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Originally published In Press as doi:10.1074/jbc.M107231200 on October 31, 2001

J. Biol. Chem., Vol. 277, Issue 2, 1332-1339, January 11, 2002
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ERK-1/2 and p38 Kinase Oppositely Regulate Nitric Oxide-induced Apoptosis of Chondrocytes in Association with p53, Caspase-3, and Differentiation Status*

Song-Ja KimDagger §, Jung-Won JuDagger , Chun-Do OhDagger §, Young-Mee YoonDagger §, Woo Keun SongDagger , Jae-Hong KimDagger , Yung Joon YooDagger , Ok-Sun Bang||, Shin-Sung Kang||, and Jang-Soo ChunDagger **

From the Dagger  National Research Laboratory, Department of Life Science, Kwangju Institute of Science and Technology, Kwangju 500-712, Korea and the || Department of Biology, Kyungpook National University, Taegu 702-701, Korea

Received for publication, July 30, 2001, and in revised form, September 20, 2001


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Nitric oxide regulates cartilage destruction by causing dedifferentiation and apoptosis of chondrocytes. We investigated the role of the mitogen-activated protein kinase subtypes, extracellular signal-regulated protein kinase (ERK)-1/2, and p38 kinase in NO-induced apoptosis of rabbit articular chondrocytes and their involvement in dedifferentiation. Generation of NO with sodium nitroprusside (SNP) caused dedifferentiation, as indicated by the inhibition of type II collagen expression and proteoglycan synthesis. NO additionally caused apoptosis, accompanied by p53 accumulation and caspase-3 activation. SNP treatment stimulated activation of ERK-1/2 and p38 kinase. Inhibition of ERK-1/2 with PD98059 rescued SNP-induced dedifferentiation but enhanced apoptosis up to 2-fold, whereas inhibition of p38 kinase with SB203580 enhanced dedifferentiation, with significant blockage of apoptosis. The stimulation of apoptosis by ERK inhibition was accompanied by increased p53 accumulation and caspase-3 activity, whereas the inhibitory effect of p38 kinase blockade was associated with reduced p53 accumulation and caspase-3 activity. Our results indicate that NO-induced p38 kinase functions as an induction signal for apoptosis and in the maintenance of chondrocyte phenotype, whereas ERK activity causes dedifferentiation and operates as an anti-apoptotic signal. NO generation is less proapoptotic in chondrocytes that are dedifferentiated by serial monolayer culture or phorbol ester treatment. NO-induced p38 kinase activity is low in dedifferentiated cells compared with that in differentiated chondrocytes, with lower levels of p53 accumulation and caspase-3 activity. Our findings collectively suggest that ERK-1/2 and p38 kinase oppositely regulate NO-induced apoptosis of chondrocytes, in association with p53 accumulation, caspase-3 activation, and differentiation status.


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Chondrocytes in cartilage are differentiated from mesenchymal cells during embryonic development. Differentiated chondrocytes, which are the only cell type found in normal mature cartilage, synthesize sufficient amounts of cartilage-specific extracellular matrix (ECM)1 to maintain matrix integrity (1). This homeostasis is destroyed in degenerative diseases, such as osteoarthritis and rheumatoid arthritis. Arthritis is characterized by structural and biochemical changes in chondrocytes and cartilage, including degradation of the cartilage matrix, insufficient synthesis of ECM because of loss of chondrocyte phenotype (i.e. dedifferentiation), and increased numbers of apoptotic chondrocytes (2-6).

It is now generally accepted that proinflammatory cytokines, such as interleukin-1beta and tumor necrosis factor-alpha , play a predominant role in structural and biochemical alterations in chondrocytes and cartilage (6, 7). One of the mechanisms by which cytokines elicit their effects involves the stimulation of NO production via inducible NO synthase (iNOS) (7-9). NO regulates chondrocyte and cartilage function in various ways. These include: 1) inhibition of the synthesis of cartilage-specific ECM molecules, such as type II collagen and proteoglycan by triggering dedifferentiation of chondrocytes (10, 11); 2) an increase in the number of apoptotic cells, which correlates with the extent of cartilage matrix loss (12-14); and 3) modulation of matrix metalloproteinases to cause degradation of cartilage matrix (15). Indeed, iNOS inhibitors protect against degeneration of cartilage and chondrocytes in a number of experimental models. For instance, in experimentally induced osteoarthritis in different animal species, significant correlation was observed between the level of NO production and prevalence of apoptotic cells in cartilage tissue (12, 13), and administration of an iNOS inhibitor resulted in reduced articular cartilage damage and apoptotic cell death (16). In addition, NO generation by the NO donor, sodium nitroprusside (SNP), induced apoptosis in cultured articular chondrocytes (14, 17). However, the detailed mechanism of NO-induced chondrocyte apoptosis remains to be elucidated.

A group of cysteine proteases denoted "caspases" play a central role in apoptosis. Caspases are synthesized as proenzymes that are activated by cleaving the prodomain at a specific aspartic acid cleavage site. One of the upstream signaling pathways leading to caspase activation involves the release of cytochrome c and other apoptogenic factors from injured mitochondria (18, 19). Cytochrome c release and caspase activation are mediated by the translocation of cytosolic Bax (a proapoptotic member of Bcl-2 family) to mitochondria in response to various apoptotic stimuli (20-23). Several recent studies indicate that mitochondrial integrity is regulated by a tumor suppressor protein, p53 (24-26). The function of p53 in apoptosis regulation is modulated by multiple phosphorylation of several different protein kinases, including the major subtypes of mitogen-activated protein (MAP) kinase, i.e. extracellular signal-regulated kinase (ERK)-1 and -2, c-Jun N-terminal kinase, and p38 kinase (27-31). Although previous experiments show that NO in chondrocytes activates caspases to induce apoptosis (17, 32), to date, no direct evidence identifies a role for p53 and its regulation by MAP kinase in chondrocyte apoptosis.

The MAP kinase subtypes are activated by proinflammatory cytokines, which additionally generate NO, suggesting the role for NO in MAP kinase signaling. In fact, treatment with NO donors activates MAP kinase subtypes (33). Among the MAP kinase subtypes, ERKs are activated in response to mitogen or growth factor stimulation, whereas c-Jun N-terminal kinase and p38 kinases are stimulated during cellular stress conditions. The issue of whether MAP kinase activation determines cell survival or death currently remains controversial. Several studies indicate that c-Jun N-terminal kinase and p38 kinase activation processes are associated with apoptosis, whereas ERK activation is coupled with cell survival. Thus, coordinated activation and interactions allow cells to respond to various genotoxic and survival factors by affecting a number of downstream targets (34-37).

Although it is evident that NO production in chondrocytes causes apoptotic cell death, the underlying mechanism of this process has not been well characterized so far. In this study, we investigate the signaling cascade during NO-induced cell death in rabbit articular chondrocytes. Using SNP as a NO donor to examine the direct effects of NO generation, we initially describe the role of MAP kinase subtypes, ERK-1/2 and p38 kinase, activated by NO. The investigation additionally focuses on the characterization of the role of MAP kinase subtypes in caspase-3 regulation (an executioner of apoptosis) and p53 (a known signaling molecule upstream of caspase-3), both of which respond to various apoptotic stimuli. We additionally evaluate the relationship between apoptosis and dedifferentiation of chondrocytes, two processes simultaneously initiated by NO in the mature cartilage cells. We report here that ERK-1/2 and p38 kinase oppositely regulate NO-induced dedifferentiation and apoptosis of chondrocytes and that the reverse effects of the two MAP kinase proteins on apoptosis are triggered by the converse regulation of p53 and caspase-3 in a differentiation status-dependent pathway.

    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Monolayer Culture of Rabbit Articular Chondrocytes and Experimental Culture Condition-- Rabbit articular chondrocytes were released from cartilage slices of 2-week-old New Zealand White rabbits by enzymatic digestion. To summarize, cartilage slices were aseptically dissected and then dissociated enzymatically for 6 h in 0.2% collagenase type II (381 units/mg solid; Sigma) in phosphate-buffered saline. Individual cells were then obtained by collecting the supernatant after brief centrifugation. The cells were resuspended in Dulbecco's modified Eagle's medium (Invitrogen) supplemented with 10% (v/v) fetal bovine calf serum, 50 µg/ml streptomycin, and 50 units/ml penicillin, after which they were plated on culture dishes at a density of 5 × 104 cells/cm2. The medium was changed every 2 days after seeding, and the cells reached confluence in ~5 days. The primary culture before confluence, designated as passage (P) 0, was subcultured up to P4 by plating cells at a density of 5 × 104 cells/cm2 in each subculture.

SNP was used as a NO donor. In the experiments measuring SNP dose response, primary chondrocytes were treated with various concentrations of SNP for 24 h in Dulbecco's modified Eagle's medium containing 10% fetal calf serum. The time course of the SNP response was determined by incubating cells with 1 mM SNP for the indicated time period (0-48 h). To explore the signaling cascade on SNP-induced dedifferentiation and apoptosis, we used 100 µM caspase-3 inhibitor z-Asp-Glu-Val-Asp-fluoromethyl ketone (Bachem, Heidelberg, Germany), 100 µM pan-caspase inhibitor z-Val-Ala-Asp (O-methyl)-fluoromethyl ketone (Bachem), MEK-1/2 inhibitor PD98059, and p38 kinase inhibitor SB203580. Chondrocytes were preincubated with each inhibitor for 30 min, followed by coincubation with SNP.

Determination of Chondrocyte Differentiation Status-- Loss of chondrocyte phenotype, dedifferentiation, was determined by examining the accumulation of sulfated glycosaminoglycan with Alcian Blue, as described previously (38, 39), or by the expression of type II collagen studied by Western or Northern blot analysis. To summarize, type II collagen expression was detected using antibodies purchased from Chemicon (Temecula, CA) by Western blot analysis, as described below. For Northern blot analysis of type II collagen, total RNA was isolated using RNA STAT-60 (TEL-TEST, Inc., Friendswood, TX). In this analysis 5 µg/lane of RNA was denatured and fractionated on formaldehyde/agarose gels, and the separated RNA was transferred to S & S Nytran N nylon membranes. Prehybridization and hybridization were performed in 250 mM Na2HPO4, pH 7.2, 7% SDS, 1 mM EDTA for 3 and 12 h, respectively. Rabbit type II collagen transcript was probed with partial cDNA generated by reverse transcriptase PCR using RNA isolated from P0 cells. The forward PCR primer for type II collagen was 5'-GACCCCATGCAGTACATGCG-3' and the reverse primer was 5'-AGCCGCCATTGATGGTCTCC-3'. High specific activity random-primed probes were prepared from the PCR product (370 bp) using the T7 QuickPrime kit (Amersham Biosciences, Inc.) in the manner specified by the supplier. Filters were washed three times with 0.2× SSC/0.1% SDS, dried, and exposed to Kodak X-Omat film with intensifying screens at -80 °C.

NO Assay-- NO production was measured by estimating the stable NO metabolite, nitrite, in conditioned medium using a spectrophotometric method based on the Griess reaction (40). Following culture of chondrocytes for the specified times, 100 µl of the culture supernatants was mixed with 100 µl of Griess reagent (1% sulfanilamide, 0.1% naphthyl ethylenediamine dihydrochloride, and 1.25% H3PO4) and incubated for 10 min at room temperature. Nitrite concentrations were determined by measuring the absorbance at 550 nm in an enzyme-linked immunosorbent assay reader (Molecular Devices, Sunnyvale, CA).

Determination of Apoptosis-- Apoptotic death of chondrocytes was determined by examining DNA fragmentation using standard procedures. Briefly, cell pellets were resuspended in 750 µl of ice-cold lysis buffer (20 mM Tris-HCl, 10 mM EDTA, and 0.5% Triton X-100, pH 8.0) for 45 min with occasional shaking. DNA was extracted with phenol and precipitated with alcohol. The pellet was dried and resuspended in 100 µl of 20 mM Tris-HCl, pH 8.0. After digesting RNA with RNase (0.1 mg/ml) at 37 °C for 1 h, the samples (5 µg) were electrophoresed through a 1% agarose gel in 450 nM Tris-acetate-EDTA buffer, pH 8.0. DNA was photographed under UV light. Alternatively, apoptotic cell death was determined by terminal deoxynucleotidyl transfer-mediated nick end labeling according to the manufacturer's protocol (Roche Molecular Biochemicals). Apototic cell death was quantified by flow cytometry based on the number of cells with fragmented DNA. Briefly, the cells were harvested by centrifugation, washed in ice-cold phosphate-buffered saline, and fixed in 80% ethanol that had been precooled to -20 °C. The fixed cells were resuspended in phosphate-buffered saline containing 50 µg/ml propidium iodide, 0.1% Nonidet P-40, and 100 µg/ml RNase A (Sigma) and incubated in the dark for 1 h. The number of cells with fragmented DNA was then quantified using 1.2 × 104 cells on a FACSort flow cytometer using the Cellquest analysis program (Becton Dickinson, Mountain View, CA).

Caspase-3 Assay-- Activation of caspase-3 was determined by Western blot analysis using an antibody specific to the active form of caspase-3, purchased from New England Biolabs (Beverly, MA). Caspase-3 activity was determined by measuring the absorbance at 405 nm after cleavage of synthetic substrate Ac-Asp-Glu-Val-Asp-chromophore p-nitroaniline. Briefly, chondrocytes were collected by scraping in ice-cold phosphate-buffered saline and brief centrifugation and lysed on ice for 10 min in the cell lysis buffer provided in the CLONTECH A ApoAlertTM CPP32 colorimetric assay kit. The lysates were reacted with 50 µM Ac-Asp-Glu-Val-Asp-chromophore p-nitroaniline in a reaction buffer (0.1 M Hepes, 20% glycerol, 10 mM dithiothreitol, and protease inhibitors, pH 7.4). The mixtures were maintained at 37 °C for 1 h in a water bath and subsequently analyzed in an enzyme-linked immunosorbent assay reader (Molecular Devices, Sunnyvale, CA). The enzyme activity was calculated from a standard curve prepared using p-nitroanaline. The relative levels of pNA were normalized against the protein concentration of each extract.

Western Blot Analysis-- Whole cell lysates were prepared by extracting proteins using a buffer containing 50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 1% Nonidet P-40, and 0.1% SDS, supplemented with protease inhibitors (10 µg/ml leupeptin, 10 µg/ml pepstatin A, 10 µg/ml aprotinin, and 1 mM of 4-(2-aminoethyl) benzenesulfonyl fluoride) and phosphatase inhibitors (1 mM NaF and 1 mM Na3VO4). The proteins were size-fractionated by SDS-polyacrylamide gel electrophoresis and transferred to a nitrocellulose membrane. The nitrocellulose sheet was then blocked with 3% nonfat dry milk in Tris-buffered saline. Protein expression was determined using antibodies purchased from the following sources: mouse anti-p53 monoclonal antibody and rabbit active caspase-3 polyclonal antibody from New England Biolabs, ERK-2 from Transduction Laboratories (Lexington, KY), and p38 kinase from Santa Cruz Biotechnology (Santa Cruz, CA). The blots were developed using a peroxidase-conjugated secondary antibody and an ECL system.

MAP Kinase Assay-- Activation of ERK-1 and -2 was examined by Western blot analysis using antibodies specific to activated, tyrosine- and threonine-phosphorylated ERK-1/2 (New England Biolabs), as described previously (38, 39). Activity of p38 kinase was determined by immunocomplex kinase assay, as described previously (38, 39). Briefly, cell lysates were prepared in a lysis buffer containing 20 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1% Triton X-100, 2.5 mM sodium pyrophosphate, 1 mM beta -glycerolphosphate, and inhibitors of proteases and phosphatases. The samples were precipitated with rabbit polyclonal anti-p38 kinase antibody (Santa Cruz Biotechnology Inc.), and immune complexes were collected by binding to protein A-Sepharose beads (Pierce). After washing with lysis buffer, the beads were resuspended in 20 µl of kinase reaction buffer containing 25 mM Tris-HCl, pH 7.5, 5 mM beta -glycerolphosphate, 2 mM dithiothreitol, 0.1 mM sodium orthovanadate, 10 mM MgCl2, [gamma -32P]ATP, and 1 µg of ATF-2 fusion protein as a substrate for p38 kinase (New England Biolabs). Following incubation for 30 min at 30 °C, the reaction was stopped by the addition of 4× Laemmli's sample buffer followed by boiling. The samples were resolved by electrophoresis, and phosphorylation of ATF-2 was then determined by autoradiography.

Data Analyses and Statistics-- The results are expressed as the means ± S.E. values calculated from the specified number of determinations. A Student's t test was used to compare individual treatments with their respective control values. A probability of p < 0.05 was taken as denoting a significant difference.

    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Nitric Oxide Causes Dedifferentiation and Apoptosis of Chondrocytes-- Rabbit articular chondrocytes in primary culture were treated with the NO generator SNP, and NO production was measured by estimating the stable metabolite nitrite. As expected, SNP-treated chondrocytes produced NO in a dose- and time-dependent manner (Fig. 1). The effects of NO production on the differentiation status of chondrocytes were determined by examining expression of type II collagen and sulfated proteoglycan synthesis. Western and Northern blot analyses of chondrocytes treated with SNP for 24 h demonstrated that expression of type II collagen was significantly reduced at 1 mM SNP and undetectable at 3 mM SNP. Fig. 2A depicts the evident inhibition of type II collagen by SNP (1 mM), following 24 h of incubation. Consistent with the expression pattern of type II collagen, SNP treatment led to a dose-dependent decrease in the accumulation of sulfated proteoglycan (Fig. 2B). These results confirm that generation of NO causes dedifferentiation of primary cultured rabbit articular chondrocytes.


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  		Fig. 1.   Generation of NO in SNP-treated chondrocytes. Rabbit articular chondrocytes were treated with the indicated concentrations of SNP for 24 h (A) or with 1 mM SNP for the specified time periods (B). NO production was measured by estimating the stable NO metabolite, nitrite, in conditioned medium, using a spectrophotometric method based on the Griess reaction. The data represent the average values with standard deviation (n = 3).


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  		Fig. 2.   NO causes dedifferentiation of articular chondrocytes. Chondrocytes were treated with the indicated concentrations of SNP for 24 h (upper panel of A) or with 1 mM SNP for specified time periods (lower panel of A). The expression of type II collagen was determined by Northern (NB) and Western (WB) blot analysis (A). Accumulation of sulfated glycosaminoglycan in cells treated with the indicated concentrations of SNP for 24 h was quantified by Alcian Blue staining (B). The data in A represent the results of a typical experiment conducted four times, and B signifies the average values with standard deviation (n = 4).

Treatment of chondrocytes with SNP (1 mM) for 24 h additionally caused apoptotic cell death, as determined by DNA fragmentation (Fig. 3A), fluorescence-activated cell sorter sorting of propidium iodide-stained cells with fragmented DNA (Fig. 3B), and terminal deoxynucleotidyl transfer-mediated nick end labeling assay (Fig. 3C). Quantitation of SNP-treated cells with FACSort flow cytometer indicated dose- and time-dependent apoptosis (Fig. 3, D and E). Specifically, after SNP treatment for 24 h at concentrations of 1 and 3 mM, 19 and 43% cells underwent apoptosis (Fig. 3D), and cell death was observed as early as 6 h (Fig. 3E). Similar results were observed when apoptotic cells were quantified by terminal deoxynucleotidyl transfer-mediated nick end labeling assay (data not shown).


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  		Fig. 3.   NO causes apoptosis of chondrocytes. A, chondrocytes were untreated (lane C) or treated with 1 mM SNP for 24 h (lane S), and apoptotic cell death was determined by examining DNA fragmentation by electrophoresis. Lane M, molecular size markers. B, apoptotic cell death from control or SNP-treated (1 mM, 24 h) cells was determined by counting cells with fragmented DNA with fluorescence-activated cell sorter analysis. C, apoptosis of chondrocytes was determined by terminal deoxynucleotidyl transfer-mediated nick end labeling assay. Nuclei of SNP-treated cells were stained with propidium iodide (PI). D and E, chondrocytes were treated with the indicated concentrations of SNP for 24 h (D) or with 1 mM SNP for the specified time periods (E). Apoptotic cell death was quantified with the propidium iodide method, using a FACSorter flow cytometer. The data in A-C represent the results of a typical experiment conducted three times, and D and E represent the average values with standard deviation (n = 4).

Opposite Roles of ERK-1/2 and p38 Kinase in NO-induced Dedifferentiation of Chondrocytes-- To investigate the signaling pathway stimulating dedifferentiation and apoptosis of SNP-treated chondrocytes, we initially examined changes in the activity of ERK-1/2 and p38 kinase. These two MAP kinase subtypes were selected for study, based on results from previous analyses (38) indicating that the proteins conversely regulate chondrocyte differentiation. When cells were treated with 1 mM SNP, ERK-1/2 activity was transiently increased, as determined by phosphorylation status of the protein (Fig. 4A). Levels of ERK-1/2 phosphorylation began to increase at 3 h, reached peak levels at 12 h, and decreased thereafter. Similar to the pattern of ERK activation, p38 kinase activity was also transiently increased in SNP-treated cells (Fig. 4A). The effects of SNP on ERK-1/2 and p38 kinase activity were dose-dependent, as shown in Fig. 4B. NO-induced ERK and p38 kinase activation were blocked by PD98059, an inhibitor of ERK-1/2 upstream kinase (41), and SB203580, an inhibitor of p38 kinase (42), respectively (Fig. 4C).


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  		Fig. 4.   Activation ERK-1/2 and p38 kinase in SNP-treated chondrocytes. A and B, rabbit articular chondrocytes were treated with 1 mM SNP for the indicated time periods (A) or for 24 h with the specified concentrations of SNP (B). Activation of ERK-1/2 was determined by Western blot analysis, using antibody specific to activated ERK-1/2. p38 activity was determined by immunocomplex kinase assay, using ATF-2 as a substrate. Expression of ERK-1 and p38 MAP kinase was determined by Western blot analysis. C, chondrocytes were treated with 1 mM SNP for 24 h in the absence or presence of 30 µM PD98059 or SB203580. ERK-1/2 and p38 kinase activity was determined as described above. The data represent the results of a typical experiment conducted four times with comparable results.

To determine the association between ERK-1/2 activation and dedifferentiation, chondrocytes were treated with SNP in the presence and absence of PD98059, and expression of chondrocyte markers was examined. The addition of PD98059 to SNP-treated chondrocytes blocked the increased phosphorylation of ERK-1/2 (Fig. 5A, lower panel) and slowed the inhibition of both type II collagen expression (Fig. 5A, upper panel) and proteoglycan synthesis (Fig. 5C). The addition of SB203580 inhibited NO-induced p38 kinase activity, as determined by an immunocomplex kinase assay (Fig. 5B, lower panel). In contrast to the effects of ERK inhibition, blockade of p38 kinase accelerated NO-induced decrease of type II collagen expression (Fig. 5B, upper panel) and proteoglycan synthesis (Fig. 5C). The effects of PD98059 and SB203580 on proteoglycan synthesis at each concentration of SNP were statistically significant, with a probability of p < 0.05. These data suggest that NO-induced ERK-1/2 activation is responsible for the dedifferentiation of chondrocytes, whereas p38 kinase functions to maintain chondrocyte phenotype.


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  		Fig. 5.   ERK-1/2 and p38 kinase oppositely regulate NO-induced dedifferentiation of chondrocytes. Articular chondrocytes were treated with the indicated concentrations of SNP for 24 h in the absence or presence of PD98059 (A and C) or SB203580 (B and C). Expression of type II collagen was determined by Western blot analysis (A and B), and accumulation of sulfated glycosaminoglycan was quantified by Alcian Blue staining. The data in A and B depict the results of a typical experiment conducted four times, and B represents the average values with standard deviation (n = 4).

Opposite Roles of ERK-1/2 and p38 Kinase in NO-induced Apoptosis of Chondrocytes-- The role of ERK-1/2 and p38 kinase in NO-induced apoptosis of chondrocytes was next investigated. Upon inhibition of NO-induced ERK-1/2 activation with PD98059, chondrocyte apoptosis was significantly enhanced in a dose-dependent manner (Fig. 6). In the presence of 30 µM PD98059, SNP-induced apoptosis (1 mM) was increased by 1.9-fold, compared with untreated control cells. In contrast, inhibition of p38 kinase with SB203580 significantly blocked NO-induced apoptosis in a dose-dependent manner (Fig. 6). In cells treated with 30 µM SB203580, the percentage of SNP-stimulated apoptotic cells (19%) was decreased to 4.3%.


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  		Fig. 6.   ERK-1/2 and p38 kinase oppositely regulate NO-induced apoptosis of chondrocytes. Articular chondrocytes were treated with 1 mM SNP for 24 h in the presence of indicated concentrations of PD98059 or SB203580. Apoptotic cell death was quantified with the propidium iodide method using a FACSorter flow cytometer. The data represent the average values with standard deviation (n = 3).

In an attempt to elucidate the mechanism of MAP kinase-regulated apoptosis, we examined caspase-3, an executioner of cell death, and p53, a signaling molecule upstream of caspase-3, both of which are activated in response to various apoptotic stimuli. As shown in Fig. 7A, SNP treatment induced active caspase-3 and accumulation of p53 in a dose- and time-dependent manner. Increased apoptosis caused by inhibition of ERK-1/2 was accompanied by increased p53 and active caspase-3 accumulation (Fig. 7B). In contrast, decreased apoptosis caused by p38 kinase inhibition was associated with decreased accumulation of p53 and active caspase-3 (Fig. 7C). Our data imply that opposite regulation of apoptosis by ERK-1/2 and p38 MAP kinase is associated with converse effects on the modulation of p53 and caspase-3 pathway.


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  		Fig. 7.   ERK-1/2 and p38 kinase oppositely regulate accumulation of p53 and activation of caspase-3. A, chondrocytes were treated with the indicated concentrations of SNP for 24 h (upper panel) or with 1 mM SNP for the specified time periods (lower panel). B and C, cells were treated with 1 mM SNP for 24 h in the presence of indicated concentrations of PD98059 (B) or SB203580 (C). p53 and active caspase-3 were detected by Western blot analyses. The data represent the average values with standard deviation (n = 4).

In addition to the generation of active caspase-3 (determined by Western blot analysis), SNP treatment resulted in a dramatic increase in caspase-3 activity (Fig. 8A). Inhibition of ERK-1/2 with PD98059 potentiated NO-induced caspase-3 activity, whereas p38 kinase inhibition blocked caspase-3 activation. Addition of the caspase-3 inhibitor, z-Asp-Glu-Val-Asp-fluoromethyl ketone, significantly blocked NO-induced caspase-3 activity (Fig. 8A) and apoptosis (Fig. 8B) in chondrocytes, suggesting a pivotal role for the caspase-3 in NO-induced apoptosis. We suggest that the activities of proapoptotic p38 kinase and anti-apoptotic ERK depend on the modulation of caspase-3.


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  		Fig. 8.   Inhibition of caspase-3 activity blocks NO-induced apoptosis. Chondrocytes were treated with 1 mM SNP for 24 h in the absence or presence of indicated concentrations of 30 µM PD98059 or SB203580. Caspase-3 activity was determined using a colorimetric substrate (A), and apoptosis was quantified by FACSorter (B). The data represent the average values with standard deviation (n = 3).

NO-induced Chondrocyte Apoptosis Is Dependent on Differentiation Status-- The above results imply that ERK-1/2 and p38 kinase oppositely regulate both dedifferentiation and apoptosis, two processes simultaneously initiated by NO generation. These data also suggest that NO-induced dedifferentiation and apoptosis are interrelated based on the observations that inhibition of ERK slows dedifferentiation with the enhanced apoptosis, whereas inhibition of p38 kinase accelerates dedifferentiation with the blockade of apoptosis. We therefore further investigated the relationship between NO-induced dedifferentiation and apoptosis in chondrocytes. For this purpose, chondrocytes were passaged to P4 by a serial monolayer culture or treated with 100 nM phorbol 12-myristate 13-acetate (PMA) for 2 days to cause dedifferentiation (43, 44). Under these conditions, expression of type II collagen and synthesis of proteoglycan was almost completely inhibited (Fig. 9A), confirming the loss of cell phenotype. SNP treatment (1 mM for 24 h) of cells dedifferentiated by either PMA treatment (Fig. 9B) or serial monolayer culture (Fig. 9C) resulted in 7.8 and 4.6% apoptotic cells, compared with 19% in differentiated cells. Our findings suggest that NO-induced chondrocyte apoptosis depends on the differentiated phenotype of these cells.


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  		Fig. 9.   NO-induced apoptosis depends on the differentiation status of chondrocytes. A, dedifferentiation of chondrocytes. P0 chondrocytes were treated with 100 nM PMA for 48 h (PMA) or subcultured to passage 4 (P4). Accumulation of sulfated glycosaminoglycan was quantified by Alcian Blue staining (upper panel) and expression of type II collagen was determined by Western blot analysis (lower panel). Con, control. B, passage 0 chondrocytes were left untreated or treated with indicated concentrations of PMA for 24 h. The cells were further incubated for 24 h in the presence or absence of 1 mM SNP in PMA containing medium. C, P0 or P4 cells were treated with the indicated concentrations of SNP for 24 h, and apoptotic cell death was quantified using FACScan. The data represent the average values with standard deviation (n = 3).

We subsequently examined the molecular mechanism for the requirement of differentiated chondrocyte phenotype in NO-induced apoptosis. As shown in Fig. 10A, SNP treatment induced a similar degree of NO production in both differentiated and dedifferentiated chondrocytes. However, consistent with observations of decreased apoptosis, NO-induced accumulation of p53 was significantly reduced in dedifferentiated cells (Fig. 10B). Formation (Fig. 10B) and activity (Fig. 10C) of caspase-3 were also significantly reduced in dedifferentiated cells. In PMA-induced dedifferentiated chondrocytes at P0, the addition of SNP caused ERK-1/2 activation, similar to differentiated chondrocytes. Dedifferentiation caused by serial monolayer culture significantly elevated ERK-1/2 activity, and the addition of SNP slightly increased the phosphorylation level. In contrast to ERK-1/2, SNP-induced activation of p38 kinase was considerably reduced, both in monolayer culture- and PMA-induced dedifferentiated cells (Fig. 10B). The results collectively indicate that reduced apoptosis in dedifferentiated cells on SNP treatment is due to inability to activate p38 kinase, p53 accumulation, and caspase-3 activation.


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  		Fig. 10.   NO does not induce activation of p38 kinase and caspase-3 or accumulation of p53 in dedifferentiated chondrocytes. A, P0 chondrocytes, dedifferentiated chondrocytes by PMA treatment 24 h (P0+PMA), or P4 cells were treated with the indicated concentrations of SNP for 24 h, and generation of NO was determined by estimating the release of the stable NO metabolite, nitrite. B and C, P0 chondrocytes, chondrocytes dedifferentiated by PMA treatment for 24 h (P0+PMA), or P4 cells were treated with the indicated concentrations of SNP for 24 h, and active caspase-3, p53, phosphorylated ERK-1/2, and p38 kinase activity was determined as described for Fig. 4 (B). Caspase-3 activity was determined, using a colorimetric substrate (C). The data in A and C represent the average values with standard deviation (n = 3), and B represents the results of a typical experiment conducted three times.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Nitric oxide, produced by inflammatory cytokines in chondrocytes, is implicated as a critical mediator of arthritis. Compared with the normal state, arthritic cartilage produces a large amount of NO in both spontaneous and proinflammatory cytokine-stimulated conditions (16). A high level of nitrite/nitrate is found in the synovial fluid and serum of arthritis patients (45, 46), because of increased levels of iNOS (46, 47). NO-induced cartilage destruction is caused by active degradation of ECM via modulation of matrix metalloproteinases (15), as well as induction of dedifferentiation (10, 11) and apoptosis (12-14). The molecular basis of the signaling pathway of NO-induced modulation of phenotype and apoptosis is yet to be properly characterized. Our study provides evidence for the opposite regulation of apoptosis and dedifferentiation of articular chondrocytes by two MAP kinase subtypes, ERK-1/2 and p38 kinase. We further demonstrate that the differential effects of the two proteins on apoptosis are closely associated with converse regulation of p53 accumulation and caspase-3 activation in a differentiation status-dependent pathway, as depicted in Fig. 11.


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  		Fig. 11.   Schematic summary of NO-induced dedifferentiation and apoptosis of articular chondrocytes. NO-induced p38 kinase functions to maintain differentiated phenotypes of chondrocytes and/or to inhibit dedifferentiation and causes apoptosis, whereas ERK activity causes dedifferentiation and/or inhibits the maintenance of differentiated phenotype and functions as anti-apoptotic signal. The opposite effects of ERK and p38 kinase in apoptosis are brought about by converse regulation of p53 and caspase-3 in a differentiation status-dependent pathway.

Signaling Pathways in NO-induced Chondrocyte Apoptosis-- To elucidate the signaling pathway leading to apoptosis in SNP-treated chondrocytes, we focused on ERK-1/2 and p38 kinase, based on the observation by other investigators that these two MAP kinase subtypes are activated by NO generation (33) and that chondrocyte function (including differentiation) is oppositely regulated by these two protein subtypes, as revealed in a previous study by our group (38). Here, we show that ERK-1/2 and p38 kinases activated by NO in a similar kinetic pattern regulate apoptosis conversely. By using specific inhibitors of the MAP kinase subtypes, we established that p38 kinase activation is a primary signal for apoptosis, whereas ERK-1/2 functions as an anti-apoptotic signal (Fig. 11). Similar observations regarding the role of MAP kinase in survival and apoptosis were noted in chondrocytes as well as other cell types. For example, a recent study by Shakibaei et al. (32) showed that ERK functions as a survival signal. Their group demonstrated that survival of human chondrocytes on type II collagen or integrin beta 1 requires ERK signaling, and inhibition of ERK causes cellular apoptosis. Additionally, the proapoptotic role of p38 kinase has been established in chondrocytes (17, 21).

The converse effects of ERK and p38 kinase on chondrocyte apoptosis appear to be associated with the accumulation of p53 tumor suppressor protein. A study by Yatsugi et al. (48) suggested the possible involvement of p53 in chondrocyte apoptosis and cartilage destruction. Immunohistochemical analyses of p53 in the articular cartilage of patients with rheumatoid arthritis and osteoarthritis showed increased staining of the tumor suppressor protein in chondrocytes with apoptotic morphology. Accumulation of p53 plays a major role in a variety of cellular responses, including apoptosis. It is believed that phosphorylation of p53 is the mechanism leading to accumulation, which results in prolonged protein half-life because of inhibition of ubiquitination and degradation (49, 50). Within the MAP kinase family, p53 is phosphorylated either directly or indirectly by c-Jun N-terminal kinase (27, 51), p38 kinase (28-30, 52, 53), and ERK (30). Despite a contradictory report indicating that p53 phosphorylation by p38 kinase induces transcriptional activity without affecting accumulation (31), it is generally recognized that phosphorylation is the primary mechanism that causes accumulation. Although we did not determine the detailed mechanism of p38 kinase-mediated accumulation of p53 in this study, we hypothesize that NO-induced p38 kinase activity in chondrocytes phosphorylates p53 either directly or indirectly, resulting in accumulation.

Compared with the numerous studies on the role of p38 kinase in p53 regulation, there is less evidence to indicate phosphorylation and/or accumulation of p53 by ERK-1/2. Earlier analyses show that, depending on the type of cells and extracellular stimuli, ERK activity leads to accumulation of p53 (54) or protects against apoptosis, possibly by modulating p53 accumulation (55). However, our experiments clearly indicate that NO-induced ERK-1/2 in articular chondrocytes functions as an anti-apoptotic signal, in association with its ability to reduce p53 accumulation and caspase-3 activation. Although further experiments are required to address whether the functions of ERK-1/2 and p38 kinase in p53 accumulation result from the phosphorylation of the tumor suppressor protein, to our knowledge, this is the first report to indicate the association of p53 with NO-induced chondrocyte apoptosis and opposite regulation by ERK-1/2 and p38 kinase.

To date, the mechanism of p53 regulation of apoptosis has not been clearly elucidated. One of the proposed mechanisms for p53 activity is via the activation of caspase. The p53 protein transcriptionally regulates Bax, a proapoptotic member of Bcl-2 family (56). In response to apoptotic signals, Bax is redistributed from the cytosol to the mitochondria, where it causes a decline in mitochondrial membrane potential, followed by cytochrome c release and caspase activation (18-24, 26). The findings that: 1) NO-induced apoptosis is mediated by caspase-3 activation and associated with p53 accumulation in a p38 kinase-dependent pathway and 2) potentiation of caspase-3 activation by the inhibition of ERK-1/2 or inhibition of caspase-3 by the blockade of p38 kinase is closely analogous with the level of p53 accumulation support our hypothesis that NO-induced ERK and p38 kinase regulate apoptosis via a p53- and caspase-3-dependent pathway (Fig. 11).

Recently, Notoya et al. (17) reported that NO production in human osteoarthritic chondrocytes induces apoptosis via caspase-3 activation. Consistent with our theory, these authors suggest that NO-induced p38 kinase activity leads to caspase-3 activation, which in turn causes apoptosis, as confirmed by the observation that inhibition of p38 kinase by SB203580 and caspase-3 by z-Asp-Glu-Val-Asp-fluoromethyl ketone blocked NO-induced apoptosis. However, in contrast to our results, their data demonstrated that inhibition of ERK with PD98059 protected NO-induced apoptosis, indicating that NO-induced ERK also functions as a proapoptotic signal. This discrepancy may arise from the different states of cells utilized in the two sets of experiments. Whereas we employed normal articular chondrocytes, Notoya et al. used degenerative osteoarthritic chondrocytes, which have different characteristics. Osteoarthritic chondrocytes produce NO, both via proinflammatory cytokines and spontaneously, whereas normal chondrocytes do not produce NO spontaneously (9, 57). Therefore, further studies on the kinetics of MAP kinase activation, p53 accumulation, and caspase activation are necessary to distinguish between mechanisms of NO-induced apoptosis in normal and osteoarthritic chondrocytes.

Regulation of NO-induced Dedifferentiation and Its Relationship with Apoptosis-- Cartilage destruction by NO production in degenerative disease is due to active degradation of ECM by matrix metalloproteinase, as well as insufficient synthesis of ECM molecules caused by dedifferentiation and apoptosis of chondrocytes. In addition to NO effects, the differentiated phenotype of chondrocytes is rapidly lost through other environmental changes such as exposure to phorbol ester (44) or during a serial monolayer culture (43). Signaling pathways during chondrocyte dedifferentiation are not clearly understood. In this study, we demonstrate that NO-induced dedifferentiation of chondrocytes is conversely mediated by ERK-1/2 and p38 kinase; although NO-induced ERK-1/2 activation is responsible for the induction of dedifferentiation and/or inhibition of the maintenance of differentiated phenotype, p38 kinase functions to maintain chondrocyte phenotype and/or inhibition of dedifferentiation (Fig. 11). The opposing roles of ERK and p38 kinase are also observed in interleukin-1beta -induced dedifferentiation of chondrocytes (data not shown). Signaling pathways during dedifferentiation appear to differ, depending on the cause for cellular phenotype loss. We observed that chondrocyte dedifferentiation induced by a serial monolayer culture was accompanied by sustained activation of ERK-1/2 (Fig. 10B), and inhibition of ERK-1/2 by PD98059 blocked dedifferentiation.2 In contrast, no change in ERK activity was noted during dedifferentiation induced by prolonged treatment with phorbol ester (Fig. 10C) or retinoic acid (data not shown).

Because NO-induced ERK-1/2 and p38 kinase oppositely regulate both dedifferentiation and apoptosis, we investigated the relationship between these processes in chondrocytes. The ability of NO to activate p38 kinase was significantly reduced in dedifferentiated cells in our experiments. This might explain the reduced accumulation of p53 and caspase-3 activity in dedifferentiated cells. The molecular mechanisms of reduced activation of p38 kinase, p53 accumulation, and caspase-3 activation in dedifferentiated cells remain to be determined. However, it is known that cellular responses to extracellular stimuli are different in differentiated and dedifferentiated chondrocytes. For example, interleukin-1 in primary chondrocytes has growth inhibitory effects, dependent on the differentiation status of cells. In cells that are dedifferentiated by serial monolayer culture, this inhibitory effect is lost because of reduced expression of iNOS (58).

In summary, we demonstrate that NO-induced ERK and p38 kinase have opposite effects on dedifferentiation and apoptosis, in that although p38 kinase functions as an apoptosis signal and maintains chondrocyte phenotype, ERK activity causes dedifferentiation and operates as an anti-apoptotic signal. These effects are closely associated with the accumulation of p53 and caspase-3 activity. In addition, the proapoptotic role of p38 kinase is dependent on differentiation status, as confirmed by the finding that NO does not effectively activate p38 kinase, p53 accumulation, caspase-3 activation, or apoptosis in differentiated chondrocytes.

    FOOTNOTES

* This work was supported by grants from the Korea Ministry of Science and Technology (Life Phenomena and Function Research Group).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§ Supported in part by the Brain Korea 21 program.

Supported by Korean Science and Engineering Foundation (Interdisciplinary Research Project 1999-2-207-004-5).

** To whom correspondence should be addressed: Dept. of Life Science, Kwangju Institute of Science and Technology, Pook-Gu, Kwangju, 500-712, Korea. Tel.: 82-62-970-2497; Fax: 82-62-970-2484; E-mail: jschun@kjist.ac.kr.

Published, JBC Papers in Press, October 31, 2001, DOI 10.1074/jbc.M107231200

2 Y.-M. Yoon, S.-J. Kim, C.-D. Oh, J.-W. Ju, W. K. Song, Y. J. Yoo, T.-L. Huh, and J.-S. Chun, submitted for publication.

    ABBREVIATIONS

The abbreviations used are: ECM, extracellular matrix, ERK, extracellular signal-regulated protein kinase; MAP, mitogen-activated protein; iNOS, inducible nitric-oxide synthase; Pn, passage n; PMA, phorbol 12-myristate 13-acetate; SNP, sodium nitroprusside.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

1. Sandell, L. J., and Adler, P. (1999) Frontiers Biosci. 4, 731-742
2. Sandell, L. J., and Aigner, T. (2000) Arthritis Res. 3, 107-113
3. Poole, A. R. (1999) Frontiers Biosci. 4, 662-670
4. Frenkel, S. R., and Di Cesare, P. E. (1999) Frontiers Biosci. 4, 671-685
5. Smith, R. L. (1999) Frontiers Biosci. 4, 704-712
6. Denko, C. W., and Malemud, C. J. (1999) Frontiers Biosci. 4, 686-693
7. Martel-Pelletier, J., Alaaeddine, N., and Pelletier, J. P. (1999) Frontiers Biosci. 4, 694-703
8. Stadler, J., Stefanovic-Racic, M., Billiar, T. R., Curran, R. D., McIntyre, L. A., Georgescu, H. I., Simmons, R. L., and Evans, C. H. (1991) J. Immunol. 147, 3915-3920
9. Amin, A. R., and Abramson, S. B. (1998) Curr. Opin. Rheumatol. 10, 263-268
10. Cao, M., Westerhausen-Larson, A., Niyibizi, C., Kavalkovich, K., Georgescu, H. I., Rizzo, C. F., Hebda, P. A., Stefanovic-Racic, M., and Evans, C. H. (1997) Biochem. J. 324, 305-310
11. Taskiran, D., Stefanovic-Racic, M., Georgescu, H. I., and Evans, C. H. (1994) Biochem. Biophys. Res. Commun. 200, 142-148
12. Hashimoto, S., Ochs, R. L., Komiya, S., and Lotz, M. (1998) Arthritis Rheum. 41, 1632-1638
13. Blanco, F. J., Guitian, R., Vazquez-Martul, E., de Toro, F. J., and Galdo, F. (1998) Arthritis Rheum. 41, 284-289
14. Blanco, F. J., Schwarz, R. L., Ochs, H., and Lotz, M. (1995a) Am. J. Pathol. 146, 75-85
15. Murrell, G. A. C., Jang, D., and Williams, R. J. (1995) Biochem. Biophys. Res. Commun. 206, 15-21
16. Pelletier, J. P., Jovanovic, D., Fernandes, J. C., Manning, P. T., Connor, J. R., Currie, M. G., Di, Battista, J. A., and Martel-Pelletier, J. (1998) Arthritis Rheum. 41, 1275-1286
17. Notoya, K., Jovanovic, D. V., Reboul, P., Martel-Pelletier, J., Mineau, F., and Pelletier, J. P. (2000) J. Immunol. 165, 3402-3410
18. Kluck, R. M., Bossy-Wetzel, E., Green, D. R., and Newmeyer, D. D. (1997) Science 275, 1132-1136
19. Yang, J., Liu, X. S., Bhalla, K., Kim, C. N., Ibrado, A. M., Cai, J. Y., Peng, T. I., Jones, D. P., and Wang, X. (1997) Science. 275, 1129-1132
20. Shimizu, S., Narita, M., and Tsujimoto, Y. (1999) Nature 399, 483-487
21. Ghatan, S., Larner, S., Kinoshita, Y., Hetman, M., Patel, L., Xia, Z., Youle, R. J., and Morrison, R. S. (2000) J. Cell Biol. 150, 335-347
22. Hsu, Y. T., Wolter, K. G., and Youle, R. J. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 3668-3672
23. Marzo, I., Brenner, C., Zamzami, N., Jürgensmeier, J. M., Susin, S. A., Vieira, H. L. A., Prévost, M. C., Xie, X., Matsuyama, S., Reed, J. C., and Kroemer, G. (1998) Science. 281, 2027-2031
24. Schuler, M., Bossy-Wetzel, E., Goldstein, J. C., Fitzgerald, P., and Green, D. R. (2000) J. Biol. Chem. 275, 7337-7342
25. Marchenko, N. D., Zaika, A., and Moll, U. M. (2000) J. Biol. Chem. 275, 16202-16212
26. Xiang, H., Kinoshita, Y., Knudson, C. M., Korsmeyer, S. J., Schwartzkroin, P. A., and Morrison, R. S. (1998) J. Neurosci. 18, 1363-1373
27. Fuchs, S. Y., Adler, V., Pincus, M. R., and Ronai, Z. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 10541-10546
28. Huang, C., Ma, W. Y., Maxiner, A., Sun, Y., and Dong, Z. (1999) J. Biol. Chem. 274, 12229-12235
29. Bulavin, D. V., Saito, S., Hollander, M. C., Sakaguchi, K., Anderson, C. W., Appella, E., and Fornace, A. J., Jr. (1999) EMBO J. 18, 6845-6854
30. She, Q. B., Chen, N., and Dong, Z. (2000) J. Biol. Chem. 275, 20444-20449
31. Sanchez-Prieto, R., Rojas, J. M., Taya, Y., and Gutkind, J. S. (2000) Cancer Res. 60, 2464-2472
32. Shakibaei, M., Schulze-Tanzil, G., de Souza, P., John, T., Rahmanzadeh, M., Rahmanzadeh, R., and Merker, H. J. (2001) J. Biol. Chem. 276, 13289-13294
33. Callsen, D., and Brune, B. (1999) Biochemistry 38, 2279-2286
34. Xia, Z., Dickens, M., Raingeaud, J., Davis, R. J., and Greenberg, M. E. (1995) Science 270, 1326-1331
35. Robinson, M. J., and Cobb, M. H. Curr. Opin. Cell Biol. 9, 180-186
36. Schaeffer, H. J., and Weber, M. J. (1999) Mol. Cell. Biol. 19, 2435-2444
37. Chang, L., and Karin, M. (2001) Nature 410, 37-40
38. Oh, C.-D., Chang, S.-H., Yoon, Y.-M., Lee, S.-J., Lee, Y.-S., Kang, S.-S., and Chun, J.-S. (1999) J. Biol. Chem. 275, 5613-5619
39. Yoon, Y.-M., Oh, C.-D., Kim, D.-Y., Lee, Y.-S., Park, J.-W., Huh, T.-L., Kang, S.-S., and Chun, J.-S. (2000) J. Biol. Chem. 275, 12353-12359
40. Green, L. C., Wagner, D. A., Glogowski, J., Skipper, P. L., Wishnok, J. S., and Tannenbaum, S. R. (1982) Anal. Biochem. 126, 131-138
41. Alessi, D. R., Cuenda, A., Cohen, P., Dudley, D. T., and Saltiel, A. R. (1995) J. Biol. Chem. 270, 27489-27494
42. Cuenda, A., Rouse, J., Doza, Y. N., Meier, R., Cohen, P., Gallagher, T. F., Young, P. R., and Fee, J. C. (1995) FEBS Lett. 364, 229-233
43. Benya, P. D., Padilla, S. R., and Nimni, M. E. (1978) Cell 15, 1313-1321
44. Finer, M. H., Gerstenfeld, L. C., Young, D., Doty, P., and Boedtker, H. (1985) Mol. Cell. Biol. 6, 1415-1424
45. Farrell, A. J., Blake, D. R., Palmer, R. M., and Moncada, S. (1992) Ann. Rheum. Dis. 51, 1219-1222
46. McInnes, I. B., Leung, B. P., Field, M., Wei, X. Q., Huang, F. P., Sturrock, R. D., Kinninmonth, A., Weidner, J., Mumford, R., and Liew, F. Y. (1996) J. Exp. Med. 184, 1519-1524
47. Grabowski, P. S., Wright, P. K., Van't Hof, R. J., Helfrich, M. H., Ohshima, H., and Ralston, S. H. (1997) Br. J. Rheumatol. 36, 651-655
48. Yatsugi, N., Tsukazaki, T., Osaki, M., Koji, T., Yamashita, S., and Shindo, H. (2000) J. Orthop. Sci. 5, 150-156
49. Kubbutat, M. H., Jones, S. N., and Vousden, K. H. (1997) Nature 387, 299-303
50. Haupt, Y., Maya, R., Kazaz, A., and Oren, M. (1997) Nature 387, 296-299
51. Hu, M. C., Qiu, W. R., and Wang, Y. P. (1997) Oncogene. 15, 2277-2287
52. Sayed, M., Kim, S. O., Salh, B. S., Issinger, O. G., and Pelech, S. L. (2000) J. Biol. Chem. 275, 16569-16573
53. Keller, D., Zeng, X., Li, X., Kapoor, M., Iordanov, M. S., Taya, Y., Lozano, G., Magun, B., and Lu, H. (1999) Biochem. Biophys. Res. Commun. 261, 464-471
54. Persons, D. L., Yazlovitskaya, E. M., and Pelling, J. C. (2000) J. Biol. Chem. 275, 35778-35785
55. Anderson, C. N., and Tolkovsky, A. M. (1999) J. Neurosci. 19, 664-673
56. Budihardjo, I., Oliver, H., Lutter, M., Luo, X., and Wang, X. (1999) Annu. Rev. Cell Dev. Biol. 15, 269-290
57. Pelletier, J. P., Mineau, F., Ranger, P., Tardif, G., and Martel-Pelletier, J. P. (1996) Osteoarthritis Cart. 4, 77-84
58. Blanco, F. J., Geng, Y., and Lotz, M. (1995b) J. Immunol. 154, 4018-4026

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