in allergen-induced asthma J. Clin. Invest. 105,1455-1463[Medline]
  2. Holgate, S. T. (1999) The epidemic of allergy and asthma Nature 402,B2-B4[CrossRef][Medline]
  3. Bochner, B. S., Undem, B. J., Lichtenstein, L. M. (1994) Immunological aspects of allergic asthma Annu. Rev. Immunol. 12,295-335[CrossRef][Medline]
  4. Rothenberg, M. E. (1998) Eosinophilia N. Engl. J. Med. 338,1592-1600[Free Full Text]
  5. Benayoun, L., Druilhe, A., Dombret, M. C., Aubier, M., Pretolani, M. (2003) Airway structural alterations selectively associated with severe asthma Am. J. Respir. Crit. Care Med. 167,1360-1368[Abstract/Free Full Text]
  6. Agosti, J. M., Altman, L. C., Ayars, G. H., Loegering, D. A., Gleich, G. J., Klebanoff, S. J. (1987) The injurious effect of eosinophil peroxidase, hydrogen peroxide, and halides on pneumocytes in vitro J. Allergy Clin. Immunol. 79,496-504[CrossRef][Medline]
  7. Barnes, P. J., Chung, K. F., Page, C. P. (1998) Inflammatory mediators of asthma: an update Pharmacol. Rev. 50,515-596[Abstract/Free Full Text]
  8. Erjefalt, J. S., Persson, C. G. (2000) New aspects of degranulation and fates of airway mucosal eosinophils Am. J. Respir. Crit. Care Med. 161,2074-2085[Free Full Text]
  9. Denzler, K. L., Borchers, M. T., Crosby, J. R., Cieslewicz, G., Hines, E. M., Justice, J. P., Cormier, S. A., Lindenberger, K. A., Song, W., Wu, W., et al (2001) Extensive eosinophil degranulation and peroxidase-mediated oxidation of airway proteins do not occur in a mouse ovalbumin-challenge model of pulmonary inflammation J. Immunol. 167,1672-1682[Abstract/Free Full Text]
  10. Giembycz, M. A., Lindsay, M. A. (1999) Pharmacology of the eosinophil Pharmacol. Rev. 51,213-340[Abstract/Free Full Text]
  11. Jong, E. C., Henderson, W. R., Klebanoff, S. J. (1980) Bactericidal activity of eosinophil peroxidase J. Immunol. 124,1378-1382[Abstract]
  12. Reeves, E. P., Lu, H., Jacobs, H. L., Messina, C. G., Bolsover, S., Gabella, G., Potma, E. O., Warley, A., Roes, J., Segal, A. W. (2002) Killing activity of neutrophils is mediated through activation of proteases by K+ flux Nature 416,291-297[CrossRef][Medline]
  13. Schraufstatter, I., Hyslop, P. A., Jackson, J. H., Cochrane, C. G. (1988) Oxidant-induced DNA damage of target cells J. Clin. Invest. 82,1040-1050
  14. Slungaard, A., Vercellotti, G. M., Walker, G., Nelson, R. D., Jacob, H. S. (1990) Tumor necrosis factor alpha/cachectin stimulates eosinophil oxidant production and toxicity towards human endothelium J. Exp. Med. 171,2025-2041[Abstract/Free Full Text]
  15. MacPherson, J. C., Comhair, S. A., Erzurum, S. C., Klein, D. F., Lipscomb, M. F., Kavuru, M. S., Samoszuk, M. K., Hazen, S. L. (2001) Eosinophils are a major source of nitric oxide-derived oxidants in severe asthma: characterization of pathways available to eosinophils for generating reactive nitrogen species J. Immunol. 166,5763-5772[Abstract/Free Full Text]
  16. Brennan, M. L., Wu, W., Fu, X., Shen, Z., Song, W., Frost, H., Vadseth, C., Narine, L., Lenkiewicz, E., Borchers, M. T., et al (2002) A tale of two controversies.Defining both the role of peroxidases in nitrotyrosine formation in vivo using eosinophil peroxidase and myeloperoxidase-deficient mice, and the nature of peroxidase-generated reactive nitrogen species J. Biol. Chem. 277,17415-17427[Abstract/Free Full Text]
  17. Wu, W., Chen, Y., Hazen, S. L. (1999) Eosinophil peroxidase nitrates protein tyrosyl residues. Implications for oxidative damage by nitrating intermediates in eosinophilic inflammatory disorders J. Biol. Chem. 274,25933-25944[Abstract/Free Full Text]
  18. Wu, W., Chen, Y., d'Avignon, A., Hazen, S. L. (1999) 3-Bromotyrosine and 3,5-dibromotyrosine are major products of protein oxidation by eosinophil peroxidase: potential markers for eosinophil-dependent tissue injury in vivo Biochemistry 38,3538-3548[CrossRef][Medline]
  19. Kaminsky, D. A., Mitchell, J., Carroll, N., James, A., Soultanakis, R., Janssen, Y. (1999) Nitrotyrosine formation in the airways and lung parenchyma of patients with asthma J. Allergy Clin. Immunol. 104,747-754[CrossRef][Medline]
  20. Kim, D. K., Cho, E. S., Um, H. D. (2000) Caspase-dependent and -independent events in apoptosis induced by hydrogen peroxide Exp. Cell Res. 257,82-88[CrossRef][Medline]
  21. Gotoh, Y., Cooper, J. A. (1998) Reactive oxygen species- and dimerization-induced activation of apoptosis signal-regulating kinase 1 in tumor necrosis factor-alpha signal transduction J. Biol. Chem. 273,17477-17482[Abstract/Free Full Text]
  22. Lee, Y. J., Shacter, E. (1999) Oxidative stress inhibits apoptosis in human lymphoma cells J. Biol. Chem. 274,19792-19798[Abstract/Free Full Text]
  23. Vercammen, D., Beyaert, R., Denecker, G., Goossens, V., Van Loo, G., Declercq, W., Grooten, J., Fiers, W., Vandenabeele, P. (1998) Inhibition of caspases increases the sensitivity of L929 cells to necrosis mediated by tumor necrosis factor J. Exp. Med. 187,1477-1485[Abstract/Free Full Text]
  24. Reed, J. C., Green, D. R. (2002) Remodeling for demolition: changes in mitochondrial ultrastructure during apoptosis Mol. Cell 9,1-3[CrossRef][Medline]
  25. Aoki, H., Kang, P. M., Hampe, J., Yoshimura, K., Noma, T., Matsuzaki, M., Izumo, S. (2002) Direct activation of mitochondrial apoptosis machinery by c-Jun N-terminal kinase in adult cardiac myocytes J. Biol. Chem. 277,10244-10250[Abstract/Free Full Text]
  26. Rustin, P. (2002) Mitochondria, from cell death to proliferation Nat. Genet. 30,352-353[CrossRef][Medline]
  27. Ankarcrona, M., Dypbukt, J. M., Bonfoco, E., Zhivotovsky, B., Orrenius, S., Lipton, S. A., Nicotera, P. (1995) Glutamate-induced neuronal death: a succession of necrosis or apoptosis depending on mitochondrial function Neuron 15,961-973[CrossRef][Medline]
  28. Tournier, C., Hess, P., Yang, D. D., Xu, J., Turner, T. K., Nimnual, A., Bar-Sagi, D., Jones, S. N., Flavell, R. A., Davis, R. J. (2000) Requirement of JNK for stress-induced activation of the cytochrome c-mediated death pathway Science 288,870-874[Abstract/Free Full Text]
  29. Li, P., Nijhawan, D., Budihardjo, I., Srinivasula, S. M., Ahmad, M., Alnemri, E. S., Wang, X. (1997) Cytochrome c and dATP-dependent formation of Apaf-1/caspase-9 complex initiates an apoptotic protease cascade Cell 91,479-489[CrossRef][Medline]
  30. Davis, R. J. (2000) Signal transduction by the JNK group of MAP kinases Cell 103,239-252[CrossRef][Medline]
  31. Slungaard, A., Mahoney, J. R., Jr (1991) Thiocyanate is the major substrate for eosinophil peroxidase in physiologic fluids.Implications for cytotoxicity J. Biol. Chem. 266,4903-4910[Abstract/Free Full Text]
  32. Malkinson, A. M., Dwyer-Nield, L. D., Rice, P. L., Dinsdale, D. (1997) Mouse lung epithelial cell lines-tools for the study of differentiation and the neoplastic phenotype Toxicology 123,53-100[CrossRef][Medline]
  33. Magdaleno, S. M., Wang, G., Mireles, V. L., Ray, M. K., Finegold, M. J., DeMayo, F. J. (1997) Cyclin-dependent kinase inhibitor expression in pulmonary Clara cells transformed with SV40 large T antigen in transgenic mice Cell Growth Differ 8,145-155[Abstract]
  34. van der Vliet, A., Eiserich, J. P., Halliwell, B., Cross, C. E. (1997) Formation of reactive nitrogen species during peroxidase-catalyzed oxidation of nitrite.A potential additional mechanism of nitric oxide-dependent toxicity J. Biol. Chem. 272,7617-7625[Abstract/Free Full Text]
  35. Janssen, Y. M., Matalon, S., Mossman, B. T. (1997) Differential induction of c-fos, c-jun, and apoptosis in lung epithelial cells exposed to ROS or RNS Am. J. Physiol. 273,L789-L796
  36. Englert, R. P., Shacter, E. (2002) Distinct modes of cell death induced by different reactive oxygen species: amino acyl chloramines mediate hypochlorous acid-induced apoptosis J. Biol. Chem. 277,20518-20526[Abstract/Free Full Text]
  37. Cohn, V. H., Lyle, J. (1966) A fluorometric assay for glutathione Anal. Biochem. 14,434-440[CrossRef][Medline]
  38. Persinger, R. L., Blay, W. M., Heintz, N. H., Hemenway, D. R., Janssen-Heininger, Y. M. (2001) Nitrogen dioxide induces death in lung epithelial cells in a density-dependent manner Am. J. Respir. Cell Mol. Biol. 24,583-590[Abstract/Free Full Text]
  39. Espey, M. G., Xavier, S., Thomas, D. D., Miranda, K. M., Wink, D. A. (2002) Direct real-time evaluation of nitration with green fluorescent protein in solution and within human cells reveals the impact of nitrogen dioxide vs. peroxynitrite mechanisms Proc. Natl. Acad. Sci. USA 99,3481-3486[Abstract/Free Full Text]
  40. Janssen, Y. M., Heintz, N. H., Mossman, B. T. (1995) Induction of c-fos and c-jun proto-oncogene expression by asbestos is ameliorated by N-acetyl-L-cysteine in mesothelial cells Cancer Res 55,2085-2089[Abstract/Free Full Text]
  41. Harris, C. A., Johnson, E. M., Jr (2001) BH3-only Bcl-2 family members are coordinately regulated by the JNK pathway and require Bax to induce apoptosis in neurons J. Biol. Chem. 276,37754-37760[Abstract/Free Full Text]
  42. Budd, R. C. (2001) Activation-induced cell death Curr. Opin. Immunol. 13,356-362[CrossRef][Medline]
  43. Coleman, M. L., Sahai, E. A., Yeo, M., Bosch, M., Dewar, A., Olson, M. F. (2001) Membrane blebbing during apoptosis results from caspase-mediated activation of ROCK I Nat. Cell Biol. 3,339-345[CrossRef][Medline]
  44. Sebbagh, M., Renvoize, C., Hamelin, J., Riche, N., Bertoglio, J., Breard, J. (2001) Caspase-3-mediated cleavage of ROCK I induces MLC phosphorylation and apoptotic membrane blebbing Nat. Cell Biol. 3,346-352[CrossRef][Medline]
  45. Uehata, M., Ishizaki, T., Satoh, H., Ono, T., Kawahara, T., Morishita, T., Tamakawa, H., Yamagami, K., Inui, J., Maekawa, M., Narumiya, S. (1997) Calcium sensitization of smooth muscle mediated by a Rho-associated protein kinase in hypertension Nature 389,990-994[CrossRef][Medline]
  46. Arlandson, M., Decker, T., Roongta, V. A., Bonilla, L., Mayo, K. H., MacPherson, J. C., Hazen, S. L., Slungaard, A. (2001) Eosinophil peroxidase oxidation of thiocyanate.Characterization of major reaction products and a potential sulfhydryl-targeted cytotoxicity system J. Biol. Chem. 276,215-224[Abstract/Free Full Text]
  47. Janssen-Heininger, Y. M., Macara, I., Mossman, B. T. (1999) Cooperativity between oxidants and tumor necrosis factor in the activation of nuclear factor (NF)-kappaB: requirement of Ras/mitogen-activated protein kinases in the activation of NF-kappaB by oxidants Am. J. Respir. Cell Mol. Biol. 20,942-952[Abstract/Free Full Text]
  48. Wang, X., Martindale, J. L., Liu, Y., Holbrook, N. J. (1998) The cellular response to oxidative stress: influences of mitogen-activated protein kinase signalling pathways on cell survival Biochem. J. 333,291-300
  49. Benhar, M., Dalyot, I., Engelberg, D., Levitzki, A. (2001) Enhanced ROS production in oncogenically transformed cells potentiates c-Jun N-terminal kinase and p38 mitogen-activated protein kinase activation and sensitization to genotoxic stress Mol. Cell. Biol. 21,6913-6926[Abstract/Free Full Text]
  50. Gibson, E. M., Henson, E. S., Villanueva, J., Gibson, S. B. (2002) MEK kinase 1 induces mitochondrial permeability transition leading to apoptosis independent of cytochrome c release J. Biol. Chem. 277,10573-10580[Abstract/Free Full Text]
  51. Hanayama, R., Tanaka, M., Miwa, K., Shinohara, A., Iwamatsu, A., Nagata, S. (2002) Identification of a factor that links apoptotic cells to phagocytes Nature 417,182-187[CrossRef][Medline]
  52. Li, M., Carpio, D. F., Zheng, Y., Bruzzo, P., Singh, V., Ouaaz, F., Medzhitov, R. M., Beg, A. A. (2001) An essential role of the NF-kappa B/Toll-like receptor pathway in induction of inflammatory and tissue-repair gene expression by necrotic cells J. Immunol. 166,7128-7135[Abstract/Free Full Text]



This article has been cited by other articles:


Home page
 Am. J. Respir. Cell Mol. Bio.Home page
C. Pantano, V. Anathy, P. Ranjan, N. H. Heintz, and Y. M. W. Janssen-Heininger
Nonphagocytic Oxidase 1 Causes Death in Lung Epithelial Cells via a TNF-RI-JNK Signaling Axis
Am. J. Respir. Cell Mol. Biol., April 1, 2007; 36(4): 473 - 479.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Lung Cell. Mol. Physiol.Home page
M. E. Poynter, R. L. Persinger, C. G. Irvin, K. J. Butnor, H. van Hirtum, W. Blay, N. H. Heintz, J. Robbins, D. Hemenway, D. J. Taatjes, et al.
Nitrogen dioxide enhances allergic airway inflammation and hyperresponsiveness in the mouse
Am J Physiol Lung Cell Mol Physiol, January 1, 2006; 290(1): L144 - L152.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
S. Kumar, L. Gupta, Y. S. Han, and C. Barillas-Mury
Inducible Peroxidases Mediate Nitration of Anopheles Midgut Cells Undergoing Apoptosis in Response to Plasmodium Invasion
J. Biol. Chem., December 17, 2004; 279(51): 53475 - 53482.
[Abstract] [Full Text] [PDF]

Home page
 Mol. Cell. Biol.Home page
P. Shrivastava, C. Pantano, R. Watkin, B. McElhinney, A. Guala, M. L. Poynter, R. L. Persinger, R. Budd, and Y. Janssen-Heininger
Reactive Nitrogen Species-Induced Cell Death Requires Fas-Dependent Activation of c-Jun N-Terminal Kinase
Mol. Cell. Biol., August 1, 2004; 24(15): 6763 - 6772.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available