Free Radical Biology and Medicine 86 (2015) 219–227

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Original Contribution

c-Jun N-terminal kinase attenuates TNFα signaling by reducing Nox1-dependent endosomal ROS production in vascular smooth muscle cells Hyehun Choi n, Anna Dikalova, Ryan J. Stark, Fred S. Lamb Department of Pediatrics, Vanderbilt University Medical Center, Nashville, TN 37232, USA

art ic l e i nf o Article history: Received 13 January 2015 Received in revised form 23 April 2015 Accepted 12 May 2015 Available online 19 May 2015 Keywords: Tumor necrosis factor-α NADPH oxidase 1 c-Jun N-terminal kinase Endosome Superoxide

a b s t r a c t Tumor necrosis factor-α (TNFα), a proinflammatory cytokine, causes vascular smooth muscle cell (VSMC) proliferation and migration and promotes inflammatory vascular lesions. Nuclear factor-kappa B (NF-κB) activation by TNFα requires endosomal superoxide production by Nox1. In endothelial cells, TNFα stimulates c-Jun N-terminal kinase (JNK), which inhibits NF-κB signaling. The mechanism by which JNK negatively regulates TNFα-induced NF-κB activation has not been defined. We hypothesized that JNK modulates NF-κB activation in VSMC, and does so via a Nox1-dependent mechanism. TNFαinduced NF-κB activation was TNFR1- and endocytosis-dependent. Inhibition of endocytosis with dominant-negative dynamin (DynK44A) potentiated TNFα-induced JNK activation, but decreased ERK activation, while p38 kinase phosphorylation was not altered. DynK44A attenuated intracellular, endosomal superoxide production in wild-type (WT) VSMC, but not in NADPH oxidase 1 (Nox1) knockout (KO) cells. siRNA targeting JNK1 or JNK2 potentiated, while a JNK activator (anisomycin) inhibited, TNFα-induced NF-κB activation in WT, but not in Nox1 KO cells. TNFα-stimulated superoxide generation was enhanced by JNK1 inhibition in WT, but not in Nox1 KO VSMC. These data suggest that JNK suppresses the inflammatory response to TNFα by reducing Nox1-dependent endosomal ROS production. JNK and endosomal superoxide may represent novel targets for pharmacologic modulation of TNFα signaling and vascular inflammation. & 2015 Elsevier Inc. All rights reserved.

Introduction Vascular smooth muscle cells (VSMCs) tightly regulate proliferation, contraction, inflammation, and extracellular matrix deposition and mitogen-activated protein kinases (MAPKs) play a critical role in these processes [1]. VSMCs are critical components of vascular inflammatory lesions, such as atherosclerotic plaques [2,3]. Circulating levels of the proinflammatory cytokine, tumor necrosis factor-α (TNFα), are elevated in patients with cardiovascular disease including

Abbreviations: ASK, apoptosis signal-regulating kinase; DynK44A, dominantnegative dynamin; eGFP, enhanced green fluorescent protein; ERK, extracellular signal-regulated kinase; ESR, electron spin resonance; JNK, c-Jun N-terminal kinase; MAPK, mitogen-activated protein kinase; NADPH, reduced nicotinamideadenine dinucleotide phosphate; NF-κB, nuclear factor-kappa B; Nox, NADPH oxidase; ROS, reactive oxygen species; siRNA, small interfering ribonucleic acid; TMH, 1-hydroxy-4-methoxy-2,2,6,6-tetramethylpiperidine; TNFα, tumor necrosis factor-α; TNFR, tumor necrosis factor-α receptor; TRAF, TNFR-associated factor; VSMC, vascular smooth muscle cell n Correspondence to: Vanderbilt University Medical Center, Department of Pediatrics, 2215 Garland Avenue, Light Hall-1055, Nashville, TN 37232-3122. Fax: þ 1 615 936 3467. E-mail address: [email protected] (H. Choi). http://dx.doi.org/10.1016/j.freeradbiomed.2015.05.015 0891-5849/& 2015 Elsevier Inc. All rights reserved.

hypertension [4], ischemic heart disease [5,6], and myocardial injury [7]. TNFα promotes VSMC proliferation and migration [8,9], thus contributing to the development of inflammatory vascular lesions [10]. TNFα signals are transduced through two distinct receptors, TNFα receptor 1 (TNFR1) and TNFR2. Depending on conditions, TNFR1 can activate VSMC proliferation, apoptosis, or necrosis. TNFR1 induces nuclear factor-kappa B (NF-κB) activation [11], which is associated with enhanced cell survival and proliferation, but also has a “death domain” which recruits adaptor proteins that support signaling pathways that lead to apoptosis and cell death [12]. In contrast, there is evidence that TNFR2 may actually exert a protective effect in the cardiovascular system, by an as yet unclear mechanism [13,14]. TNFR2 can mediate either NF-κB signal activation or termination via a TNFR-associated factor 2 (TRAF2)-dependent process [15]. Endocytosis regulates both activation and termination of signaling by ligand-bound receptors and TNFR1 internalization alters signal transduction [16,17]. Dynamin,
100 kDa GTPase, is an essential protein for TNFα receptor internalization [16,18]. Dynamin is required for membrane budding and pinches off endocytic vesicles, separating them from the plasma membrane [16,19]. We reported previously that in VSMC endosomes are a site where TNFα-induced superoxide and associated reactive oxygen species (ROS) are produced. Endosomal

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ROS are required for NF-κB activation and proliferation [20–22]. These “signaling” endosomes contain the nicotinamide adenine dinucleotide phosphate (NADPH) oxidase 1 (Nox1) [20,23], which is required for intravesicular superoxide production [24]. The critical role of Nox1 is demonstrated by the fact that VSMC proliferation and migration are reduced in Nox1 null cells and Nox1 plays a critical role in neointimal formation by supporting these events as well as extracellular matrix production [25]. Increased expression of Nox1 is observed in neointimal VSMCs where it promotes activation of matrix metalloproteinase9 [26]. Furthermore, Nox1 overexpression potentiates vascular smooth muscle hypertrophy in transgenic mice [27]. TNFα activates MAPKs, including c-Jun N-terminal kinase (JNK), p38 kinase, and extracellular signal-regulated kinase (ERK) [28]. These MAPKs mediate multiple downstream effects of TNFα including apoptosis, proliferation, and differentiation. We have shown previously that TNFα-mediated JNK and p38 activation are potentiated while ERK activation is not altered when endocytosis is inhibited in endothelial cells. These changes promote TNFα-induced cell death [29]. In contrast to endothelial cells, VSMCs tend to proliferate rather than die in response to TNFα [9]. We proposed that MAPKs affect NF-κB activation by TNFα, balancing ROSdependent inflammatory signaling in VSMC. The current study explores how subcellular localization of MAPK activation by TNFα modifies the inflammatory response of VSMC. Just as in endothelial cells, JNK activation, which does not require endocytosis of TNFα receptor, potently inhibits NF-κB activation. This effect of JNK is exerted via modulation of ROS production by endosomal Nox1.

Enzo Life Sciences (Farmingdale, NY). Other chemicals were purchased from Sigma-Aldrich (St. Louis, MO). Cultured cells Primary aortic VSMC from C57/BL6 mice were explanted as previously described [30]. Briefly, after mice were euthanized using carbon dioxide (CO2), thoracic aortas were excised and cleaned from fat tissue, and the endothelial layer was removed by passing the pins and cut into 2-to 4-mm-square sections in an ice-cold phosphatebuffered saline. The fragments were placed in the culture dish with medium and maintained in an incubator at 37 1C in a humidified 5% CO2 atmosphere. After 7–10 days, the tissues were removed and the migrated cells were maintained in Dulbecco's modified Eagle's medium (DMEM; Life Technologies, Grand Island, NY) supplemented with 10% fetal bovine serum (FBS), 1% penicillin/streptomycin, 1X minimum essential medium nonessential amino acids, 1X vitamin, and 20 mM Hepes. Nox1 KO cells [31] were a generous gift from Dr. Francis Miller (University of Iowa). siRNA transfection siRNA (negative control; D-001206-13, TNFR1; M-060201-01, TNFR2; M-043973-01, JNK1; M-040128-01, JNK2; M-040134-00) were purchased from Dharmacon (Lafayette, CO). siRNA (100 nM) was incubated with Lipofectamine 2000 (Life Technologies) in serum-free medium for 20 min. The resultant complex of siRNA– Lipofectamine 2000 was added to the cells in DMEM containing 5% FBS and then maintained for 3 days before performing experiments.

Materials and methods

NF-κB activity

Reagents

NF-κB induction was assessed by infection of VSMC with replication-deficient adenovirus containing a luciferase reporter gene driven by NF-κB transcriptional activation. Cells were infected for 40 h followed by exposure to TNFα (10 ng/mL) in

SP600125, SB203580, and U0126 were purchased from Cell Signaling Technology (Danvers, MA). Dynasore was obtained from

Fig. 1. TNFα-induced intracellular ROS production is attenuated by endocytosis inhibition in WT but not in Nox1 KO VSMCs. Total intracellular superoxide production was measured using TMH (A, C) or CAT1H (B, D). (A and B) TNFα increases superoxide production. *Po 0.05 compared to Control (n ¼3 to 4). (C and D) After infection with adenovirus of eGFP or DynK44A for 2 days, superoxide production was measured as described under Materials and methods. Results are presented as mean 7 SEM after normalization to protein concentration. *P o0.05 compared to AdeGFPþ TNFα in WT (n ¼4).

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serum-free DMEM for 6 h. Inhibitors and an activator were incubated for 30 min prior to TNFα exposure. In experiments requiring longer TNFα exposure (24 and 48 h), DMEM with 5% FBS was used. For siRNA transfected cells, adenovirus was infected 1 day after transfection of siRNA. Luciferase activity (relative light units) was measured in reporter lysis buffer, according to the protocol of the manufacturer (Promega, Madison, WI) and normalized to protein concentration (BCA protein assay). Adenoviral-mediated gene transfer Adenoviruses were obtained from Gene Transfer Vector Core at the University of Iowa (Iowa City, IA). Control viruses (eGFP) or the dynamin1 dominant-negative mutant (DynK44A) was added to VSMC (80% confluence, 10–30 MOI) in DMEM containing 5% FBS. Experiments were performed after 48 h. Western blot analysis Cells were serum-deprived (0.5% serum) for 3 h and then stimulated with TNFα (10 ng/mL). Protein extracts (40–60 μg) were separated by electrophoresis on a polyacrylamide gel (10%) and transferred to nitrocellulose membranes. Nonspecific binding was blocked with 5% skim milk in Tris-buffered saline solution with Tween 20 (0.1%) for 1 h at room-temperature. Membranes were then incubated with primary antibodies overnight at 4 1C. Antibodies were as follows: p-JNK, JNK, p-p38, p38, p-ERK, ERK (Cell Signaling Technology, Danvers, MA), tubulin (Vanderbilt Antibody Core, Nashville, TN), JNK1, and JNK2 (Santa Cruz Biotechnology, Dallas, TX). After incubation with fluorescent secondary antibodies, signals were developed using the Odyssey Imaging System (LI-COR Biosciences, Lincoln, NE) and quantified densitometrically. Results were normalized to the indicated protein and expressed as arbitrary units. Detection of reactive oxygen species Fluorescent detection of endosomal ROS was measured using OxyBURST Green H2HFF-BSA (Invitrogen, Life Technologies) [20]. In order to promote endocytosis of BSA-conjugated OxyBURST in VSMC, prior exposure to cationic ferritin was used as previously described [32]. Briefly, cells were grown on chamber slides. After washing with PBS, cationic ferritin (0.2 mg/mL) was incubated in PBS for 1 min at 37 1C and then quickly rinsed with PBS. Cells were then incubated in 50 mg/mL OxyBURST with or without TNFα at 37 1C. After 30 min, cells were fixed in 4% paraformaldehyde. Nuclear counterstaining was performed with To-Pro-3 (Life Technologies) for 10 min. Coverslides were mounted with ProLong Gold antifade reagent (Life Technologies) and imaged by fluorescence confocal microscopy. Fluorescence intensity was quantified using ImageJ software. Control experiments for quantification of endocytosis were performed with identical methods using dextran-conjugated to Texas Red (50 mg/mL, Life Technologies). For quantification of superoxide production, 1-hydroxy-4-methoxy-2,2,6,6-tetramethylpiperidine (TMH; Enzo Life Sciences) and 1-hydroxy-2,2,6,6-tetramethylpiperidin-4-yl-trimethylammonium chloride (CAT1H; Enzo Life Sciences) were used. TMH is cell permeable and used for detection of total intracellular superoxide. CAT1H is a cell-impermeable spin probe commonly used to detect extracellular superoxide which we also employed to detect endosomal superoxide [33,34]. After siRNA transfection of cells for 3 days or infection with adenovirus expressing eGFP or DynK44A for 2 days, the cells were incubated with Krebs/Hepes containing TMH (0.5 mmol/L) or CAT1H (0.5 mmol/L) and TNFα (10 ng/mL) for 20 min. The cells were then washed, scraped, snap-frozen, and placed in an electron spin resonance (ESR) finger Dewar under liquid nitrogen. ESR spectra were recorded using the following ESR settings: field sweep, 80 G;

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microwave frequency, 9.39 GHz; microwave power, 2 mW; modulation amplitude, 5 G; conversion time, 327.68 ms; time constant, 5242.88 ms; 512 points resolution; and receiver gain, 1  104. The signal was normalized to protein concentration. Statistical analysis Values are mean7standard error of the mean (SEM), and ‘n’ represents the number of independently performed experiments in cultured cells. Graphs were generated using Graph Pad Prism 5.0 (GraphPad Software, San Diego, CA). Statistical differences were calculated by Student's t test or one-way ANOVA. Post hoc comparisons were performed using Newman-Keuls analysis to compare all groups. A P value less than 0.05 was considered to be statistically significant.

Results TNFR1 and TNFR2 in NF-κB activation There are two distinct TNFα receptors, TNFR1 and TNFR2. To determine the contribution of these receptor subtypes to TNFαinduced inflammation in VSMC, we measured NF-κB activation in the presence of siRNA targeting TNFR1 or 2. TNFR1 siRNA completely blocked TNFα-induced NF-κB activation, but TNFR2 siRNA did not change the response significantly, despite very effective knockdown of mRNA (Suppl. Fig. 1). Thus, this response appears to be almost completely TNFR1 dependent.

WT Ad-virus eGFP TNFα

Dyn

Nox1 KO eGFP

y Dyn

- + - + - + - + p-JNK

JNK Tubulin

p-ERK

ERK Tubulin

p-p38 p38 Tubulin Fig. 2. Inhibition of endocytosis potentiates TNFα-induced phosphorylation of JNK, but diminishes phosphorylation of ERK. Western blots quantify phosphorylated and total MAPKs after 10 min exposure to TNFα (10 ng/mL) following infection with eGFP or DynK44A (Dyn). (A–C) Representative images show phosphorylated (p-) and total JNK (A), ERK (B), and p38 (C). Bar graphs show the relative abundance of phosphorylated proteins after normalization to total protein expression. Results are presented as mean 7SEM in each experimental group after normalization to control (eGFP in WT) quantities. *Po 0.05 compared to eGFP control in WT. †P o0.05. ‡Po 0.05 compared to eGFP with TNFα in WT (n ¼4 to 6).

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WT

Nox1 KO

WT

Nox1 KO

Fig. 3. NF-κB activation by TNFα is reduced by inhibition of endocytosis in both WT and Nox1 KO VSMC. Data are normalized to control levels (fold change) for WT or KO cells independently. (A and C) After 30 min incubation with DMSO or dynasore (15, 40, or 80 μM), TNFα was applied for 6 h. *Po 0.05 vs control. †Po 0.05 compared to TNFα without dynasore (n¼ 3 to 6). (B and D) Cells were infected with adenovirus expressing eGFP or DynK44A. *P o 0.05 vs eGFP only (control). †Po 0.05 (n¼ 4 to 6).

TNFα-induced intracellular ROS production TNFα-induced superoxide production was measured using the spin probe TMH for total intracellular, and CAT1H for endosomal superoxide. TNFα increased superoxide measured by both spin probes in WT cells (Fig. 1A and B). The increase in endosomal superoxide following TNFα was confirmed using OxyBURST Green (Suppl. Fig. 2). TNFα-induced superoxide production was not significantly altered by mitoTEMPO (25 nmol/L, 1 h), a mitochondrial selective antioxidant (TNFα 241.375.7, TNFα þ mitoTEMPO 22573.9, NS, n¼4). Both spin probes also demonstrate that WT cells have higher TNFα-induced superoxide production than Nox1 KO cells (Fig. 1C and D). Total intracellular (Fig. 1C) and endosomal (Fig. 1D) superoxide detected following TNFα treatment was reduced by inhibition of endocytosis with recombinant dominantnegative dynamin (AdDynK44A) in WT cells, but not in Nox1 KO cells. In addition, the quantitative decrease in superoxide production (pmol/mg of protein) induced by endocytosis inhibition (AdDynK44A) was similar for TMH, which distributes to the entire cell, and CAT1H, which can only enter the cell via endocytosis. Taken together, these data suggest that the majority of the intracellular superoxide produced by VSMCs in response to TNFα is produced by Nox1 and localized to newly formed endosomes. MAPK activation modulated by endocytosis The impact of receptor endocytosis on TNFα-induced MAPK (JNK, ERK, p38) activation was investigated. In VSMC, phosphorylation of MAPK by TNFα peaks after 10–15 min of stimulation and declines thereafter [9,35]. We tested phosphorylation of MAPK after 10 min of TNFα stimulation and observed significant phosphorylation of JNK, ERK, and p38. Inhibition of endocytosis by DynK44A caused differential modulation of MAPK pathways, resulting in potentiation of JNK and reduction of ERK

phosphorylation in WT VSMC (Fig. 2A and B). In contrast, Nox1 KO cells demonstrated no significant increase in JNK or ERK phosphorylation following TNFα. Phosphorylation of p38 by TNFα in KO cells was still significant, but was greatly reduced compared to WT cells (Fig. 2C). Endocytosis inhibition by DynK44A in Nox1 KO cells did not further modify JNK or ERK activation, but blocked the weak p38 activation observed in Nox1 KO cells (Fig. 2). Importantly, Nox1 KO cells do not have a generalized defect in endocytosis, uptake of 10,000 MW Texas Red-labeled dextran was similar in WT and KO cells (Suppl. Fig. 3).

Endocytosis-dependent NF-κB activation NF-κB activation was measured in WT and Nox1 KO VSMC. The increment in NF-κB activity induced by TNFα was greater in WT than in Nox1 KO cells (Fig. 3). In order to determine whether TNFα-mediated NF-κB activation is endocytosis dependent, a dynamin inhibitor (dynasore) [36] or recombinant dominantnegative dynamin (Ad-DynK44A) [37] was used. In WT and Nox1 KO VSMC, NF-κB activation was reduced by both dynasore (Fig. 3A and C) and Ad-DynK44A (Fig. 3B and D). The effect of dynasore was concentration dependent. NF-κB activation in WT cells was significantly reduced by 15 mM dynasore, a concentration that had no effect on the response to TNFα in Nox1 KO cells. The response of KO cells was reduced by 40 or 80 mM dynasore, reflecting reduced sensitivity to endocytosis inhibition compared to WT cells (Fig. 3A and C). Ad-DynK44A reduced resting NF-κB activity as well as the response to TNFα in both WT and KO cells (Fig. 3B and D). These data suggest that Nox1 KO cells have developed an alternative mechanism of NF-κB activation that does not require Nox1. This pathway has reduced sensitivity to endocytosis inhibition compared to normal TNFα signaling.

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Fig. 4. TNFα-induced NF-κB activation is reduced by ERK inhibition (U0126) and enhanced by JNK inhibition (SP600125) in WT VSMC. Results are normalized to the control value. (A) Inhibitors were incubated for 30 min prior to TNFα stimulation. After 6 h incubation with TNFα, U0126 attenuated, but SP600125 potentiated, NF-kB activation. C, Control (DMSO); U, U0126, ERK inhibitor (10 μM); SB, SB203580, p38 inhibitor (10 μM); SP, SP600125, JNK inhibitor (3 μM). *Po 0.05 vs control. †Po 0.05 (n¼4). (B) Inhibition of NF-κB activation by dynasore (40 μM) persists in the presence of SP600125. *P o0.05 vs control. †Po 0.05 (n¼ 6). C: Time dependence of enhancement of NFκB activation in response to TNFα by SP600125. *P o0.05 vs TNFα in each time period (n ¼3 to 7).

Impact of MAPK on NF-κB activation

Attenuation of NF-κB activation by JNK requires Nox1

Previously we have shown that in endothelial cells, NF-κB activation by TNFα is potentiated by JNK inhibition [29]. To determine whether this effect is preserved across other cell types involved in vascular inflammation, we tested the effect of JNK and other MAPK inhibitors on TNFα-induced VSMC inflammation. In WT VSMC, MAPK inhibitors did not affect NF-κB activity at rest, in the absence of TNFα stimulation. SB203580 (10 mM), a p38 inhibitor, did not alter TNFα-induced NF-κB activation while ERK inhibition by U0126 (10 mM) modestly attenuated TNFα-induced NF-κB activation. As in endothelial cells, a JNK inhibitor (SP600125, 3 mM) significantly potentiated the response to TNFα (Fig. 4A). To consider whether this potentiating effect of SP600125 on NF-κB activation was related to endocytosis, SP600125 was coincubated with dynasore. In the presence of dynasore, SP600125 no longer increased NF-κB activation (Fig. 4B), suggesting that JNK attenuates steps in NF-κB signaling that occur subsequent to endocytosis. Thus, we determined whether JNK alters the endocytosis process itself. Inhibition of JNK did not affect TNFα-induced endocytosis of labeled dextran (Suppl. Fig. 4). We next investigated the time dependence of the ability of JNK inhibition to potentiate NF-κB signaling. NF-κB activation in response to TNFα was maximal at 6 h and maintained for 24 h, but returned to baseline after 48 h. JNK inhibition significantly potentiated NF-κB activation in response to TNFα at 6, 16, and 24 h, but unlike what we observed in endothelial cells [29], SP600125 did not extend the duration of NF-κB activation (Fig. 4C).

Pharmacologic inhibition of JNK increased NF-κB signaling (Fig. 4). We sought to confirm this effect using siRNA targeting of JNK1 and JNK2. These siRNA were very effective at selectively suppressing protein abundance for the JNK isoforms (Suppl. Fig. 5). We also considered whether the increase in NF-κB activation induced by JNK inhibition required Nox1. This seemed plausible because NF-κB activation requires formation of signaling endosomes containing Nox1 [20]. First, we confirmed that in WT cells, both JNK1 and JNK2 siRNA potentiated NF-κB activation by TNFα (Fig. 5A). Similar effects were observed when JNK1 and JNK2 siRNA were combined (data not shown). However, in comparison to WT cells, inhibition of JNK1 or JNK2 using siRNA did not affect NF-κB activation in Nox1 KO cells following TNFα exposure (Fig. 5B). To further confirm that JNK activation inhibits NF-κB signaling, we used an additional pharmacological tool, a JNK activator (Anisomycin, 0.1 mM). As shown in Fig. 5C and D, anisomycin blocked TNFα-induced NF-κB activation in WT, but not in Nox1 KO cells. These data suggest that JNK attenuates NF-κB signaling by TNFα in a Nox1-dependent manner. Inhibition of ROS production by JNK Since Nox1 is required for JNK to have an impact on NF-κB activation, we hypothesized that JNK might directly affect Nox1 activity. We therefore measured endosomal ROS production in both WT and Nox1 KO VSMC. Endosomal superoxide production was selectively detected using OxyBURST green fluorescence (H2HFF-BSA). JNK1 siRNA

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WT

Nox1 KO

Fig. 5. JNK inhibits Nox1-dependent NF-κB activation. After transfection with siRNA (Control, JNK1, or JNK2), NF-κB activation was measured. (A) siRNA targeting JNK1 or 2 increased NF-κB activation in WT VSMC (n¼ 6 to 8). (B) JNK1 and 2 siRNA did not alter the effect of TNFα on NF-κB activation in Nox1 KO VSMC (n¼ 6 to 13). *Po 0.05 vs control in control siRNA. †Po 0.05 compared to TNFα in control siRNA. (C and D) Anisomycin (0.1 μM), a JNK activator blocked NF-κB activation by TNFα in WT not Nox1 KO cells. *Po 0.05 vs control. †Po 0.05 (n¼ 6).

enhanced the TNFα-stimulated OxyBURST signal in WT VSMCs. By comparison, minimal superoxide production was detected with or without JNK1 siRNA in Nox1 KO cells (Fig. 6A). In order to confirm these observations, we also detected intracellular superoxide production using TMH. These experiments confirmed that inhibition of JNK1 with siRNA potentiated superoxide generation by TNFα compared to control siRNA in WT. In comparison with WT, Nox1 KO cells produced less superoxide and siJNK1 did not change this (Fig. 6B). Finally, we used CAT1H to quantitatively assay endosomal superoxide and observed a very similar effect of siJNK (siControlþTNFα 21779, siJNK1þTNFα 32477 pmol/mg of protein, n¼4). All of these data suggest that suppression of Nox1-dependent endosomal superoxide production is associated with JNK activation following exposure to TNFα.

Discussion The key finding of this study is that JNK regulates NF-κB activation by TNFα via TNFR1-dependent modulation of Nox1dependent superoxide production. Inhibition of endocytosis enhances JNK phosphorylation by TNFα, suggesting that this process occurs without receptor endocytosis. In contrast, dynamin-dependent endocytosis and Nox1 are required for ERK1/2 and NF-κB activation. Despite this difference in localization, JNK inhibits Nox1-dependent endosomal superoxide production and suppresses subsequent NF-κB activation (Fig. 7). Endocytosis of receptors is known to affect signal transduction and this phenomenon has been widely investigated [38]. TNFR1 endocytosis can impact both proapoptotic and antiapoptotic signaling [16]. We

have shown previously that in VSMC, endosomal ROS production is required for NF-κB activation [20]. This endocytosis-dependent signaling pathway also exists in endothelial cells and inhibition of JNK promotes NF-κB activation, cell survival, and proliferation [29]. In the present study, we show that TNFα-induced endosomal superoxide production and subsequent NF-κB activation follow endocytosis of TNFR1 and this signaling is similarly modulated by JNK. This effect of JNK appears to be mediated by reduction of Nox1-dependent superoxide production. It remains to be determined if activated JNK phosphorylates its relevant target proteins before the endocytic vesicles leave the plasma membrane, or if JNK acts intracellularly at the level of the endocytic vesicle or early endosome. However, JNK does not exert its effect by disrupting the endocytic process itself (Suppl. Fig. 4). Nox1-dependent superoxide production is required for activation of NF-κB by TNFα. Antioxidant treatment (N-acetylcysteine), as well as both acute (diphenylene iodonium, DPI) and subacute (Nox1 antisense or dominant-negative Rac1) inhibition of Nox1, all dramatically impair NF-κB activation by TNFα in VSMC [20]. In contrast, we now observe that Nox1 KO cells have effective TNFα-stimulated NFκB activity, even though this effect of TNFα is blunted compared to WT (Fig. 3). These data provide evidence of compensation for the chronic loss of Nox1. Through this alternative mechanism, Nox1 KO cells activate NF-κB following TNFα exposure, but this pathway has reduced sensitivity to dynasore (Fig. 3C). This suggests that a larger component of this compensatory signal pathway originates at the plasma membrane. Further study will be required to define the mechanism of NF-κB activation in Nox1 KO VSMC. Multiple MAPKs are activated by TNFα in VSMC [28,35]. We confirm that TNFα increases phosphorylation of JNK, ERK, and p38 in WT VSMC. We have previously shown that JNK activation by

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siControl+TNFα

225

siJNK1+TNFα

WT

Nox1KO

WT siControl +TNFα

WT siJNK1 +TNFα

KO siControl +TNFα

40G

KO siJNK1 +TNFα

Fig. 6. TNFα-induced ROS production is potentiated by JNK1 inhibition in WT VSMC, but not in Nox1 KO cells. (A) After transfection with siRNA targeting control sequence or JNK1 for 3 days, endosomal ROS were measured with OxyBURST green fluorescence as described under Materials and methods. Quantification of OxyBURST signal showed an increase associated with JNK1 inhibition. Confocal images are representative of three independent experiments. *†P o0.05 compared to siControlþ TNFα in WT. (B) TMH, quantifying total intracellular superoxide production following TNFα was reduced in Nox1 KO cells compared to WT and increased by JNK1 inhibition only in WT cells. Representative ESR spectra for TMH (left) and mean data (right). *P o0.05 compared to siControlþ TNFα in WT (n¼4).

TNFα is potentiated by endocytosis inhibition in endothelial cells [29]. We now observe a similar effect in VSMC, suggesting once again that JNK activation occurs even in the absence of TNFR endocytosis. Interestingly, in endothelial cells, ERK activation was not affected by endocytosis inhibition [29], while in VSMC, ERK phosphorylation was markedly diminished, indicating that ERK1/2 activation occurs primarily following TNFR endocytosis. Thus, the impact of receptor endocytosis on TNFα-induced MAPK signaling appears to be cell type dependent. Both JNK and ERK activation were disrupted in Nox1 KO VSMC, suggesting that superoxide produced by Nox1 is required for MAPK signaling events occurring both at the plasma membrane and in endosomes. Previous studies have demonstrated that downstream changes in gene expression related to NF-κB activation can inhibit subsequent JNK activation [39,40]. Suppression of NF-κB induces the JNK cascade through the transcriptional upregulation of gadd45beta, which attenuates JNK signaling [40]. The IκB kinase/NF-κB pathway inhibits TNFα-mediated JNK activation, partly through Xchromosome-linked inhibitor of apoptosis protein [39]. We now confirm our previous observation that the converse also occurs: JNK activation suppresses NF-κB signaling in endothelial cells [29] and in VSMC (Figs. 4 and 5). In endothelial cells, JNK inhibition potentiates NF-κB activation in response to TNFα and this effect is maintained for over 48 h [29]. By comparison, the magnitude and duration of this effect were somewhat smaller in VSMC. Considered broadly, JNK and p38 signaling favor cell death [41], whereas ERK1/2 activation promotes proliferation and cell survival

[42]. TNFα signaling can impact both of these opposing directions of cell fate. Just as in endothelial cells [29], the balance of these influences is affected by TNFR endocytosis in VSMC. The ability of JNK and subsequent c-Jun activation to induce apoptosis is well documented [43]. However, the mechanism by which JNK modulates cell proliferation, and may even promote oncogenesis, is not nearly as well defined. JNK activation is an important signaling step in apoptosis, inflammation, and matrix degradation [44,45]. There are three isoforms, JNK1, 2, and 3. JNK1 and 2 are ubiquitously expressed while JNK3 is restricted to the brain, heart, and testes. JNK1 and 2 can have distinct substrate affinities [46] and play differential signaling roles. We observed no significant difference in the ability of siRNA targeted to JNK1 or 2 to enhance TNFαinduced NF-κB activation (Fig. 5). Inhibition of JNK signaling with SP600125, which targets all isoforms of JNK, has been demonstrated to suppress cartilage and bone destruction in inflammatory arthritis [47] and was beneficial in cardiomyopathy caused by mutation of the lamin A/C gene [48]. However, JNK1-deficient mice were not protected from TNF-mediated arthritis [49]. JNK1deficient mice spontaneously develop intestinal tumors [50] and are more prone to develop skin tumors following treatment with TPA (12-O-tetradecanoylphorbol-13-acetate) [51]. In view of the known ability of inflammation to promote oncogenesis, these phenotypes may be related to impairment of JNK-mediated suppression of NF-κB as described herein. Our work also suggests that site specificity of JNK activation adds an important additional layer of complexity to its role in TNFα signaling.

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metabolism. The next generation of anti-inflammatory agents may need to target ROS in a site-specific manner.

TNFR1

Nox1

TNFα Plasma Membrane

Conclusions Endosome

JNK

O2·Nox1

O

TNFR1

ERK

JNK attenuates TNFα-mediated inflammatory response by reducing Nox1-dependent endosomal superoxide production. This study demonstrates that JNK is a critical regulator of the complex inflammatory balance between cell proliferation and death that is induced by TNFR1. Optimal modification of TNFα signaling for clinical benefit will require a complete understanding of this complex process in order to develop anti-inflammatory agents that target ROS in a site-specific manner.

NF-κB

Conflict of interest Fig. 7. Overview of TNFα-mediated signaling in VSMCs. In response to TNFα, TNFR1 triggers superoxide production by Nox1 within endosomes. Interfering with endocytosis completely blocks intracellular superoxide (O2  –) production and alters activation of both ERK and JNK. TNFα-induced NF-κB activation is associated with TNFR1 endocytosis and is partly dependent on ERK activation. JNK activation is potentiated when endocytosis is inhibited, suggesting that the process occurs without endocytosis. JNK activation reduces NF-κB activation by suppression of Nox1-dependent ROS production.

Our results provide the first demonstration of a specific mechanism for JNK modulation of NF-κB activation, whereby JNK reduces Nox1-dependent ROS production. There are several links between JNK, Nox1, and ROS. JNK, also known as stressactivated protein kinase, is known to be activated by ROS [52]. TNFα-induced JNK activation is dependent on the Nox complex protein p47phox and is inhibited by antioxidant treatment [53]. Coexpression of TRAF4 and p47phox increases ROS production and JNK activation [54]. JNK activation by ROS can occur through the apoptosis signal-regulating kinase 1 (ASK1) MAPKKK, which activates the MAPKK-JNK signal pathway. ASK1 is activated when oxidation of thioredoxin causes it to dissociate from its complex with ASK1 [55,56]. In addition to ASK1, Src kinase can also mediate JNK activation by ROS [57]. The current finding that inhibition of JNK1 potentiates TNFα-induced endosomal superoxide production in VSMC suggests that JNK activation by ROS may provide a negative feedback loop for control of ROS production by Nox1. There are over 50 known substrates for JNK. These include nuclear targets such as transcription factors (c-Jun, ATF2, Elk-1, c-myc, p53) and nuclear hormone receptors (PPARγ, glucocorticoid receptor, retinoic acid receptor). JNK also phosphorylates cytoplasmic proteins including: (1) adaptor proteins (JIP1 and 3 and 14-3-3), (2) pro- and antiapoptotic mitochondrial proteins such as Bim and Bcl2, (3) other kinases such as p90RSK and Akt, and (4) regulators of cell movement such as paxillin and kinesin heavy chain [58]. Like many of these other JNK targets, Nox1 activity clearly impacts cell fate and motility [27,59]. Thus it seems plausible that Nox1 or a member of the Nox1 protein complex (likely to include p22phox, p47phox, NoxA1, and Rac1 in VSMC) is also a substrate for JNK. Recently, Nox1 was shown to be phosphorylated at T429 by PKCβ1 following exposure of VSMCs to TNFα [60]. Further work will be required to define the molecular mechanism by which JNK negatively modulates Nox1 activity. Increased superoxide production leads to excessive VSMC growth, endothelial dysfunction, monocyte migration, lipid peroxidation, inflammation, and other processes contributing to vascular damage [61,62]. Vascular injury by ROS contributes to the development of cardiovascular diseases, such as atherosclerosis, hypertension, and heart failure [63]. However, clinical studies have generally not shown significant benefit of nonselective antioxidant therapy on the development or progression of cardiovascular diseases [64–66]. The current results emphasize the complex role of Nox1 and ROS in inflammatory signaling and the need to fully understand ROS localization and

The authors declare that they have no conflict of interests.

Acknowledgment This project was supported by a post-doctoral fellowship from the American Heart Association [13POST16950048 to H.C.].

Appendix A. Supporting information Supplementary data associated with this article can be found in the online version at http://dx.doi.org/10.1016/j.freeradbiomed. 2015.05.015. References [1] Sprague, A. H.; Khalil, R. A. Inflammatory cytokines in vascular dysfunction and vascular disease. Biochem. Pharmacol. 78:539–552; 2009. [2] Dzau, V. J.; Braun-Dullaeus, R. C.; Sedding, D. G. Vascular proliferation and atherosclerosis: NEw perspectives and therapeutic strategies. Nat. Med 8:1249–1256; 2002. [3] Feil, S.; Fehrenbacher, B.; Lukowski, R.; Essmann, F.; Schulze-Osthoff, K.; Schaller, M.; Feil, R. Transdifferentiation of vascular smooth muscle cells to macrophage-like cells during atherogenesis. Circ. Res. 115:662–667; 2014. [4] Genctoy, G.; Altun, B.; Alper Kiykim, A.; Arici, M.; Erdem, Y.; Çağlar, M.; Yasavul, Ü.; Turgan, Ç.; Çağlar, Ş. TNF alpha  308 genotype and renin– angiotensin system in hemodialysis patients: an effect on inflammatory cytokine levels? Artific. Organs 29:174–178; 2005. [5] Vaddi, K.; Nicolini, F. A.; Mehta, P.; Mehta, J. L. Increased secretion of tumor necrosis factor-alpha and interferon-gamma by mononuclear leukocytes in patients with ischemic heart disease. Relevance in superoxide anion generation. Circulation 90:694–699; 1994. [6] Tentolouris, C.; Tousoulis, D.; Antoniades, C.; Bosinakou, E.; Kotsopoulou, M.; Trikas, A.; Toutouzas, P.; Stefanadis, C. Endothelial function and proinflammatory cytokines in patients with ischemic heart disease and dilated cardiomyopathy. Int. J. Cardiol. 94:301–305; 2004. [7] Matsumori, A.; Yamada, T.; Suzuki, H.; Matoba, Y.; Sasayama, S. Increased circulating cytokines in patients with myocarditis and cardiomyopathy. Br. Heart J. 72:561–566; 1994. [8] Peppel, K.; Zhang, L.; Orman, E. S.; Hagen, P. O.; Amalfitano, A.; Brian, L.; Freedman, N. J. Activation of vascular smooth muscle cells by TNF and PDGF: overlapping and complementary signal transduction mechanisms. Cardiovasc. Res. 65:674–682; 2005. [9] Goetze, S.; Xi, X. P.; Kawano, Y.; Kawano, H.; Fleck, E.; Hsueh, W. A.; Law, R. E. TNF-alpha-induced migration of vascular smooth muscle cells is MAPK dependent. Hypertension 33:183–189; 1999. [10] Zhang, H.; Park, Y.; Wu, J.; Chen, X.; Lee, S.; Yang, J.; Dellsperger, K. C.; Zhang, C. Role of TNF-alpha in vascular dysfunction. Clin. Sci. (Lond.) 116:219–230; 2009. [11] Wajant, H.; Scheurich, P. TNFR1-induced activation of the classical NF-kappaB pathway. FEBS J. 278:862–876; 2011. [12] Cabal-Hierro, L.; Lazo, P. S. Signal transduction by tumor necrosis factor receptors. Cell Signal. 24:1297–1305; 2012. [13] Luo, D.; Luo, Y.; He, Y.; Zhang, H.; Zhang, R.; Li, X.; Dobrucki, W. L.; Sinusas, A. J.; Sessa, W. C.; Min, W. Differential functions of tumor necrosis factor receptor 1 and 2 signaling in ischemia-mediated arteriogenesis and angiogenesis. Am. J. Pathol. 169:1886–1898; 2006.

H. Choi et al. / Free Radical Biology and Medicine 86 (2015) 219–227

[14] Higuchi, Y.; McTiernan, C. F.; Frye, C. B.; McGowan, B. S.; Chan, T. O.; Feldman, A. M. Tumor necrosis factor receptors 1 and 2 differentially regulate survival, cardiac dysfunction, and remodeling in transgenic mice with tumor necrosis factor-alpha-induced cardiomyopathy. Circulation 109:1892–1897; 2004. [15] Rodriguez, M.; Cabal-Hierro, L.; Carcedo, M. T.; Iglesias, J. M.; Artime, N.; Darnay, B. G.; Lazo, P. S. NF-kappaB signal triggering and termination by tumor necrosis factor receptor 2. J. Biol. Chem. 286:22814–22824; 2011. [16] Schutze, S.; Tchikov, V.; Schneider-Brachert, W. Regulation of TNFR1 and CD95 signalling by receptor compartmentalization. Nat. Rev. Mol. Cell. Biol. 9:655–662; 2008. [17] Schneider-Brachert, W.; Tchikov, V.; Neumeyer, J.; Jakob, M.; Winoto-Morbach, S.; Held-Feindt, J.; Heinrich, M.; Merkel, O.; Ehrenschwender, M.; Adam, D.; Mentlein, R.; Kabelitz, D.; Schutze, S. Compartmentalization of TNF receptor 1 signaling: internalized TNF receptosomes as death signaling vesicles. Immunity 21:415–428; 2004. [18] Loerke, D.; Mettlen, M.; Yarar, D.; Jaqaman, K.; Jaqaman, H.; Danuser, G.; Schmid, S. L. Cargo and dynamin regulate clathrin-coated pit maturation. PLoS Biol. 7:e1000057; 2009. [19] Henley, J. R.; Krueger, E. W. A.; Oswald, B. J.; McNiven, M. A. Dynaminmediated Internalization of Caveolae. J. Cell Biol. 141:85–99; 1998. [20] Miller, F. J.; Filali, M.; Huss, G. J.; Stanic, B.; Chamseddine, A.; Barna, T. J.; Lamb, F. S. Cytokine activation of nuclear factor κb in vascular smooth muscle cells requires signaling endosomes containing Nox1 and ClC-3. Circ. Res. 101:663–671; 2007. [21] Miller Jr. F. J.; Chu, X.; Stanic, B.; Tian, X.; Sharma, R. V.; Davisson, R. L.; Lamb, F. S. A differential role for endocytosis in receptor-mediated activation of Nox1. Antioxid. Redox Signal. 12:583–593; 2010. [22] Matsuda, J. J.; Filali, M. S.; Moreland, J. G.; Miller, F. J.; Lamb, F. S. Activation of swelling-activated chloride current by tumor necrosis factor-α requires ClC-3dependent endosomal reactive oxygen production. J. Biol. Chem. 285:22864–22873; 2010. [23] Lassegue, B.; Clempus, R. E. Vascular NAD(P)H oxidases: specific features, expression, and regulation. Am. J. Physiol. Regul. Integr. Comp. Physiol 285: R277–R297; 2003. [24] Brandes, R. P.; Kreuzer, J. Vascular NADPH oxidases: molecular mechanisms of activation. Cardiovasc. Res. 65:16–27; 2005. [25] Lee, M. Y.; San Martin, A.; Mehta, P. K.; Dikalova, A. E.; Garrido, A. M.; Datla, S. R.; Lyons, E.; Krause, K. H.; Banfi, B.; Lambeth, J. D.; Lassegue, B.; Griendling, K. K. Mechanisms of vascular smooth muscle NADPH oxidase 1 (Nox1) contribution to injury-induced neointimal formation. Arterioscler. Thromb. Vasc. Biol. 29:480–487; 2009. [26] Xu, S.; Shriver, A. S.; Jagadeesha, D. K.; Chamseddine, A. H.; Szocs, K.; Weintraub, N. L.; Griendling, K. K.; Bhalla, R. C.; Miller Jr. F. J. Increased expression of Nox1 in neointimal smooth muscle cells promotes activation of matrix metalloproteinase-9. J. Vasc. Res. 49:242–248; 2012. [27] Dikalova, A.; Clempus, R.; Lassegue, B.; Cheng, G.; McCoy, J.; Dikalov, S.; San Martin, A.; Lyle, A.; Weber, D. S.; Weiss, D.; Taylor, W. R.; Schmidt, H. H.; Owens, G. K.; Lambeth, J. D.; Griendling, K. K. Nox1 overexpression potentiates angiotensin II-induced hypertension and vascular smooth muscle hypertrophy in transgenic mice. Circulation 112:2668–2676; 2005. [28] Aggarwal, B. B. Signalling pathways of the TNF superfamily: a double-edged sword. Nat. Rev. Immunol. 3:745–756; 2003. [29] Choi, H.; Nguyen, H. N.; Lamb, F. S. Inhibition of endocytosis exacerbates TNFα-induced endothelial dysfunction via enhanced JNK and p38 activation. Am. J. Physiol. Heart Circ. Physiol 306:H1154–H1163; 2014. [30] Metz, R.; Patterson, J.; Wilson, E. Vascular smooth muscle cells: isolation, culture, and characterization. In: Peng, X., Antonyak, M., editors. Cardiovascular development. Humana Press; 2012. p. 169–176. [31] Zimmerman, M. C.; Takapoo, M.; Jagadeesha, D. K.; Stanic, B.; Banfi, B.; Bhalla, R. C.; Miller Jr. F. J. Activation of NADPH oxidase 1 increases intracellular calcium and migration of smooth muscle cells. Hypertension 58:446–453; 2011. [32] Sprague, E. A.; Kelley, J. L.; Suenram, C. A.; Valente, A. J.; Abreu-Macomber, M.; Schwartz, C. J. Stimulation of albumin endocytosis by cationized ferritin in cultured aortic smooth muscle cells. Am. J. Pathol. 121:433–443; 1985. [33] Widder, J. D.; Chen, W.; Li, L.; Dikalov, S.; Thöny, B.; Hatakeyama, K.; Harrison, D. G. Regulation of tetrahydrobiopterin biosynthesis by shear stress. Circ. Res. 101:830–838; 2007. [34] Dikalov, S. I.; Kirilyuk, I. A.; Voinov, M.; Grigor'ev, I. A. EPR detection of cellular and mitochondrial superoxide using cyclic hydroxylamines. Free Radic. Res. 45:417–430; 2011. [35] Camici, G. G.; Steffel, J.; Akhmedov, A.; Schafer, N.; Baldinger, J.; Schulz, U.; Shojaati, K.; Matter, C. M.; Yang, Z.; Luscher, T. F.; Tanner, F. C. Dimethyl sulfoxide inhibits tissue factor expression, thrombus formation, and vascular smooth muscle cell activation: a potential treatment strategy for drug-eluting stents. Circulation 114:1512–1521; 2006. [36] Macia, E.; Ehrlich, M.; Massol, R.; Boucrot, E.; Brunner, C.; Kirchhausen, T. Dynasore, a cell-permeable inhibitor of dynamin. Dev. Cell 10:839–850; 2006. [37] Damke, H.; Baba, T.; Warnock, D. E.; Schmid, S. L. Induction of mutant dynamin specifically blocks endocytic coated vesicle formation. J. Cell Biol. 127:915–934; 1994. [38] Sorkin, A.; von Zastrow, M. Endocytosis and signalling: intertwining molecular networks. Nat. Rev. Mol. Cell. Biol. 10:609–622; 2009.

227

[39] Tang, G.; Minemoto, Y.; Dibling, B.; Purcell, N. H.; Li, Z.; Karin, M.; Lin, A. Inhibition of JNK activation through NF-kappaB target genes. Nature 414:313–317; 2001. [40] De Smaele, E.; Zazzeroni, F.; Papa, S.; Nguyen, D. U.; Jin, R.; Jones, J.; Cong, R.; Franzoso, G. Induction of gadd45beta by NF-kappaB downregulates proapoptotic JNK signalling. Nature 414:308–313; 2001. [41] Matsuzawa, A.; Ichijo, H. Stress-responsive protein kinases in redox-regulated apoptosis signaling. Antioxid. Redox Signal. 7:472–481; 2005. [42] Nelson, P. R.; Yamamura, S.; Mureebe, L.; Itoh, H.; Kent, K. C. Smooth muscle cell migration and proliferation are mediated by distinct phases of activation of the intracellular messenger mitogen-activated protein kinase. J. Vasc. Surg. 27:117–125; 1998. [43] Dhanasekaran, D. N.; Reddy, E. P. JNK signaling in apoptosis. Oncogene 27:6245–6251; 2008. [44] 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. Requirement of JNK for stress-induced activation of the cytochrome c-mediated death pathway. Science 288:870–874; 2000. [45] Kyriakis, J. M.; Avruch, J. Sounding the alarm: protein kinase cascades activated by stress and inflammation. J. Biol. Chem. 271:24313–24316; 1996. [46] Gupta, S.; Barrett, T.; Whitmarsh, A. J.; Cavanagh, J.; Sluss, H. K.; Derijard, B.; Davis, R. J. Selective interaction of JNK protein kinase isoforms with transcription factors. EMBO J. 15:2760–2770; 1996. [47] Han, Z.; Boyle, D. L.; Chang, L.; Bennett, B.; Karin, M.; Yang, L.; Manning, A. M.; Firestein, G. S. c-Jun N-terminal kinase is required for metalloproteinase expression and joint destruction in inflammatory arthritis. J. Clin. Invest. 108:73–81; 2001. [48] Wu, W.; Muchir, A.; Shan, J.; Bonne, G.; Worman, H. J. Mitogen-activated protein kinase inhibitors improve heart function and prevent fibrosis in cardiomyopathy caused by mutation in lamin A/C gene. Circulation 123:53–61; 2011. [49] Koller, M.; Hayer, S.; Redlich, K.; Ricci, R.; David, J. P.; Steiner, G.; Smolen, J. S.; Wagner, E. F.; Schett, G. JNK1 is not essential for TNF-mediated joint disease. Arthritis Res. Ther. 7:R166–R173; 2005. [50] Tong, C.; Yin, Z.; Song, Z.; Dockendorff, A.; Huang, C.; Mariadason, J.; Flavell, R. A.; Davis, R. J.; Augenlicht, L. H.; Yang, W. c-Jun NH2-terminal kinase 1 plays a critical role in intestinal homeostasis and tumor suppression. Am. J. Pathol. 171:297–303; 2007. [51] She, Q. B.; Chen, N.; Bode, A. M.; Flavell, R. A.; Dong, Z. Deficiency of c-Jun-NH (2)-terminal kinase-1 in mice enhances skin tumor development by 12-Otetradecanoylphorbol-13-acetate. Cancer Res. 62:1343–1348; 2002. [52] Shen, H. M.; Liu, Z. G. JNK signaling pathway is a key modulator in cell death mediated by reactive oxygen and nitrogen species. Free Radic. Biol. Med. 40:928–939; 2006. [53] Gu, Y.; Xu, Y. C.; Wu, R. F.; Souza, R. F.; Nwariaku, F. E.; Terada, L. S. TNFalpha activates c-Jun amino terminal kinase through p47(phox). Exp. Cell Res. 272:62–74; 2002. [54] Xu, Y. C.; Wu, R. F.; Gu, Y.; Yang, Y. S.; Yang, M. C.; Nwariaku, F. E.; Terada, L. S. Involvement of TRAF4 in oxidative activation of c-Jun N-terminal kinase. J. Biol. Chem. 277:28051–28057; 2002. [55] Saitoh, M.; Nishitoh, H.; Fujii, M.; Takeda, K.; Tobiume, K.; Sawada, Y.; Kawabata, M.; Miyazono, K.; Ichijo, H. Mammalian thioredoxin is a direct inhibitor of apoptosis signal‐regulating kinase (ASK) 1. EMBO J. 17:2596–2606; 1998. [56] Min, W.; Lin, Y.; Tang, S.; Yu, L.; Zhang, H.; Wan, T.; Luhn, T.; Fu, H.; Chen, H. AIP1 recruits phosphatase PP2A to ASK1 in tumor necrosis factor–induced ASK1-JNK activation. Circ. Res. 102:840–848; 2008. [57] Yoshizumi, M.; Abe, J.; Haendeler, J.; Huang, Q.; Berk, B. C. Src and Cas mediate JNK activation but not ERK1/2 and p38 kinases by reactive oxygen species. J. Biol. Chem. 275:11706–11712; 2000. [58] Bogoyevitch, M. A.; Kobe, B. Uses for JNK: the many and varied substrates of the c-Jun N-terminal kinases. Microbiol. Mol. Biol. Rev. 70:1061–1095; 2006. [59] Kim, Y. S.; Morgan, M. J.; Choksi, S.; Liu, Z. G. TNF-induced activation of the Nox1 NADPH oxidase and its role in the induction of necrotic cell death. Mol. Cell 26:675–687; 2007. [60] Streeter, J.; Schickling, B. M.; Jiang, S.; Stanic, B.; Thiel, W. H.; Gakhar, L.; Houtman, J. C.; Miller Jr. F. J. Phosphorylation of Nox1 regulates association with NoxA1 activation domain. Circ. Res. 115:911–918; 2014. [61] Berk, B. C. Redox signals that regulate the vascular response to injury. Thromb. Haemost. 82:810–817; 1999. [62] Rao, G. N.; Berk, B. C. Active oxygen species stimulate vascular smooth muscle cell growth and proto-oncogene expression. Circ. Res. 70:593–599; 1992. [63] Pham-Huy, L. A.; He, H.; Pham-Huy, C. Free radicals, antioxidants in disease and health. Int. J. Biomed. Sci. 4:89–96; 2008. [64] Dietary supplementation with n-3 polyunsaturated fatty acids and vitamin E after myocardial infarction: results of the GISSI-Prevenzione trial. Gruppo Italiano per lo Studio della Sopravvivenza nell'Infarto miocardico. Lancet 354:447-455; 1999. [65] Yusuf, S.; Dagenais, G.; Pogue, J.; Bosch, J.; Sleight, P. Vitamin E supplementation and cardiovascular events in high-risk patients. The Heart Outcomes Prevention Evaluation Study Investigators. N. Engl. J. Med. 342:154–160; 2000. [66] MRC/BHF Heart Protection Study of antioxidant vitamin supplementation in 20,536 high-risk individuals: a randomised placebo-controlled trial. Lancet 360:23-33; 2002.