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CARDIOVASCULAR
Graduate Interdisciplinary Program in Genetics and Genomics (S.P.) and Department of Pharmacology (Q.M.C.), College of Medicine, University of Arizona, Tucson, Arizona
Received September 5, 2004; accepted November 30, 2004.
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Abstract |
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). This type of cell is traditionally thought to be terminally differentiated and not capable of dividing in adult hearts. In response to an increased workload or cell loss, CMCs often enlarge their size and become hypertrophic. This scenario is evident after myocardial infarction, chronic cardiac ischemia, or an increase in pressure load caused by hypertension. Hypertrophy can be initially beneficial to compensate for the new demand on cardiac output. However, sustained and extensive hypertrophy of CMCs eventually contributes to heart failure (Francis et al., 1995; Frey and Olson, 2003). After myocardial infarction, heart fibroblasts (HFs) are crucial for healing infarcted areas and creating scar tissue. Normally, these cells provide structural support for the contractile function of CMCs by maintaining the homeostasis of the extracellular matrix through production of proteases and matrix proteins. In numerous types of heart disease, fibroblasts have been shown to participate in myocardium remodeling. Fibrosis in the heart derives from overstimulation or malfunction of fibroblasts, resulting in increased stiffness of local areas or of the heart overall. The consequence of fibrosis is blockage of contractile function.
Oxidative stress has been linked to cardiac remodeling and heart failure (Chen et al., 2001b). Reactive oxygen species (ROS) are released during ischemia and ischemic reperfusion. In addition, catecholamines and angiotensin II (AngII), two endocrine factors known to induce cardiac hypertrophy and cardiac remodeling, have been shown to increase the production of oxidants (Miller et al., 1996; Griendling and Alexander, 1997). Three branches of MAPKs are activated by ROS: JNKs, ERK1/2, and p38. MAPK activation precedes, and is often necessary for, CMC hypertrophy or myocardial fibrosis (Molkentin and Dorn, 2001). An initiator of the MAPK pathway located on the plasma membrane would be a convenient target for designing drugs that inhibit ROS from inducing these disease phenotypes in the heart.
One unsolved question is whether different cell types share similar initiation pathways of MAPK activation by ROS. Some reports have linked oxidants with a direct activation of the nonreceptor tyrosine kinase Src in mouse fibroblasts and erythrocytes (Abe et al., 1997; Mallozzi et al., 1999), whereas others have shown that small G proteins like G
i and Go contribute to ROS-induced MAPK activation in CMCs (Nishida et al., 2000). Examining two different cell types within an organ is useful for addressing whether different cell types share similar initiators in MAPK activation by ROS. Traditionally, it is assumed that oxidants diffuse through the plasma membrane to initiate the signaling pathways. Recent evidence suggests a role of growth factor receptors on the plasma membrane as the initiator of MAPK signaling by ROS.
Peptide growth factors, such as epidermal growth factor (EGF), bind to their receptors in the plasma membrane to initiate a cascade of signaling events. The EGF receptor (EGFR) is a receptor tyrosine kinase that is ubiquitously expressed in a variety of cell types, with the most abundant expression in epithelial cells and many cancer cells (Carpenter, 2000; Mendelsohn and Baselga, 2000; Prenzel et al., 2001). EGFR belongs to a family containing three other members (ErbB2, ErbB3, and ErbB4) that undergo homodimerization or heterodimerization to induce autophosphorylation and receptor tyrosine kinase activation in response to ligand binding (Mendelsohn and Baselga, 2000; Schlessinger, 2002). The EGFR dimers are shuttled from the caveolae into clathrin-coated pits for internalization and ligand release before recycling back to the plasma membrane (Carpenter, 2000). Activation of the receptor tyrosine kinase triggers downstream Ras-Raf-MAPK signaling. Specific inhibitors of receptor tyrosine kinases have been developed to block the downstream signaling and abolish the biological consequence of growth factors (Mendelsohn and Baselga, 2000). Such inhibitors have been made for the EGFR, platelet-derived growth factor receptor (PDGFR), and fibroblast growth factor receptor (FGFR). The availability of these inhibitors allows us to test the involvement of these receptors in ROS-induced MAPK activation.
The EGFR has been shown to mediate ROS-induced MAPK activation in vascular endothelial cells and renal epithelial cells (Chen et al., 2001a; Zhougang and Schnellmann, 2004). In CMCs and vascular smooth muscle cells, AngII has been found to transactivate the EGFR upon binding to its own nontyrosine kinase receptor AngII type 1 (Frank and Eguchi, 2003; Shah and Catt, 2003). AngII induces the production of ROS in vascular smooth muscle cells (Griendling and Alexander, 1997; Ushio-Fukai et al., 1999). These findings lead us to test here whether EGFR mediates MAPK signaling pathways activated by ROS in CMCs and HFs.
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Materials and Methods |
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; Coronella-Wood et al., 2004) and seeded at a density of 0.5 x 106 cells per well of a six-well plate (Falcon; BD Biosciences Discovery Labware, Bedford, MA) in low-glucose Dulbecco's modified Eagle's medium (DMEM) containing 10% fetal bovine serum (FBS). Two days later, the cells were rinsed with phosphate-buffered saline, and the media were replaced with 0.5% FBS low-glucose DMEM for a 2-day starvation. At the time of H2O2 treatment, more than 90% of CMCs express myosin heavy chain (Tu et al., 2003; Coronella-Wood et al., 2004).
HFs were retained by differential plating during the preparation of CMCs. HFs were cultured in high-glucose DMEM containing 10% FBS and allowed to grow in 100-mm dishes. Because HFs grow voluntarily, unlike CMCs or endothelial cells, the cells were subcultured once to reduce the possible contamination by CMCs and endothelial cells. The second-passage HFs were seeded into six-well plates at a density of 0.3 x 106 cells per well. One day after seeding, the fibroblasts were rinsed with phosphate-buffered saline, and the media were changed to 0.5% FBS high-glucose DMEM for a 2-day starvation. Nearly all of the cells in the HF preparation express vimentin (a marker of fibroblasts) but not desmin (a marker for CMCs) or -von Willebrand factor (a marker for endothelial cells).
Drug Treatment and Sample Harvesting. HFs and CMCs were treated and harvested identically. Most inhibitors were purchased from Calbiochem (San Diego, CA) except for BB2516, which was a gift from Dr. G. Timothy Bowden (Arizona Cancer Center, University of Arizona). Serum-starved cells were pretreated with AG-1478 for 30 min or other inhibitors for 1 h before being treated with 100 µM H2O2 for the time indicated in the text. The cells were harvested in a lysis buffer [20 mM HEPES (pH 7.4), 2 mM EGTA, 50 mM 
-glycerophosphate, 1% Triton X-100, 10% glycerol, and 1 mM dithiothreitol with 1 mM Na3VO4, 2 µg/ml leupeptin, 10 µg/ml aprotinin, and 400 µM phenylmethanesulfonyl fluoride freshly added]. Samples were then stored on ice for 15 min before centrifugation at 12,000 rpm at 4°C for 10 min in a Beckman Microfuge R centrifuge (Beckman Coulter, Fullerton, CA). Supernatants were transferred to fresh tubes, and protein concentration was determined using the Bradford protein assay solution according to the manufacturer's instructions (Bio-Rad, Hercules, CA).
Western Blot Protocol. Cell lysate containing 20 to 30 µg of proteins was loaded in each well of a 10% SDS-polyacrylamide gel (Bio-Rad). The gel was run at 60 V for 10 min and then 90 V for 2.5 h to achieve the desired separation. Proteins were then transferred to polyvinylidene fluoride membranes by electrophoresis at 65 V for 3 h at 4°C. Membranes were blocked with 5% nonfat milk and rinsed briefly in Tris-buffered saline/Tween 20 before incubation in a primary antibody solution overnight at 4°C with rocking. All of the primary antibodies were diluted in Tris-buffered saline/Tween 20 solution containing 5% bovine serum albumin and 0.1% sodium azide (1:400 for p-JNK, catalog no. 9251; 1:1000 for p-ERK1/2, catalog no. 9101; and 1:500 for p-p38, catalog no. 9211; Cell Signaling Technology Inc., Beverly, MA). After the primary antibody incubation, membranes were washed five times in Tris-buffered saline/Tween 20 and then placed in an anti-rabbit secondary antibody conjugated with horseradish peroxidase (no. 65-6120, 1:2500 dilution for p-JNK and p-p38 and 1:5000 dilution for p-ERK1/2; Zymed Laboratories, South San Francisco, CA) for a 2-h incubation at room temperature. Membranes were then washed five times before exposure to enhanced chemiluminescence reagents (Sigma-Aldrich, St. Louis, MO). After detection of phosphorylated proteins, membranes were stripped from their bound antibodies using either 0.1 N NaOH or Restore Buffer (Pierce, Rockford, IL). The membranes were then blotted with antibodies to total JNK, ERK, or p38 (1:1000 dilution for all antibodies; catalog no. 9252 for JNKs, no. 9102 for ERK1/2, and no. 9212 for p38; Cell Signaling Technology Inc.) as described previously for measuring the total level of the proteins.
To probe for EGFR and vinculin, cell lysates containing 50 µg of proteins were run on a 6.5% SDS-polyacrylamide gel and transferred and treated as described previously. Primary antibodies were diluted 1:2000 for anti-EGFR (no. 06-129; Upstate Biotechnology, Lake Placid, NY) or 1:1000 for anti-vinculin (no. V9131; Sigma-Aldrich). Secondary antibodies were anti-sheep (1:12,000, no. 61-8620) for the EGFR blot and anti-mouse (1:5000, no. 81-6520) for the vinculin blot (both from Zymed Laboratories). The EGFR blot was developed using Femto enhanced chemiluminescence reagents (no. 34095; Pierce).
To quantify the band intensities, the images of Western blot films were scanned, and the bands were quantitated using software from Alpha Innotech (San Leandro, CA) with an output in Excel. The band intensities of the control groups were set to 1, and all of the treatment readings are therefore considered -fold increases over the control within the same experiment.
Immunoprecipitation. Cell lysates containing 300 to 500 µg of protein were used for immunoprecipitations. Samples were mixed with the antibody against insulin receptor substrate 1 (IRS-1; 1:100 dilution, polyclonal rabbit, catalog no. 06-248; Upstate Biotechnology) and rotated at 4°C for 2 h. Protein A-conjugated Sepharose beads (30 µl, 1:1 dilution) were added to the samples and rotated at 4°C for 1 h. Beads were collected by centrifugation at 3000 rpm for 1 min, and the supernatant was aspirated. Beads were then washed with 1 ml of PAN buffer [10 mM PIPES (pH 7), 100 mM NaCl, and 20 µg/ml aprotinin] with Nonidet P40 (0.5%) and then 1 ml of PAN buffer without Nonidet P40. The bound proteins were dissociated from the beads by 10-min boiling in 20 µl of 2x Laemmli buffer before Western blot analysis. Blots were incubated with primary antibodies (phosphotyrosine PY99, 1:500, no. sc-7020; Santa Cruz Biotechnology, Inc., Santa Cruz, CA; phosphoserine, 1:800, no. P3430, Sigma-Aldrich; IRS-1, 1:1000), and the bound antibody was detected by enhanced chemiluminescence reaction following incubation with secondary antibody conjugated with horseradish peroxidase.
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; Coronella-Wood et al., 2004). The conserved phosphorylation sites for activation of JNKs, ERK1/2, or p38 are Thr183/Tyr185, Thr202/Tyr204, or Thr180/Tyr182, respectively. Dose-response studies were performed to confirm that 100 µM H2O2 is the appropriate concentration to use for identifying the upstream regulators of MAPKs in two cell types: CMCs and HFs. The results show that although 30 µM H2O2 can induce phosphorylation of JNKs and ERKs in CMCs (Fig. 1A), obvious increases of phosphorylated forms of all three MAPKs at the signature sites were observed with H2O2 at 40 µM or more for both cell types (Fig. 1). The highest levels of phospho-JNKs, -ERKs, or -p38 were observed with 100 or 200 µM H2O2 (Fig. 1). The dose-response study suggests that 100 µM is the most appropriate dose to use because 200 µM H2O2 caused a decrease of phospho-ERKs from its peak level in CMCs (Fig. 1A) and reduced the level of total ERKs in HFs (Fig. 1B). Measurements of total JNKs, ERK1/2, and p38 protein levels indicate that H2O2 at 100 µM or less did not alter the protein level of these kinases (Fig. 1). Time-course studies comparing HFs with CMCs found that all three MAPKs increased their phosphorylation at the signature sites within 30 min of treatment with 100 µM H2O2 (Fig. 2). The induction of phosphorylated forms of JNKs, ERK1/2, and p38 seemed to reach a peak at 30 min and then began to decline after 60 min in both cell types (Fig. 2), with an exception of p38 in CMCs, which remained at a considerable level of phosphorylation even 2 h after H2O2 treatment (Fig. 2A). These data suggest 30 min is a suitable time frame for testing the upstream regulators of MAPK activation by oxidants.
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We determined the presence of EGFR in CMCs and HFs by Western blot analysis and found detectable levels of the receptor in both cell types (Fig. 3A). CMCs seem to contain a higher level of EGFR compared with HFs (Fig. 3A). Treatment with 5 ng/ml EGF for 10 min or 100 µM H2O2 for 10 min did not alter the level of the receptor (Fig. 3A). In certain experimental systems, chemical stress causes covalently linked dimerization of EGFR. We did not observe an increase in intensity of the bands at the molecular weight corresponding to EGFR dimers with H2O2 treatment in either cell type (Fig. 3A).
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One way to determine the involvement of the EGFR in mediating MAPK activation by oxidants is to measure the impact of EGF pretreatment, assuming that the pretreatment induces receptor internalization. EGF treatment alone caused peak phosphorylation of JNKs and ERK1/2 at 10 min in CMCs and HFs (Fig. 3, B and C). The levels of phosphorylated JNKs and ERK1/2 returned toward the basal level after 20 min (Fig. 3, B and C). The activation of p38 by EGF was minimal in either cell type at 10 min and not detectable at other time points (Fig. 3, B and C). Given the decline of EGF-induced MAPK activation after 20 min, this time point allows us to examine whether H2O2 can subsequently cause further activation of these kinases. After EGF pretreatment in CMCs, H2O2 failed to further activate JNKs and ERK1/2 (Fig. 3B). In contrast, EGF pretreatment in HFs did not alter the response to H2O2 in the activation of JNKs and ERK1/2 (Fig. 3C). In both cell types, EGF pretreatment slightly reduced the level of p38 activation by H2O2 without changing the time course (Fig. 3, B and C).
AG-1478 is an ATP-binding site inhibitor of EGFR tyrosine kinase, providing a tool to determine the involvement of EGFR in H2O2-induced MAPK activation. Treating cells with 250 nM AG-1478 prevented EGF from activating JNKs, ERK1/2, and p38 in either cell type (Fig. 3, B and C). The presence of AG-1478 also prevents EGF from altering the overall pattern of response to H2O2 (Fig. 3, B and C). In contrast, with H2O2 treatment, AG-1478 partially inhibited JNK and ERK1/2 activation without affecting p38 activation in CMCs and HFs significantly (Fig. 3, B and C).
PD158780, a competitive inhibitor of the EGFR tyrosine kinase ATP-binding site, differs from AG-1478 structurally and was used to verify the inhibitory effect of AG-1478 on H2O2-induced activation of JNKs, ERK1/2, and p38. CMCs or HFs were treated with 10 µM PD158780 before the exposure of H2O2. Results from PD158780 were essentially comparable to those of AG-1478 between the two cell types (Figs. 4, 5, 6).
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The nonreceptor Src family tyrosine kinase is known to be a key relay of signals from activated tyrosine kinase receptors such as EGFR to downstream targets. To inhibit the Src family of tyrosine kinases, we used the pharmacologic inhibitor PP2 (10 µM). PP3 is a structural analog of PP2 but does not have a Src inhibitory effect and therefore serves as a negative control. In CMCs and HFs, PP2 could inhibit the phosphorylation of JNKs and ERK1/2 after H2O2 treatment, whereas PP3 had no such inhibitory effect (Figs. 4 and 5). The inhibitory effect of PP2 on p38 activation was less significant in CMCs and was not detected in HFs (Fig. 6).
Several lines of evidence link matrix metalloproteinases (MMPs) to the transactivation of EGFR (Prenzel et al., 2001). Inhibitors of MMPs, GM6001 and BB2516, were used to test whether MMPs contribute to H2O2-induced MAPK activation. GM6001 and BB2516 are broad-spectrum hydroximate inhibitors of MMPs but are structurally distinct and have different potencies (Whittaker et al., 1999). An insignificant inhibition, if any, was observed on JNKs or ERK1/2 activation by H2O2 in CMCs with these two chemicals (Figs. 4A and 5A). In contrast, these two inhibitors were found to abolish JNK or ERK1/2 activation by H2O2 treatment in HFs (Figs. 4B and 5B). Neither MMP inhibitor significantly reduced the level of oxidant-induced p38 phosphorylation in each cell type (Fig. 6).
CMCs and HFs also express the FGFR (Liu et al., 1995), PDGFR (Brostrom et al., 2002), and insulin-like growth factor receptor (Velloso et al., 1998). The pharmacologic inhibitors of the FGFR tyrosine kinase or PDGFR tyrosine kinase allowed us to test whether these receptors serve as upstream regulators of ROS-induced MAPK activation. As Fig. 7 shows, inhibitors of FGFR and PDGFR, AG1295 (20 µM) and DMBI (10 µM), respectively, did not affect the phosphorylation of JNKs, ERK1/2, and p38 in CMCs or HFs after H2O2 treatment (Fig. 7, A and B).
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To explore whether the insulin receptor, which is known to be present on CMCs, might be activated by H2O2, we tested for tyrosine phosphorylation of IRS-1 because this event is immediately downstream of insulin receptor activation (Velloso et al., 1998). IRS-1 was isolated by immunoprecipitation, followed by Western blot analysis for tyrosine and serine phosphorylation. In contrast to the positive control treatment of insulin, IRS-1 was not tyrosine-phosphorylated by H2O2 treatment (Fig. 7C). Serine phosphorylation measurements also failed to detect a positive sign of IRS-1 phosphorylation in H2O2-treated cells (Fig. 7C). Collectively, these results suggest that the FGFR, PDGFR, or insulin receptor may not be significant contributors to MAPK activation by H2O2 treatment in CMCs and HFs.
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Whether EGFR mediates oxidative stress signaling during the progression of heart failure in vivo remains to be determined. It is commonly thought that the extracellular concentration of H2O2 caused by ischemia is approximately 6 µM, which increases to 11 µM during reperfusion in the myocardium (Zweier et al., 1987). Many cell types have been found to release H2O2 at a rate of 0.02 to 30 nmol/min/106 cells (Kinnula et al., 1991), suggesting the possibility that the lower doses of H2O2 used in this study are somewhat relevant to pathophysiologic conditions. Nevertheless, ischemia and reperfusion have been shown to induce activation of JNKs, ERKs, and p38 in the myocardium (Bogoyevitch et al., 1996; Yue et al., 2000). Ideally, a signature of EGFR activation, such as oxidant-specific phosphorylation sites, will be useful in determining the role of EGFR in MAPK activation in vivo and in understanding the mechanism underlying H2O2-induced EGFR activation. As the natural ligand of EGFR, EGF causes autophosphorylation of the EGFR at key residues, including Tyr1068, Tyr1148, and Tyr1173, which are critical for the propagation of the stimulatory signal through creation of binding sites for adaptors such as Grb2 and phospholipase C 
(Bertics et al., 1988; Sorkin et al., 1992). More recently, phosphorylation of Tyr845 in the kinase domain of EGFR has been linked to Src activity (Tice et al., 1999). However, our primary cell culture model did not accommodate reliable analysis of specific tyrosine phosphorylation sites on EGFR. Compared with renal proximal tubule epithelial cells or tumor cells, primary CMCs and HFs may not contain the level of EGFR sufficient for analysis of phosphorylation sites using traditional technology such as immunoprecipitation and Western blot. Detection of the level of EGFR protein in our experimental systems requires a method that is much more sensitive than regular Western blot analysis (Materials and Methods). In A549 lung carcinoma cells, H2O2 treatment led to strong phosphorylation of EGFR at Tyr845 and weaker phosphorylation of Tyr1068 and Tyr1173 (Ravid et al., 2002). In renal proximal tubular cells, H2O2 stimulates phosphorylation of EGFR at Tyr845 and Tyr1068 residues via an Src-mediated pathway (Zhougang and Schnellmann, 2004). These reports suggest the possibility that H2O2 may activate EGFR through multiple phosphorylation sites.
Pretreatment with EGF before H2O2 stimulation showed the relative importance of EGFR in mediating oxidant signaling in CMCs compared with HFs. In CMCs, EGF pretreatment presumably leads to EGFR internalization, and the lack of EGFR on the surface of the cells causes an inability of subsequent H2O2 treatment to activate JNKs and ERK1/2. An alternative explanation is that EGF pretreatment had drained the reservoir of intermediate signaling molecules needed to mediate H2O2-induced JNK and ERK1/2 activation. In either case, the dependence on EGFR is clear for JNKs and ERK1/2 activation in CMCs after oxidative stress. In contrast to the finding in CMCs, HFs failed to respond to EGF pretreatment by down-regulating H2O2-induced activation of JNKs and ERK1/2. AG-1478 seems to be a specific inhibitor for EGFR with an IC50 <3 nM and is not effective in inhibiting ErbB2 or other members of the EGFR family (IC50, >10,000 nM for ErbB2) (Mendelsohn and Baselga, 2000). The apparently conflicting data between EGF pretreatment and AG-1478 pretreatment in HFs suggest the possibility that dimerization with other members of the EGFR family may participate in the regulation of oxidant signaling in these cells. EGF can activate EGFR homodimers and EGFR-ErbB2 heterodimers, but it is less effective with ErbB1-3 and ErbB1-4 heterodimers and has no effect on other combinations (2-3, 2-3, 3-4) (Riese and Stern, 1998). The homodimers of EGFR undergo rapid internalization, whereas association of the activated EGFR with ErbB2 has a reduced internalization capacity but comparable signaling capacity relative to EGFR homodimers (Carpenter, 2000). Given the lower abundance of EGFR in HFs compared with CMCs (Fig. 2A), it is possible that heterodimerization with other ErbB family members is more significant on EGF stimulation in HFs; therefore, EGFR is not as vulnerable to down-regulation in HFs as in CMCs.
Our data with MMP inhibitors suggest that H2O2 may activate extracellular or membrane-bound MMPs in HFs. MMPs are capable of cleaving and therefore activating EGF-like factors from the extracellular matrix or those bound to the cell exterior. For example, the matrix protein laminin-5 contains an EGF-like domain that can be released by MMPs and therefore activate the EGFR (Schenk et al., 2003). In the case of EGFR transactivation by AngII, activated MMPs release heparin-binding EGF from the extracellular matrix to activate EGFR (Shah and Catt, 2003). H2O2 has been shown to activate MMPs in HFs (Siwik et al., 2001). Therefore, there are two scenarios that may mediate MAPK activation by H2O2: H2O2 activates EGFR via a ligand-independent manner or causes release of an EGF-like factor that in turn activates EGFR. The ligand-mediated activation of EGFR in HFs is supported by our data with MMP inhibitors. The failure of EGF pretreatment to alter H2O2-induced activation of ERK1/2 and JNKs also indirectly supports this ligand-dependent activation of EGFR theory in HFs. In contrast, the lack of an MMP inhibitor effect on CMCs suggests the possibility of ligand-independent activation of EGFR by H2O2 in these cells. The data with EGF pretreatment are consistent with this possibility because down-regulation of EGFR in CMCs by EGF pretreatment inhibited JNK or ERK1/2 activation by H2O2. In addition to these simplistic explanations, a triple membrane-passing signaling mechanism is postulated to link G protein-coupled receptors to receptor tyrosine kinase activation through membrane-bound MMPs (Prenzel et al., 2001). Kim et al. (2002) have shown that just such a system is activated when HFs are stimulated by the G protein-coupled receptor agonist isoproterenol. This suggests the possibility that HFs may possess this type of signaling web in mediating MAPK activation by H2O2.
Our data with the Src inhibitor PP2 parallel those observed with EGFR inhibitors, suggesting that Src and EGFR may lie in the same pathway leading to JNKs and ERK1/2 activation. In most circumstances, PP2 displayed a higher efficiency in inhibiting activation of ERK1/2 and JNKs by H2O2 than other inhibitors used in this study. Evidence has been accumulated indicating that H2O2 activates Src, which can mediate MAPK signaling during oxidative stress (Gawler, 1998; Chen et al., 2001a). Src can phosphorylate EGFR at Tyr845, causing stabilization of the activated state of the kinase domain (Sato et al., 1995; Leu and Maa, 2003). Upon phosphorylation of these sites, a well known network of SH2 and SH3 domain-containing proteins, such as Grb2, Ras, Raf, and Shc, are activated and mobilized from their membrane-associated positions to start the MAPK signaling cascade (Gawler, 1998). Conversely, Src is also a well known downstream target of EGFR tyrosine kinase. Therefore, in the case of ligand-independent activation of EGFR, the sequential relationship between Src and EGFR remains an open question. In conclusion, despite the fact that H2O2 can enter into cells through diffusion, we show here a role of a trans-membrane protein EGFR in mediating activation of MAPKs by oxidative stress.
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Footnotes |
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ABBREVIATIONS: CMC, cardiomyocyte; HF, heart fibroblast; ROS, reactive oxygen species; AngII, angiotensin II; MAPK, mitogen-activated protein kinase; JNK, c-Jun N-terminal kinase; ERK, extracellular signal-related kinase; EGF, epidermal growth factor; EGFR, epidermal growth factor receptor; PDGFR, platelet-derived growth factor receptor; FGFR, fibroblast growth factor receptor; DMEM, Dulbecco's modified Eagle's medium; FBS, fetal bovine serum; BB2516, [2S-[N4(R*),2R*,3S*]]-N4-[2,2-dimethyl-1-[(methylamino)carbonyl]propyl]-N1,2-dihydroxy-3-(2-methylpropyl)butanediamide (marimastat); AG-1478, 4-(3'-chloroanilino)-6,7-dimethoxy-quinazoline; IRS-1, insulin receptor substrate 1; PD158780, 4-[(3-bromophenyl)amino]-6-(methylamino)-pyrido[3,4-d]pyridimine; PP2, 4-amino-5-(4-chlorophenyl)-7-(t-butyl)pyrazolo[3,4-d]pyrimidine; PP3, 4-amino-7-phenylpyrazol[3,4-d]pyrimidine; MMP, matrix metalloproteinase; GM6001, N-[(2R)-2-(hydroxamidocarbonylmethyl)-4-methylpentanoyl]-L-tryptophan methylamide; AG1295, 6,7-dimethyl-2-phenylquinoxaline; DMBI, (Z)-3-[4-(dimethylamino) benzylidenyl] indolin-2-one.
Address correspondence to: Dr. Qin M. Chen, Department of Pharmacology, College of Medicine, University of Arizona, 1501 North Campbell Ave., Tucson, AZ 85724. E-mail: qchen{at}email.arizona.edu
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