ORIGINAL RESEARCH ARTICLE

Journal of

Cyclooxygenase-2 Regulates NLRP3 Inflammasome-Derived IL-1b Production

Cellular Physiology

KUO-FENG HUA,1* JU-CHING CHOU,1 SHUK-MAN KA,2 YU-LING TASI,3 ANN CHEN,4 SHIH-HSIUNG WU,5 HSIAO-WEN CHIU,1 WEI-TING WONG,1 YIH-FUH WANG,6 CHANGE-LING TSAI,6 CHEN-LUNG HO,7 AND CHENG-HSIU LIN1 1

Department of Biotechnology and Animal Science, National Ilan University, Ilan, Taiwan

2

Graduate Institute of Aerospace and Undersea Medicine, National Defense Medical Center, Taipei, Taiwan

3

Graduate Institute of Life Science, National Defense Medical Center, Taipei, Taiwan

4

Department of Pathology, Tri-Service General Hospital, National Defense Medical Center, Taipei, Taiwan

5

Institute of Biological Chemistry, Academia Sinica, Taipei, Taiwan

6

Graduate Institute of Electrical Engineering and Computer Science, National Penghu University of Science and Technology, Penghu, Taiwan

7

Division of Wood Cellulose, Taiwan Forestry Research Institute, Taipei, Taiwan

The NLR family, pyrin domain-containing 3 (NLRP3) inflammasome is a reactive oxygen species-sensitive multiprotein complex that regulates IL-1b maturation via caspase-1. It also plays an important role in the pathogenesis of inflammation-related disease. Cyclooxygenase-2 (COX-2) is induced by inflammatory stimuli and contributes to the pathogenesis of inflammation-related diseases. However, there is currently little known about the relationship between COX-2 and the NLRP3 inflammasome. Here, we describe a novel role for COX-2 in regulating the activation of the NLRP3 inflammasome. NLRP3 inflammasome-derived IL-1b secretion and pyroptosis in macrophages were reduced by pharmaceutical inhibition or genetic knockdown of COX-2. COX-2 catalyzes the synthesis of prostaglandin E2 and increases IL-1b secretion. Conversely, pharmaceutical inhibition or genetic knockdown of prostaglandin E2 receptor 3 reduced IL-1b secretion. The underlying mechanisms for the COX-2-mediated increase in NLRP3 inflammasome activation were determined to be the following: (1) enhancement of lipopolysaccharide-induced proIL-1b and NLRP3 expression by increasing NF-kB activation and (2) enhancement of the caspase-1 activation by increasing damaged mitochondria, mitochondrial reactive oxygen species production and release of mitochondrial DNA into cytosol. Furthermore, inhibition of COX-2 in mice in vivo with celecoxib reduced serum levels of IL-1b and caspase-1 activity in the spleen and liver in response to lipopolysaccharide (LPS) challenge. These findings provide new insights into how COX-2 regulates the activation of the NLRP3 inflammasome and suggest that it may be a new potential therapeutic target in NLRP3 inflammasome-related diseases. J. Cell. Physiol. 230: 863–874, 2015. © 2014 Wiley Periodicals, Inc.

COX-2 is the most thoroughly studied and best-understood mammalian dioxygenase. It is inhibited by nonsteroidal antiinflammatory drugs and plays a primary therapeutic role in the treatment of inflammation. COX-2 has also been implicated in the development of diseases, including tumorigenesis (Khan et al., 2011), diabetes mellitus (Bagi et al., 2006), diabetic nephropathy (Nasrallah et al., 2009; Cheng et al., 2011; Quilley et al., 2011), osteoarthritis (Zweers et al., 2011), and atherosclerosis (Cuccurullo et al., 2007). COX-2-targeted therapy is commonly used to treat inflammation. For example, the COX-2-selective inhibitor celecoxib is used clinically to treat rheumatoid arthritis, osteoarthritis, ankylosing spondylitis (McCormack, 2011), atherosclerosis (Jacob et al., 2008), cancer (Khan Z et al., 2011), and Alzheimer’s disease (Pasinetti and Aisen, 1998). IL-1b is synthesized as an inactive immature form (precursor of IL-1b, proIL-1b) in macrophages in response to LPS (Hsu and Wen, 2002). IL-1b release is controlled by inflammasomes, which are caspase-1-containing multiprotein complexes (Miller et al., 1997; Latz et al., 2013). The best-characterized inflammasome is the NLRP3 inflammasome. The NLRP3 inflammasome controls caspase-1 activity and IL-1b release in the innate immune system (Jin and Flavell, 2010). The NLRP3 inflammasome also controls the development of disease, including pathogen infection (Kanneganti et al., 2006), type 2 © 2 0 1 4 W I L E Y P E R I O D I C A L S , I N C .

diabetes mellitus (Schroder et al., 2010), atherosclerosis (Duewell et al., 2010), obesity (Vandanmagsar et al., 2011), silicosis (Hornung et al., 2008), Alzheimer’s disease (Halle et al., 2008), gout (Martinon et al., 2006), kidney disease (Anders and Muruve, 2011), and cancer (Okamoto et al., 2010). Caspase-1 not only controls the IL-1b release but also regulates pyroptosis, which is a form of programmed cell death that involves rapid plasma-membrane rupture (Miao

Contract grant sponsor: Ministry of Science and Technology, Taiwan; Contract grant numbers: NSC 102-2628-B-197-001-MY3, NSC 103-2923-B-197-001-MY3. *Correspondence to: Kuo-Feng Hua, Department of Biotechnology and Animal Science, National Ilan University, No. 1, Sec. 1, Shen-Lung road, Ilan, 260, Taiwan. E-mail: [email protected] Manuscript Received: 20 February 2014 Manuscript Accepted: 5 September 2014 Accepted manuscript online in Wiley Online Library (wileyonlinelibrary.com): 7 October 2014. DOI: 10.1002/jcp.24815

863

864

HUA ET AL.

et al., 2011). Pyroptosis is triggered by caspase-1 after its activation by various inflammasomes. The process results in the formation of caspase-1-mediated plasma-membrane pores that cause water influx, cell swelling and eventually plasmamembrane rupture and release of intracellular contents (Bergsbaken et al., 2009). COX-2 expression can be induced by inflammatory mediators, such as IL-1b and TNF-a (Dinarello, 2009; Zweers et al., 2011). COX-2 also plays important roles in NLRP3 inflammasome-related diseases (Pasinetti and Aisen, 1998; Bagi et al., 2006; Cuccurullo et al., 2007; Nasrallah et al., 2009; Cheng et al., 2011; Quilley et al., 2011; Zweers et al., 2011). However, the relationship between COX-2 and the NLRP3 inflammasome has not been examined. In this study, we identified a novel role for COX-2 in NLRP3 inflammasomederived IL-1b secretion and pyroptosis in macrophages.

were measured by ELISA. For caspase-1 activation, cells were incubated for 5.5 h with or without 1 mg/ml of LPS and then with or without the addition of ATP (5 mM) for the last 30 min. The levels of activated caspase-1 (p10) and pro-caspase-1 (p45) in the cells were measured by western blot. Western blot The protein expression levels of activated caspase-1 (p10), pro-caspase-1 (p45), NLRP3, proIL-1b, and COX-2 were analyzed by western blot (Liao et al., 2013). The results were quantified by densitometric analysis using ImageJ software. The densitometry fold change of each group was calculated by comparing the results with the control group. The band density is normalized to actin or p45 before fold change is calculated. ROS detection

Materials and Methods Reagents LPS (from Escherichia coli 0111:B4) and mouse antibodies against actin were purchased from Sigma (St. Louis, MO). The following reagents were obtained from Santa Cruz Biotechnology (Santa Cruz, CA): COX-2 shRNA lentiviral particles (sc-29278-V), control shRNA lentiviral particles (sc-108080), EP3 shRNA plasmid (sc-35315-sh), control shRNA plasmid (sc-108060), prostaglandin E2, the prostaglandin E2 receptor 1 antagonist SC19220, the prostaglandin E2 receptor 3 antagonist L-798106, rabbit antibodies against mouse IL-1b or caspase-1, and horseradish peroxidase-conjugated second antibodies. The prostaglandin E2 receptor 2 antagonist AH6809 and the prostaglandin E2 receptor 4 antagonist AH23848 were purchased from Cayman Chemicals (Ann Arbor, MI). All the inflammasome inducers, pNiFty2-SEAP, and LyoVecTM transfection agent were purchased from InvivoGen (San Diego, CA). IL-1b, IL-6, and TNFa ELISA kits were purchased from R&D Systems (Minneapolis, MN). Mouse anti-mouse NLRP3 antibody was purchased from Enzo Life Sciences Inc. (Exeter, Farmingdale, NY). The COX-2 inhibitor SC-791 and the COX-1 inhibitor FR 122047 were purchased from Calbiochem (La Jolla, CA). Cell culture Murine macrophage J774A.1 cells were obtained from the American Type Culture Collection (Rockville, MD). The cells were propagated in RPMI-1640 medium (Gibco Laboratories, Grand Island, NY) supplemented with 10% heat-inactivated fetal calf serum (FCS) (Biological Industries Ltd, Kibbutz Beit Haemek, Israel) and 2 mM l-glutamine (Life Technologies, Carlsbad, CA) at 37°C in a 5% CO2 incubator. Stable COX-2 knockdown J774A.1 cells were produced by infection with COX-2 shRNA lentiviral particles. Stable EP3 knockdown J774A.1 cells were produced by transfection with EP3 shRNA plasmid. Peritoneal macrophages were collected by intraperitoneal injection of 4% sterile thioglycollate medium (2 ml). Three days after injection the mice were sacrificed by cervical dislocation and the macrophages were harvested. The peritoneal macrophages were grown in RPMI-1640 medium supplemented with 2 mM l-glutamine, 100 U/ml penicillin, 100 mg/ml streptomycin, 2.5 mg/ml amphotericin B, and 10% FCS. The cells were plated in 24-well culture plates at a density of 5
105 cells/ml and allowed to adhere at 37°C. IL-1b secretion and caspase-1 activation To examine IL-1b secretion cells were incubated for 5.5 h with or without LPS (1 mg/ml), then with or without the addition of ATP (5 mM, 0.5 h), nigericin (10 mM, 0.5 h), SiO2 (200 mg/ml, 24 h), CPPD crystals (100 mg/ml, 24 h), FLA-ST (100 mg/ml, 24 h), or MDP (100 ng/ml, 24 h). The levels of IL-1b in the culture medium JOURNAL OF CELLULAR PHYSIOLOGY

General ROS production was measured by detecting the fluorescence intensity of the 20 ,70 -dichlorofluorescein, which is the oxidation product of 20 ,70 -dichlorofluorescein diacetate (Molecular Probes, Eugene, OR). For LPS-induced general ROS generation, cells were incubated with or without SC-791 (10 mM), PGE2 (10 nM), or NAC (10 mM) for 30 min and were then incubated with 20 ,70 -dichlorofluorescein diacetate (2 mM) for an additional 30 min. The cells were then stimulated with LPS (1 mg/ml) for the indicated time. To examine ATP-induced general ROS generation, cells were incubated with LPS (1 mg/ml) for 5.5 h and then incubated with or without SC-791 (10 mM) or NAC (10 mM) for 30 min before incubation with 20 ,70 dichlorofluorescein diacetate (2 mM) for additional 30 min. The cells were then stimulated with ATP (5 mM) for the indicated time. The fluorescence intensity of 20 ,70 -dichlorofluorescein was detected at an excitation wavelength of 485 nm and an emission wavelength of 530 nm on a microplate absorbance reader (Bio-Rad Laboratories, Inc). To evaluate mitochondrial ROS generation the cells were incubated with LPS (1 mg/ml) for 5.5 h and were then incubated with or without SC-791 (10 mM) for 30 min before stimulation with ATP (5 mM) for 30 min. The cells were then stained with 5 mM MitoSOX (Invitrogen, Carlsbad, CA) for 20 min. The fluorescence intensity of MitoSOX was detected at an excitation wavelength of 514 nm and an emission wavelength of 585 nm on a microplate absorbance reader. The data are presented as relative fluorescence intensity fold change of each group, which was calculated by comparison to the control group. Flow cytometry To measure the inner transmembrane potential and mitochondrial mass the cells were incubated for 5.5 h with or without LPS (1 mg/ml) in the presence or absence of PGE2 (10 nM). The cells were then incubated with or without SC-791 (10 mM) for 30 min. The cells were then stained for 15 min at 37°C with 25 nM MitoTracker Deep Red and MitoTracker Green, respectively (Invitrogen, Carlsbad, CA), followed by 30 min of ATP treatment. The cells were washed with FCS, treated with trypsin and resuspended in PBS containing 1% heat-inactivated FBS. The data were acquired with flow cytometry. Pyroptosis detection sh-COX-2 or sh-SC cells were incubated for 5.5 h with or without LPS (1 mg/ml) and were then incubated with or without ATP (5 mM) for an additional 30 min. To determine LDH release, culture supernatants were evaluated for the presence of the cytoplasmic enzyme LDH using the CytoTox 961 Non-radioactive Cytotoxicity Assay kit according to the manufacturer’s instructions (Promega, Madison, WI). The percentage cytotoxicity was calculated as 100
(experimental LDH-spontaneous LDH)/(maximum LDH

COX-2 REGULATES NLRP3 INFLAMMASOME release-spontaneous LDH). The AlamarBlue1 assay kit was used to measure proliferation according to the protocol described by the manufacturer (AbD Serotec Ltd). Cell size was determined by drawing circles around representative cells from 20 fields and calculating area using ImageJ software. PI (40 mg/ml) was added into cell cultures after treatment to determine PI uptake by cells. The PI fluorescence intensity was detected at an excitation wavelength of 536 nm and an emission wavelength of 617 nm on a microplate absorbance reader. PGE2 measurement PGE2 measurement was conducted using a PGE2 assay kit purchased from R&D Systems (catalog number: KGE004B). This assay is based on the forward sequential competitive binding technique in which PGE2 present in a sample competes with horseradish peroxidase (HRP)-labeled PGE2 for a limited number of binding sites on a mouse monoclonal antibody (R&D Systems). Conditioned media were collected from sh-SC cells and sh-COX-2 cells stimulated with LPS (1 mg/ml) for 24 h. Then, 150 ml diluted conditioned medium or serum from mice and 50 ml primary antibody solution were added into each well and incubated for 1 h at room temperature. Each well was treated with 50 ml of PGE2 conjugate and was incubated for additional 2 h at room temperature. After washing, 200 ml of substrate solution was added to each well and incubated for 30 min at room temperature (protected from light). Each well was then treated with 100 ml of stop solution and the optical density was determined using a microplate reader set to 450 nm. The assay kit is specific for PGE2 because the cross-reactivity to other prostaglandins is very low. The minimum detectable dose of PGE2 by this assay kit ranged from 16.0 to 41.4 pg/ml (according to the datasheet). NF-kB reporter assay J-Blue cells are J774A.1 macrophages stably expressing the gene for secreted embryonic alkaline phosphatase inducible by NF-kB (pNiFty2-SEAP). These cells were seeded in 24-well plates at a density of 1
105 cells/ml (0.5 ml) and grown overnight in a 5% CO2 incubator at 37°C. The cells were then pretreated with vehicle or SC-791 for 30 min and then LPS (1 mg/ml) was added and incubated for 24 h. The medium was then harvested and 20 ml aliquots were mixed with 200 ml of QUANTI-BlueTM medium (InvivoGen) in 96-well plates and incubated at 37°C for 15 min. The secreted embryonic alkaline phosphatase activity was assessed by measuring the optical density at 655 nm using an ELISA reader. Quantitative real-time PCR analysis RNA from macrophages was reverse transcribed and quantitative PCR analysis was performed using the Applied Biosystems1 StepOneTM Real-Time PCR System. All gene expression data are presented as relative expression normalized to HPRT1. The primers used were the following: mouse IL-1b, forward, 30 -CTGCAGCTGGAGAGTGTGG-30 , reverse, 50 -GGGGAACTCTGCAGACTCAA-30 ; mouse HPRT1, forward, 50 -CTGGTGAAAAGGACCTCTCG-30 , reverse, 50 TGAAGTACTCATTATAGTCAAGGGCA-30 (30). The detection of mitochondrial DNA (mtDNA) in the cytosol was performed as described (Nakahira et al., 2011). Briefly, 1
107 shSC and sh-COX-2 cells were homogenized in protease inhibitor containing buffer (100 mM pH 7.4 Tricine-NaOH, 0.25 M sucrose, 1 mM EDTA), and then centrifuged for 10 min at 700xg at 4°C. The protein concentration and volume of the supernatant was normalized and was then centrifuged for 30 min at 10,000g at 4°C. The resulting pellets were identified as the mitochondrial fraction, and the supernatants were identified as the cytosolic fraction. DNA was isolated from 200 ml of the cytosolic fraction. The mtDNA copy number was measured by quantitative PCR and JOURNAL OF CELLULAR PHYSIOLOGY

normalized to nuclear DNA (encoding 18 S ribosomal RNA) levels using a ratio of cytochrome c oxidase I DNA to nuclear DNA. The results were expressed as fold changes between the sample and the control group. The quality of the mitochondrial fraction and cytosolic fraction was monitored by detecting the protein expression level of voltage-dependent anion channel (VDAC, a marker protein for mitochondria). We found that VDAC can only be detected in the mitochondrial fraction and is not present in the cytosolic fraction (data not shown). These results indicate that the cytosolic fraction is free from mitochondria. The primers used were the following: mouse 18 S, forward, 50 -TAGAGGGACAAGTGGCGTTC-30 , reverse, 50 -CGCTGAGCCAGTCAGTGT-30 ; mouse cytochrome c oxidase I, forward, 50 -GCCCCAGATATAGCATTCCC-30 , reverse, 50 -GTTCATCCTGTTCCTGCTCC-30 . Animal experiments The experiments were performed on 8-week-old female BABL/c mice purchased from the National Laboratory Animal Breeding and Research Center (Taipei, Taiwan). The celecoxib was purchased from BioVision (Mountain View, CA, USA) and it was dissolved in buffer A (mixture of dimethylformamide, Tween-80, and saline). The mice were randomized into the following four groups: Group I: control, buffer A (100 ml) intraperitoneal injection once daily for 3 days and saline (100 ml) intraperitoneal injection once 1 h after last buffer A injection, n ¼ 10; Group II: LPS, buffer A (100 ml) intraperitoneal injection once daily for 3 days and LPS (3 mg/g body weight) intraperitoneal injection once 1 h after last buffer A injection, n ¼ 8; Group III: celecoxib þ LPS, celecoxib (10 mg/g body weight/day) intraperitoneal injection once daily for 3 days and LPS (3 mg/g body weight) intraperitoneal injection once 1 h after last celecoxib injection, n ¼ 8; Group IV: celecoxib, celecoxib (10 mg/g body weight/day) intraperitoneal injection once daily for 3 days and saline (100 ml) intraperitoneal injection once 1 h after last celecoxib injection, n ¼ 4. After treatment, serum and organs were collected for further analysis at 4 h and 24 h after LPS injection, respectively. These animal experiments were approved by the Institutional Animal Care and Use Committee of the National Defense Medical Center, Taiwan. All studies were performed according to the ethical rules in the NIH Guide for the Care and Use of Laboratory Animals. Tissue caspase-1 activity assay To measure caspase-1 activity in tissues we followed the protocol recommended for the fluorometric CaspACE1 assay system (Promega Corporation, Madison, WI). Briefly, 32 ml of ICE-like enzyme assay buffer, 2 ml of dimethyl sulfoxide, and 10 ml of 100 mM dithiothreitol were added to 75 mg of protein from tissue extract. The volume was adjusted with water to 98 ml. For each test sample, a negative control containing all of the above components plus 2 ml of 2.5 mM ICE inhibitor (Ac-YVAD-CHO) in a total volume of 98 ml and a blank control (32 ml of ICE-like enzyme assay buffer, 2 ml of sulfoxide, 10 ml of 100 mM dithiothreitol, and to 98 ml of water) were included. The mixtures were incubated at 30°C for 30 min and then 2 ml of 2.5 mM ICE substrate (Ac-YVAD-AMC) was added and the sample was incubated for 60 min at 30°C. The fluorescence intensity was measured at an excitation wavelength of 360 nm and an emission wavelength of 460 nm. The calculation of the relative fluorescence units and the caspase-1 activity for each sample and the construction of the standard curve and AMC calibration curves were performed as described in the Promega Technical Bulletin. Statistical analysis All values are given as the mean  SD. The data analysis was performed by one-way ANOVA followed by a Scheffe test.

865

866

HUA ET AL.

Results COX-2 positively regulates NLRP3 inflammasome activation

To provide direct evidence of COX-2 involvement in the activation of the NLRP3 inflammasome we stably transfected J774A.1 macrophages with shRNA plasmids targeting COX-2 (sh-COX-2). We found that COX-2 expression and PGE2 secretion in these cells was significantly lower than in cells stably transfected with a control shRNA plasmid encoding a scrambled shRNA sequence (sh-SC) (Supplemental Fig. S1A). We found that IL-1b secretion after the treatment of NLRP3 inflammasome activators, including ATP, nigericin, calcium pyrophosphate dihydrate, and SiO2 nanoparticles, was significantly lower in sh-COX-2 cells than in sh-SC cells (Fig. 1A). These effects were confirmed in murine primary peritoneal macrophages using the COX-2 specific inhibitor SC791 (Fig. 1B). Caspase-1 activation induced by LPS and ATP were significantly lower in sh-COX-2 cells than in sh-SC cells (Supplemental Fig. S1B). However, TNF-a secretion was not affected (Supplemental Fig. S1C). The reduced IL-1b secretion in sh-COX-2 cells might be caused by impaired NLRP3 inflammasome priming signals or activation signals. We incubated LPS-primed macrophages with the COX-2 inhibitor (SC-791) or COX-1 inhibitor (FR 122047) for 30 min before ATP stimulation. We found that the COX-2 inhibitor reduced IL-1b secretion and caspase-1 activation (Fig. 2A), while the COX-1 inhibitor had no significant effect (Supplemental Fig. S2A). Furthermore, neither inhibitor significantly affected IL-6 secretion independent of the NLRP3 inflammasome (Supplemental Fig. S2B). These results indicated that COX-2 regulates the ATP-mediated activation signal for the NLRP3 inflammasome. In addition, NLRP3 inflammasome activation is tightly controlled by an LPS-mediated priming signal that requires NLRP3 and proIL-1b protein synthesis (Dinarello, 2009; Bauernfeind et al., 2011). We found that the protein expression levels of NLRP3 and proIL-1b in sh-COX-2 cells were lower than in sh-SC cells (Fig. 2B). Treatment with the COX-2 inhibitor also caused a significant reduction in the protein expression levels of NLRP3 and proIL-1b in LPSstimulated macrophages (Supplemental Fig. S2C). We also found the COX-2 inhibitor reduced both NF-kB activation and IL-1b mRNA expression in LPS-activated J774A.1 macrophages (Fig. 2C). These results indicated that COX-2 regulates the LPSmediated priming signal for the NLRP3 inflammasome. We found that COX-2 is not selective for NLRP3-dependent inflammasome stimuli. The knockdown of COX-2 reduced IL1b secretion triggered by FLA-ST (flagellin from S. typhimurium) or muramyl dipeptide, which are dependent on the NLRP1 and NLRC4 inflammasomes (Latz et al., 2013), respectively. These processes are also independent of NLRP3 (Fig. 2D). IL-1b secretion triggered by FLA-ST and muramyl dipeptide was also reduced by the COX-2 inhibitor SC-791 in both J774A.1 macrophages and murine primary peritoneal macrophages (Supplemental Fig. S3A). However, the levels of TNF-a and IL-6 were not affected (Supplemental Fig. S3, B and C). Effect of COX-2 on ROS generation and mitochondrial homeostasis

ROS play an important role in NLRP3 inflammasome activation (Tschopp and Schroder, 2010; Bauernfeind et al., 2011). Because COX-2 is able to generate ROS (Fraser, 2011; Hsieh et al., 2011), we investigated whether COX-2 regulates ROS production in response to LPS and ATP stimulation. We found that LPS-induced ROS production was not affected by the COX-2 inhibitor (Fig. 3A), which indicates that COX-2 is not required for LPS-mediated ROS generation. ATP induces ROS production in LPS-primed macrophages (Moore and MacKenzie, 2009). JOURNAL OF CELLULAR PHYSIOLOGY

Figure 1. Knockdown of COX-2 attenuates NLRP3 inflammasome activation. (A) sh-COX-2 or sh-SC cells were incubated for 5.5 h with or without LPS. The cells were then treated with or without ATP, nigericin, nano SiO2, or CPPD crystals. The levels of IL-1b in the culture medium were measured by ELISA. (B) Murine primary peritoneal macrophages were incubated for 5.5 h with or without LPS. The cells were then treated for 30 min with or without COX-2 inhibitor (SC-791) in the continued presence or absence of LPS. The cells were then treated with or without ATP, nigericin, nano SiO2, or CPPD crystals. The levels of IL-1b in the culture medium were measured by ELISA. The data are expressed as the mean  SD for three separate experiments. ** and *** indicates a significant difference at the level of P < 0.01 and P < 0.001, respectively.

We found this increase was significantly reduced by the COX-2 inhibitor (Fig. 3B). These data suggest that COX-2 plays an important role in ATP-mediated ROS production. In addition, NLRP3 inflammasome activity is positively regulated by ROS derived from mitochondria (Zhou et al., 2011). We found that LPS with ATP treatment increased mitochondrial ROS level in sh-SC cells, but not in sh-COX-2 cells (Fig. 3C). Treatment with the COX-2 inhibitor also reduced mitochondrial ROS levels in LPS plus ATP activated J774A.1 macrophages and murine primary peritoneal macrophages (Supplemental Fig. S4A). We further investigated the functional mitochondrial pool in cells

COX-2 REGULATES NLRP3 INFLAMMASOME

Figure 2. Inhibition of COX-2 attenuates NLRP3 inflammasome activation. (A) J774A.1 macrophages were incubated for 5.5 h with or without LPS. The cells were then treated for 30 min with or without COX-2 inhibitor (SC-791) in the continued presence or absence of LPS. The cells were then treated for 30 min with or without ATP. The levels of IL-1b in the culture medium and activated caspase-1 (p10) in the cells were measured by ELISA and western blot, respectively. (B) sh-COX-2 or sh-SC cells were incubated for 5.5 h with or without LPS. The cells were then treated for 30 min with or without ATP. The protein expression levels of NLRP3 and proIL-1b in the cells were measured by western blot. (C) J774A.1 macrophages were incubated for 30 min with or without COX-2 inhibitor (SC-791), then for 24 h (NF-kB activation) or 2 h (IL-1b mRNA) with or without LPS in the continued presence or absence of SC-791. NF-kB activation and IL-1b mRNA expression were measured by NF-kB reporter assay and RT-PCR analysis, respectively. (D) sh-COX-2 or sh-SC cells were incubated for 5.5 h with or without LPS. The cells were then treated for 24 h with or without NLRP1 inflammasome activator FLA-ST or NLRC4 inflammasome activator MDP. The levels of IL-1b in the culture medium were measured by ELISA. The data are expressed as the mean  SD for three separate experiments. The western blot results are representative of three different experiments and the histograms are presented as the change in the ratio relative to actin or p45 compared to control group. *, ** and *** indicates a significant difference at the level of P < 0.05, P < 0.01, and P < 0.001, respectively.

JOURNAL OF CELLULAR PHYSIOLOGY

867

868

HUA ET AL.

Figure 3. Effect of COX-2 on the generation of general ROS and mitochondrial homeostasis. (A) J774A.1 macrophages were incubated for 30 min with or without SC-791 or NAC. The cells were then treated for 0–80 min with or without LPS in the continued presence or absence of SC-791 or NAC. The levels of ROS generation in the cells were measured by 20 ,70 -dichlorofluorescein diacetate. (B) J774A.1 macrophages were incubated for 5.5 h with or without LPS. The cells were then treated for 30 min with or without SC-791 or NAC in the continued presence or absence of LPS. The cells were then treated for 0–80 min with or without ATP. The levels of ROS in the cells were measured by 20 ,70 -dichlorofluorescein diacetate. (C) sh-COX-2 or sh-SC cells were incubated for 5.5 h with or without LPS. The cells were then treated for 30 min with or without ATP. The levels of mitochondrial ROS in the cells were measured by MitoSOX. (D) sh-COX-2 or sh-SC cells were incubated for 5.5 h with or without LPS. The cells were then treated for 15 min with MitoTracker Deep Red and MitoTracker Green. The cells were then treated for 30 min with or without ATP. The mitochondrial inner transmembrane potential and mitochondrial mass were measured by flow cytometry. (E) sh-COX-2 or sh-SC cells were incubated for 5.5 h with or without LPS. The cells were then treated for 30 min with or without ATP. The mitochondrial DNA release into cytosol was measured by detection of cytochrome c oxidase I DNA in cytosol. The data are expressed as the mean  SD for three separate experiments. * and ** indicates a significant difference at the level of P < 0.05 and P < 0.01, respectively.

JOURNAL OF CELLULAR PHYSIOLOGY

COX-2 REGULATES NLRP3 INFLAMMASOME with MitoTracker Deep Red, which is a fluorescent probe that stains mitochondria in live cells and accumulates in active mitochondria. To measure the total mitochondrial pool we counterstained cells with MitoTracker Green, which is a fluorescent probe that stains mitochondria regardless of mitochondrial membrane potential. As shown in Figure 3D, stimulation with LPS and ATP increased the percentage of cells with damaged mitochondria from 2.03 to 17.31% of the total population of sh-SC cells. This stimulation increased damaged mitochondria from 1.95 to 7.06% in sh-COX-2 cells. The effect of COX-2 on mitochondrial function was confirmed by inhibition of COX-2 using the specific inhibitor SC-791. The combined stimulation with LPS and ATP increased the percentage of cells with damaged mitochondria from 3.18 to 18.57% of the total population of J774A.1 macrophages. However, the COX-2 inhibitor SC-791 reduced damaged mitochondria to 10.32%. The treatment with PGE2 increased damaged mitochondria to 24.04% (Supplemental Fig. S4B). Damaged mitochondria and mitochondrial ROS induce the translocation of mitochondrial DNA (mt-DNA) into the cytosol, and cytosolic mt-DNA is a coactivator of the NLRP3 inflammasome (Zhou et al., 2011). Therefore, we examined whether the translocation of mt-DNA into cytosol was decreased in sh-COX-2 cells upon LPS and ATP stimulation. We found the stimulation increased mt-DNA release into cytosol in sh-SC cells, but not in sh-COX-2 cells (Fig. 3E). These results reflected less mitochondrial injury in COX-2 inhibition/knockdown cells than in control cells and suggest that the disruption of COX-2 protects cells from LPS and ATP stimulation. Inhibition of PGE2 receptor reduces NLRP3 inflammasome activation

PGE2 receptor 3 (EP3) and PGE2 receptor 4 (EP4) represent highaffinity receptors, whereas PGE2 receptor 1 (EP1) and PGE2 receptor 2 (EP2) require significantly higher concentrations of PGE2 for effective signaling (Kalinski, 2012). EP1 activates Ca2þ signaling and EP3 inhibits cAMP signaling (Kalinski, 2012), which both events promote the activation of NLRP3 inflammasome (Lee et al., 2012). Thus, we examined the role of EP1 and EP3 on the activation of NLRP3 inflammasome. We found that IL-1b secretion was reduced by treatment with both EP3 inhibitor (L-798106) and EP1 inhibitor (sc-19220) (Fig. 4A). However, L-798106 was approximately 3-fold more potent than sc-19220. As a result, caspase-1 activation was reduced by L-798106 but not by sc-19220 (Fig. 4B). We also found that neither sc-19220 nor L-798106 affected IL-6 secretion (Supplemental Fig. S5A). To confirm the role of EP3 in the activation of the NLRP3 inflammasome we investigated the IL-1b secretion and caspase-1 activation in cells stably transfected with shRNA plasmids targeting EP3 (sh-EP3) (Supplemental Fig. S5B). We found that IL-1b secretion and caspase-1 activation induced by LPS and ATP were significantly lower in sh-EP3 cells than in cells stably transfected with a control shRNA plasmid encoding a scrambled shRNA sequence (sh-SC) (Fig. 4C). We also found that IL-6 secretion was not significantly reduced (Supplemental Fig. S5C). In addition, LPS-induced protein expression levels of NLRP3 and proIL-1b were lower in sh-EP3 cells than in sh-SC cells (Fig. 4D). To further examine the effects of EP activation on LPS-induced signaling pathways we monitored the LPS-induced NF-kB activation in J774A.1 macrophages incubated with specific inhibitors against EP1, EP2, EP3, and EP4. We observed a slight decrease in the NF-kB activation in macrophages treated with the EP14 inhibitors (Fig. 4E). PGE2 increases NLRP3 inflammasome activation

Due to the importance of PGE2 receptor in NLRP3 inflammasome activation, we hypothesized that NLRP3 JOURNAL OF CELLULAR PHYSIOLOGY

inflammasome activation might be further increased by stimulation with exogenous PGE2. Thus, we treated cells with PGE2 before LPS priming and examined the effect on ATPinduced IL-1b secretion. The results indicate that PGE2 increased IL-1b secretion in a dose-dependent manner. Furthermore, this effect was reduced by the EP1 inhibitor sc19220 and blocked by the EP3 inhibitor L-798106 (Fig. 5A). Exogenous PGE2 stimulation was also able to restore LPS- and ATP-induced IL-1b secretion in cells pretreated with COX-2 inhibitor (Fig. 5B). PGE2 treatment also increased IL-1b secretion in mouse primary peritoneal macrophages (Supplemental Fig. S6A). Because PGE2 can induce COX-2 expression, we investigated whether the effect of exogenous PGE2 on NLRP3 inflammasome is due to more COX-2 induction induced by PGE2. We investigated the effect of PGE2 on caspase-1 activation and IL-1b secretion in sh-SC and sh-COX-2 cells. We found that exogenous PGE2 further increased LPS plus ATPinduced caspase-1 activation and IL-1b secretion in sh-SC cells. However, PGE2 increased caspase-1 activation and IL-1b secretion slightly but significantly in sh-COX-2 cells (Fig. 5C). Importantly, caspase-1 activation and IL-1b secretion in shCOX-2 cells were much lower than in sh-SC cells (Fig. 5C). In addition, exogenous PGE2 further increased LPS-induced expression levels of proIL-1b, but not NLRP3 in sh-SC cells. The expression levels of proIL-1b and NLRP3 in sh-COX-2 cells were much lower than in sh-SC cells (Fig. 5D). These results suggested that COX-2 plays an important role in PGE2-mediated effects on NLRP3 inflammasome activation. Thus, additional COX-2 expression induced by PGE2 may be responsible for this effect. Furthermore, we found that PGE2 induced ROS production and significantly increased LPS-induced ROS production in both J774A.1 and murine primary peritoneal macrophages (Supplemental Fig. S6B). PGE2 also increased mitochondrial ROS levels in LPS plus ATP activated murine primary peritoneal macrophages (Supplemental Fig. S6C). COX-2 positively regulates caspase-1 dependent pyroptosis

Pyroptosis is characterized by loss of membrane integrity, which results from caspase-1-dependent insertion of a pore into the membrane. This process leads to fluid influx, cell swelling, and lysis. To evaluate the effect of COX-2 inhibition on pyroptosis, sh-SC cells and sh-COX-2 cells were treated with LPS and ATP for 30 min and samples were assayed for LDH release. We found that LDH release induced by LPS and ATP was significantly lower in sh-COX-2 cells than in sh-SC cells (Fig. 6A, left panel). The inhibition of COX-2 also reduced LDH release in LPS and ATP activated murine primary peritoneal macrophages (Fig. 6A, right panel). To confirm cell death with an alternative assay we used the AlamarBlue1 cell viability assay. Consistent with the LDH release assay, the AlamarBlue1 analysis showed cell viability was significantly higher in sh-COX-2 cells than in sh-SC cells (Fig. 6B). The LDH and AlamarBlue1 assays suggest that exposure to LPS and ATP disrupts the plasma membrane and this effect was rescued by COX-2 inhibition. To determine whether this reduced plasma membrane integrity affected cell size we quantified cell size after exposure to LPS and ATP. Figure 6C shows an increase in cell size after exposure to LPS and ATP in sh-SC cells (mean of 165  16 mm2 for control cells compared with 206  20 mm2 for LPS and ATP exposed cells). The treatment with LPS and ATP did not increase cell size in shCOX-2 cells (mean of 164  15 mm2 for control cells compared with 166  14 mm2 for LPS and ATP exposed cells). We also examined cell uptake of the fluorescent membrane impermeable dye propidium iodide (PI) to quantitatively examine membrane damage in individual cells during LPS and ATP exposure (Fink and Cookson, 2006). The incubation of sh-SC cells with LPS and ATP resulted in an increase in PI uptake. However, LPS and ATP

869

870

HUA ET AL.

Figure 4. Inhibition of PGE2 receptors attenuates NLRP3 inflammasome activation. J774A.1 macrophages were incubated for 30 min with or without EP3 inhibitor (L-798106) or EP1 inhibitor (SC-19220). The cells were then treated for 5.5 h with or without LPS. The cells were then treated for 30 min with or without ATP. The levels of IL-1b in the culture medium (A) and activated caspase-1 (p10) in the cells (B) were measured by ELISA and western blot, respectively. (C) sh-SC and sh-EP3 cells were incubated for 5.5 h with or without LPS. The cells were then treated for 30 min with or without addition of ATP. The levels of IL-1b in the culture medium and activated caspase-1 (p10) in the cells were measured by ELISA and western blot, respectively. (D) sh-SC and sh-EP3 cells were incubated for 6 h with or without LPS. The protein expression levels of NLRP3 and proIL-1b in the cells were measured by western blot. (E) J774A.1 macrophages were incubated for 30 min with or without EP14 inhibitor. The cells were then treated for 24 h with or without LPS in the continued presence or absence of inhibitor. NF-kB activation was measured using a NF-kB reporter assay. The data are expressed as the mean  SD for three separate experiments. The western blot results are representative of three different experiments and the histograms are presented as the change in the ratio relative to p45 or actin compared to control group. * and ** indicates a significant difference at the level of P < 0.05 and P < 0.01, respectively.

JOURNAL OF CELLULAR PHYSIOLOGY

COX-2 REGULATES NLRP3 INFLAMMASOME

Figure 5. Effect of PGE2 on NLRP3 inflammasome activation. (A) J774A.1 macrophages were incubated for 30 min with or without EP3 inhibitor (L-798106) or EP1 inhibitor (SC-19220). The cells were then treated for 30 min with or without PGE2. The cells were then treated for 5.5 h with or without LPS and then with or without ATP for 30 min. The levels of IL-1b in the culture medium were measured by ELISA. (B) J774A.1 macrophages were incubated for 30 min with or without SC-791. The cells were then treated for 30 min with or without EP3 inhibitor (L-798106) or EP1 inhibitor (SC-19220). The cells were then treated for 30 min with or without PGE2 followed by 5.5 h incubation with or without PS. The cells were then treated for 30 min with or without ATP. The levels of IL-1b in the culture medium were measured by ELISA. (C) sh-SC and sh-COX-2 cells were incubated for 30 min with or without PGE2. The cells were then treated for 5.5 h with or without LPS. The cells were then treated for 30 min with or without ATP. The levels of IL-1b in the culture medium and activated caspase-1 (p10) in the cells were measured by ELISA and western blot, respectively. (D) sh-SC and sh-COX-2 cells were incubated for 30 min with or without PGE2. The cells were then treated for 6 h with or without LPS. The protein expression levels of NLRP3 and proIL-1b in the cells were measured by western blot. The data are expressed as the mean  SD for three separate experiments. The western blotting results are representative of three different experiments. * and ** indicates a significant difference at the level of P < 0.05 and P < 0.01, respectively. # indicates a significant difference at the level of P < 0.001 compared to LPS þ ATP group.

exposure did not induce an increase in PI uptake in sh-COX-2 cells (Fig. 6D). Inhibition of COX-2 suppresses NLRP3 inflammasome activation in LPS-injected mice

In this study, mice were left untreated or were injected with LPS with or without injection with COX-2 inhibitor celecoxib. We then collected serum samples 4 h after LPS injection to measure levels of PGE2 and IL-1b. We also collected the spleen and liver 24 h after injection to measure COX-2, NLRP3 levels and caspase-1 activity. As shown in Figure 7A, LPS injection resulted in a significant increase in COX-2 protein levels in the spleen and in serum PGE2 levels compared to the salineJOURNAL OF CELLULAR PHYSIOLOGY

injected controls. These effects were inhibited in the celecoxibpretreated mice. The LPS-injected mice also showed a significant increase in serum IL-1b protein levels (Fig. 7B), caspase-1 activity in the spleen and liver (Fig. 7C), and NLRP3 protein levels in the spleen and liver (Fig. 7D). These effects were significantly inhibited by pretreatment with celecoxib. In contrast, celecoxib had no significant effect on the LPS-induced increase in serum IL-6 protein levels (Fig. 7E). Discussion

COX-2 overexpression is clearly associated with the pathogenesis of many inflammation-related diseases (Pasinetti and Aisen, 1998; Redondo et al., 2011), but a causative link has

871

872

HUA ET AL.

Figure 6. Knockdown of COX-2 attenuates pyroptosis. sh-COX-2 or sh-SC cells were incubated for 5.5 h with or without LPS. The cells were then treated for 30 min with or without ATP. (A) LDH release was assayed by CytoTox 961 Non-radioactive Cytotoxicity Assay kit (For primary macrophages, the cells were incubated for 30 min with or without SC-791, then for 5.5 h with or without LPS, then with or without ATP for 30 min). (B) Cell viability was assayed by AlamarBlue1 assay kit. (C) Cell size was determined by drawing circles around representative cells from 20 fields and calculating area using ImageJ software. (D) The fluorescence intensity of PI staining was detected by a microplate absorbance reader. The data are expressed as the mean  SD for three separate experiments. *, **, and *** indicates a significant difference at the level of P < 0.05, P < 0.01, and P < 0.001, respectively.

not been established. The NLRP3 inflammasome was recently shown to be a promising therapeutic target for inflammatory disease treatment (Strowig et al., 2012). In the present study, we identified COX-2 as a novel regulator of the NLRP3 inflammasome and provided a possible explanation of how COX-2 plays an important role in NLRP3 inflammasomerelated disease. We found that COX-2 promotes NLRP3 inflammasome activation. This is the first report showing that JOURNAL OF CELLULAR PHYSIOLOGY

IL-1b secretion can be regulated by COX-2, which is a new idea that is different from the traditional relationship where COX-2 can be induced by IL-1b (Mifflin et al., 2002; Dinarello, 2009). The genetic knockdown or pharmacological inhibition of COX-2 attenuated both the priming and activation signals for the NLRP3 inflammasome. In the priming stage, inhibition of COX-2 reduced LPS-induced NF-kB activation. This priming is required for NLRP3 expression (Bauernfeind et al., 2009). The inhibition of COX-2 did not affect LPS-induced ROS generation, which is also required for NLRP3 induction in the priming step of the NLRP3 inflammasome (Bauernfeind et al., 2011). These results indicated that reduced NF-kB activation may be responsible for the decreased expression level of NLRP3 in COX-2 inhibited macrophages. Although COX-2 plays important role in the transcriptional regulation of NLRP3 expression, we cannot rule out the possibility that COX-2 might affect the ubiquitination and non-transcriptional regulation of NLRP3 that are part of the activation mechanism of the NLRP3 inflammasome (Juliana et al., 2012; Py et al., 2013). Interestingly, we found that LPS-induced NLRP3 expression was reduced by ATP. This result suggests there may be feedback regulation involved in the activation of the NLRP3 inflammasome. However, the detailed mechanism requires further investigation. In addition, LPS-induced proIL-1b expression was also reduced by ATP. It should be noted that caspase-1 activation induced by ATP cleaves proIL-1b into IL1b and may be responsible for the decreased expression level of proIL-1b after ATP treatment. Similar results were observed in a previous study (Nakahira et al., 2011). ROS not only plays an important role in the priming step of NLRP3 inflammasome, but also regulates the activation step of the NLRP3 inflammasome (Tschopp and Schroder, 2010). The induction of COX-2 expression leads to PGE2 production and is usually accompanied by an increase in ROS levels (Fraser, 2011; Hsieh et al., 2011). Conversely, the inhibition of COX-2 decreases ROS generation in human promonocytic cells infected by C. pneumonia (Mouithys-Mickalad et al., 2004) and in myeloidderived suppressor cells and macrophages from tumor-bearing mice (Veltman et al., 2010). We also found that inhibition of COX-2 reduced mitochondrial ROS generation, which has an important role in NLRP3 inflammasome activation (Kepp et al., 2011; Zhou et al., 2011). The overproduction of mitochondrial ROS promotes mitochondrial permeability and facilitates the cytosolic release of mitochondrial DNA, which stimulates activation of the NLRP3 inflammasome (Kepp et al., 2011; Zhou et al., 2011). The inhibition of COX-2 reduced mitochondrial permeability and the cytosolic release of mitochondrial DNA. These data indicated that mitochondria are involved in COX-2-mediated activation of the NLRP3 inflammasome. It has been demonstrated that cAMP inhibits NLRP3 inflammasome activation by binding to NLRP3 directly and inhibiting inflammasome assembly. Additionally, downregulation of cAMP relieves this inhibition (Lee et al., 2012). The increase of cAMP synthesis by adenylate cyclase activators or the decrease of cAMP hydrolysis by phosphodiesterase 4 inhibitors suppressed IL-1b secretion. In contrast, an adenylate cyclase inhibitor alone induced dose-dependent secretion of IL-1b in macrophages (Lee et al., 2012). It has been demonstrated that EP3 reduces cAMP levels by inhibiting adenylate cyclase (Sugimoto et al., 1992; Kalinski, 2012), whereas EP2 and EP4 stimulate the formation of intracellular cAMP (Fujino et al., 2005; Kalinski, 2012). In this study, we found that inhibition of EP3 by a specific inhibitor or knockdown by shRNA reduced NLRP3 inflammasome activation. It should be noted that reduced cAMP levels may be responsible for the increased activation of the NLRP3 inflammasome caused by EP3 signaling. Celecoxib inhibits COX-2 activity by binding with its polar sulfonamide side chain in a hydrophilic side pocket region close

COX-2 REGULATES NLRP3 INFLAMMASOME

Figure 7. Effect of COX-2 on NLRP3 inflammasome activation in vivo. (A) Protein expression levels of COX-2 in spleen and PGE2 concentration in serum. (B) IL-1b concentration in serum. (C) Caspase-1 activity in spleen and liver. (D) Protein expression levels of NLRP3 in spleen and liver. (E) IL-6 concentration in serum. The data are expressed as the mean  SD for different mice. The western blot results are representative of data obtained in different mice and the histograms are presented as the change in the ratio relative to actin compared to control group. * and ** indicates a significant difference at the level of P < 0.05 and P < 0.01, respectively.

to the active COX-2 binding site (DiPiro et al., 2008). Therefore, celecoxib should not affect COX-2 protein expression in theory. However, we found reduced COX-2 protein expression in LPS-injected mice. It should be noted that COX-2 can be induced by IL-1b (Dinarello, 2009) and PGE2 (Díaz-Muñoz JOURNAL OF CELLULAR PHYSIOLOGY

et al., 2012). Thus, downregulation of IL-1b and PGE2 in vivo may be responsible for the decreased expression level of spleen COX-2 protein caused by celecoxib administration in mice. Exogenous PGE2 further increased NLRP3 inflammasomederived IL-1b secretion, but this effect was significantly reduced

873

874

HUA ET AL.

in COX-2 knockdown macrophages. This result suggested that COX-2 plays an important role in PGE2-mediated activation of the NLRP3 inflammasome, and more COX-2 expression induced by PGE2 may be responsible for this effect. Our present results demonstrate a novel role of the well characterized molecule COX-2 in activating the NLRP3 inflammasome and suggest that COX-2-targeted therapy might be beneficial for diseases involving the NLRP3 inflammasome. These diseases include diabetes (Kellogg et al., 2007), obesity (Ghoshal et al., 2011), neural diseases (Aid and Bosetti, 2011), gout (Nalbant et al., 2005), kidney disease (Quilley et al., 2011), and cancer (Santander et al., 2012). Literature Cited Aid S, Bosetti F. 2011. Targeting cyclooxygenases-1 and -2 in neuroinflammation: Therapeutic implications. Biochimie 93:46–51. Anders HJ, Muruve DA. 2011. The inflammasomes in kidney disease. J Am Soc Nephrol 22:1007–1018. Bagi Z, Erdei N, Papp Z, Edes I, Koller A. 2006. Up-regulation of vascular cyclooxygenase-2 in diabetes mellitus. Pharmacol Rep 58:52–56. Bauernfeind FG, Horvath G, Stutz A, Alnemri ES, MacDonald K, Speert D, FernandesAlnemri T, Wu J, Monks BG, Fitzgerald KA, Hornung V, Latz E. 2009. Cutting edge: NFkappaB activating pattern recognition and cytokine receptors license NLRP3 inflammasome activation by regulating NLRP3 expression. J Immunol 183:787–791. Bauernfeind F, Bartok E, Rieger A, Franchi L, Núñez G, Hornung V. 2011. Cutting edge: Reactive oxygen species inhibitors block priming, but not activation, of the NLRP3 inflammasome. J Immunol 187:613–617. Bergsbaken T, Fink SL, Cookson BT. 2009. Pyroptosis: Host cell death and inflammation. Nat Rev Microbiol 7:99–109. Cheng H, Fan X, Moeckel GW, Harris RC. 2011. Podocyte COX-2 exacerbates diabetic nephropathy by increasing podocyte (pro) renin receptor expression. J Am Soc Nephrol 22:1240–1251. Cuccurullo C, Fazia ML, Mezzetti A, Cipollone F. 2007. COX-2 expression in atherosclerosis: the good, the bad or the ugly. Curr Med Chem 14:1595–1605. Díaz-Muñoz MD, Osma-García IC, Fresno M, Iñiguez MA. 2012. Involvement of PGE2 and the cAMP signalling pathway in the up-regulation of COX-2 and mPGES-1 expression in LPS-activated macrophages. Biochem J 443:451–461. Dinarello CA. 2009. Immunological and inflammatory functions of the interleukin-1 family. Annu Rev Immunol 27:519–550. DiPiro JT, Talbert RL, Yee GC, Matzke GR, Wells BG, Posey LM. 2008. Pharmacotherapy: A pathophysiologic approach. New York: McGraw-Hill Medical. Duewell P, Kono H, Rayner KJ, Sirois CM, Vladimer G, Bauernfeind FG, Abela GS, Franchi L, Nuñez G, Schnurr M, Espevik T, Lien E, Fitzgerald KA, Rock KL, Moore KJ, Wright SD, Hornung V, Latz E. 2010. NLRP3 inflammasomes are required for atherogenesis and activated by cholesterol crystals. Nature 464:1357–1361. Fink SL, Cookson BT. 2006. Caspase-1-dependent pore formation during pyroptosis leads to osmotic lysis of infected host macrophages. Cell Microbiol 8:1812–1825. Fraser PA. 2011. The role of free radical generation in increasing cerebrovascular permeability. Free Radic Biol Med 51:967–977. Fujino H, Salvi S, Regan JW. 2005. Differential regulation of phosphorylation of the cAMP response element-binding protein after activation of EP2 and EP4 prostanoid receptors by prostaglandin E2. Mol Pharmacol 68:251–259. Ghoshal S, Trivedi DB, Graf GA, Loftin CD. 2011. Cyclooxygenase-2 deficiency attenuates adipose tissue differentiation and inflammation in mice. J Biol Chem 286:889–898. Halle A, Hornung V, Petzold GC, Stewart CR, Monks BG, Reinheckel T, Fitzgerald KA, Latz E, Moore KJ, Golenbock DT. 2008. The NALP3 inflammasome is involved in the innate immune response to amyloid-beta. Nat Immunol 9:857–865. Hornung V, Bauernfeind F, Halle A, Samstad EO, Kono H, Rock KL, Fitzgerald KA, Latz E. 2008. Silica crystals and aluminum salts activate the NALP3 inflammasome through phagosomal destabilization. Nat Immunol 9:847–856. Hsieh YC, Mounsey RB, Teismann P. 2011. MPP(þ)-induced toxicity in the presence of dopamine is mediated by COX-2 through oxidative stress. Naunyn Schmiedebergs Arch Pharmacol 384:157–167. Hsu HY, Wen MH. 2002. Lipopolysaccharide-mediated reactive oxygen species and signal transduction in the regulation of interleukin-1 gene expression. J Biol Chem 277:22131– 22139. Jacob S, Laury-Kleintop L, Lanza-Jacoby S. 2008. The select cyclooxygenase-2 inhibitor celecoxib reduced the extent of atherosclerosis in apo E-/- mice. J Surg Res 146:135–142. Jin C, Flavell RA. 2010. Molecular mechanism of NLRP3 inflammasome activation. J Clin Immunol 30:628–631. Juliana C, Fernandes-Alnemri T, Kang S, Farias A, Qin F, Alnemri ES. 2012. Nontranscriptional priming and deubiquitination regulate NLRP3 inflammasome activation. J Biol Chem 287:36617–36622. Kalinski P. 2012. Regulation of immune responses by prostaglandin E2. J Immunol 188:21–28. Kanneganti TD, Ozören N, Body-Malapel M, Amer A, Park JH, Franchi L, Whitfield J, Barchet W, Colonna M, Vandenabeele P, Bertin J, Coyle A, Grant EP, Akira S, Núñez G. 2006. Bacterial RNA and small antiviral compounds activate caspase-1 through cryopyrin/ Nalp3. Nature 440:233–236. Kellogg AP, Wiggin TD, Larkin DD, Hayes JM, Stevens MJ, Pop-Busui R. 2007. Protective effects of cyclooxygenase-2 gene inactivation against peripheral nerve dysfunction and intraepidermal nerve fiber loss in experimental diabetes. Diabetes 56:2997–3005.

JOURNAL OF CELLULAR PHYSIOLOGY

Kepp O, Galluzzi L, Kroemer G. 2011. Mitochondrial control of the NLRP3 inflammasome. Nat Immunol 12:199–200. Khan Z, Khan N, Tiwari RP, Sah NK, Prasad GB, Bisen PS. 2011. Biology of Cox-2: An application in cancer therapeutics. Curr Drug Targets 12:1082–1093. Latz E, Xiao TS, Stutz A. 2013. Activation and regulation of the inflammasomes. Nat Rev Immunol 13:397–411. Lee GS, Subramanian N, Kim AI, Aksentijevich I, Goldbach-Mansky R, Sacks DB, Germain RN, Kastner DL, Chae JJ. 2012. The calcium-sensing receptor regulates the NLRP3 inflammasome through Ca2þ and cAMP. Nature 492:123–127. Liao PC, Chao LK, Chou JC, Dong WC, Lin CN, Lin CY, Chen A, Ka SM, Ho CL, Hua KF. 2013. Lipopolysaccharide/adenosine triphosphate-mediated signal transduction in the regulation of NLRP3 protein expression and caspase-1-mediated interleukin-1b secretion. Inflamm Res 62:89–96. Martinon F, Petrilli V, Mayor A, Tardivel A, Tschopp J. 2006. Gout-associated uric acid crystals activate the NALP3 inflammasome. Nature 440:237–241. McCormack PL. 2011. Celecoxib: A review of its use for symptomatic relief in the treatment of osteoarthritis, rheumatoid arthritis and ankylosing spondylitis. Drugs 71:2457–2489. Miao EA, Rajan JV, Aderem A. 2011. Caspase-1-induced pyroptotic cell death. Immunol Rev 243:206–214. Mifflin RC, Saada JI, Di Mari JF, Adegboyega PA, Valentich JD, Powell DW. 2002. Regulation of COX-2 expression in human intestinal myofibroblasts: mechanisms of IL-1-mediated induction. Am J Physiol Cell Physiol 282:C824–C834. Miller DK, Myerson J, Becker JW. 1997. The interleukin-1 beta converting enzyme family of cysteine proteases. J Cell Biochem 64:2–10. Moore SF, MacKenzie AB. 2009. NADPH oxidase NOX2 mediates rapid cellular oxidation following ATP stimulation of endotoxin-primed macrophages. J Immunol 83:3302–3308. Mouithys-Mickalad A, Deby-Dupont G, Dogne JM, de Leval X, Kohnen S, Navet R, Sluse F, Hoebeke M, Pirotte B, Lamy M. 2004. Effects of COX-2 inhibitors on ROS produced by Chlamydia pneumoniae-primed human promonocytic cells (THP-1). Biochem Biophys Res Commun 325:1122–1130. Nalbant S, Chen LX, Sieck MS, Clayburne G, Schumacher HR. 2005. Prophylactic effect of highly selective COX-2 inhibition in acute monosodium urate crystal induced inflammation in the rat subcutaneous air pouch. J Rheumatol 32:1762–1764. Nakahira K, Haspel JA, Rathinam VA, Lee SJ, Dolinay T, Lam HC, Englert JA, Rabinovitch M, Cernadas M, Kim HP, Fitzgerald KA, Ryter SW, Choi AM. 2011. Autophagy proteins regulate innate immune responses by inhibiting the release of mitochondrial DNA mediated by the NALP3 inflammasome. Nat Immunol 12:222–230. Nasrallah R, Robertson SJ, Hebert RL. 2009. Chronic COX inhibition reduces diabetesinduced hyperfiltration, proteinuria, and renal pathological markers in 36-week B6-Ins2 (Akita) mice. Am J Nephrol 30:346–353. Okamoto M, Liu W, Luo Y, Tanaka A, Cai XD, Norris AC, Dinarello A, Fujita M. 2010. Constitutively active inflammasome in human melanoma cells mediating autoinflammation via caspase-1 processing and secretion of interleukin-1beta. J Biol Chem 285:6477–6488. Pasinetti GM, Aisen PS. 1998. Cyclooxygenase-2 expression is increased in frontal cortex of Alzheimer’s disease brain. Neuroscience 87:319–324. Py BF, Kim MS, Vakifahmetoglu-Norberg H, Yuan J. 2013. Deubiquitination of NLRP3 by BRCC3 critically regulates inflammasome activity. Mol Cell 49:331–338. Quilley J, Santos M, Pedraza P. 2011. Renal protective effect of chronic inhibition of COX-2 with SC-58236 in streptozotocin-diabetic rats. Am J Physiol Heart Circ Physiol 300: H2316–H2322. Redondo S, Ruiz E, Gordillo-Moscoso A, Navarro-Dorado J, Ramajo M, Rodríguez E, Reguillo F, Carnero M, Casado M, Tejerina T. 2011. Overproduction of cyclo-oxygenase-2 (COX-2) is involved in the resistance to apoptosis in vascular smooth muscle cells from diabetic patients: a link between inflammation and apoptosis. Diabetologia 54:190–199. Santander S, Cebrian C, Esquivias P, Conde B, Esteva F, Jimenez P, Ortego J, Lanas A, Piazuelo E. 2012. Cyclooxygenase inhibitors decrease the growth and induce regression of human esophageal adenocarcinoma xenografts in nude mice. Int J Oncol 40:527–534. Schroder K, Zhou R, Tschopp J. 2010. The NLRP3 inflammasome: A sensor for metabolic danger. Science 327:296–300. Strowig T, Henao-Mejia J, Elinav E, Flavell R. 2012. Inflammasomes in health and disease. Nature 481:278–286. Sugimoto Y, Namba T, Honda A, Hayashi Y, Negishi M, Ichikawa A, Narumiya S. 1992. Cloning and expression of a cDNA for mouse prostaglandin E receptor EP3 subtype. J Biol Chem 267:6463–6466. Tschopp J, Schroder K. 2010. NLRP3 inflammasome activation: The convergence of multiple signalling pathways on ROS production. Nat Rev Immunol 10:210–215. Vandanmagsar B, Youm YH, Ravussin A, Galgani JE, Stadler K, Mynatt RL, Ravussin E, Stephens JM, Dixit VD. 2011. The NLRP3 inflammasome instigates obesity-induced inflammation and insulin resistance. Nat Med 17:179–188. Veltman JD, Lambers ME, van Nimwegen M, Hendriks RW, Hoogsteden HC, Aerts JG, Hegmans JP. 2010. COX-2 inhibition improves immunotherapy and is associated with decreased numbers of myeloid-derived suppressor cells in mesothelioma. Celecoxib influences MDSC function. BMC Cancer 10:464. Zhou R, Yazdi AS, Menu P, Tschopp J. 2011. A role for mitochondria in NLRP3 inflammasome activation. Nature 469:221–225. Zweers MC, de Boer TN, van Roon JJ, Bijlsma W, Lafeber FP, Mastbergen SC. 2011. Celecoxib: Considerations regarding its potential disease-modifying properties in osteoarthritis. Arthritis Res Ther 13:239.

Supporting Information

Additional supporting information may be found in the online version of this article at the publisher’s web-site.

Copyright of Journal of Cellular Physiology is the property of John Wiley & Sons, Inc. and its content may not be copied or emailed to multiple sites or posted to a listserv without the copyright holder's express written permission. However, users may print, download, or email articles for individual use.