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International Journal of Experimental Pathology logoLink to International Journal of Experimental Pathology
. 2016 Jan 14;96(6):395–405. doi: 10.1111/iep.12159

Amomum tsao‐ko fruit extract suppresses lipopolysaccharide‐induced inducible nitric oxide synthase by inducing heme oxygenase‐1 in macrophages and in septic mice

Ji‐Sun Shin 1,2,3, Suran Ryu 1,4, Dae Sik Jang 5, Young‐Wuk Cho 2,3, Eun Kyung Chung 6, Kyung‐Tae Lee 1,5,✉
PMCID: PMC4744820  PMID: 26852687

Summary

Amomum tsao‐ko Crevost et Lemarié (Zingiberaceae) has traditionally been used to treat inflammatory and infectious diseases, such as throat infections, malaria, abdominal pain and diarrhoea. This study was designed to assess the anti‐inflammatory effects and the molecular mechanisms of the methanol extract of A. tsao‐ko (AOM) in lipopolysaccharide (LPS)‐induced RAW 264.7 macrophages and in a murine model of sepsis. In LPS‐induced RAW 264.7 macrophages, AOM reduced the production of nitric oxide (NO) by inhibiting inducible nitric oxide synthase (iNOS) expression, and increased heme oxygenase‐1 (HO‐1) expression at the protein and mRNA levels. Pretreatment with SnPP (a selective inhibitor of HO‐1) and silencing HO‐1 using siRNA prevented the AOM‐mediated inhibition of NO production and iNOS expression. Furthermore, AOM increased the expression and nuclear accumulation of NF‐E2‐related factor 2 (Nrf2), which enhanced Nrf2 binding to antioxidant response element (ARE). In addition, AOM induced the phosphorylation of extracellular regulated kinase (ERK) and c‐Jun N‐terminal kinase (JNK) and generated reactive oxygen species (ROS). Furthermore, pretreatment with N‐acetyl‐l‐cysteine (NAC; a ROS scavenger) diminished the AOM‐induced phosphorylation of ERK and JNK and AOM‐induced HO‐1 expression, suggesting that ERK and JNK are downstream mediators of ROS during the AOM‐induced signalling of HO‐1 expression. In LPS‐induced endotoxaemic mice, pretreatment with AOM reduced NO serum levels and liver iNOS expression and increased HO‐1 expression and survival rates. These results indicate that AOM strongly inhibits LPS‐induced NO production by activating the ROS/MAPKs/Nrf2‐mediated HO‐1 signalling pathway, and supports its pharmacological effects on inflammatory diseases.

Keywords: Amomum tsao‐ko, HO‐1, NO, Nrf2, ROS, sepsis

Introduction

Inflammation is a normal response of living tissue to injury caused by physical or noxious chemical stimuli or microbiological toxins. However, chronic inflammation is an undesirable phenomenon that can ultimately result in inflammatory diseases, such as arthritis, asthma, multiple sclerosis, inflammatory bowel disease and atherosclerosis (Guzik et al. 2003; Rankin 2004). The pathogenesis of inflammation is a complex process that is regulated by cytokine networks and the inductions of many pro‐inflammatory genes (Guzik et al. 2003). Macrophages, which are widely distributed in the body, play a central role in orchestrating immune response through phagocytosis and pro‐inflammatory mediator secretions against foreign agents, such as lipopolysaccharide (LPS) (Kim et al. 2010a,b and c), and nitric oxide (NO; a potent pro‐inflammatory mediator) is mainly produced enzymatically in macrophages by inducible nitric oxide synthase (iNOS) at inflammatory sites, and acts as a cytotoxic agent during immune and inflammatory responses (Ryu et al. 2013). Therefore, the pharmacological suppression of NO production offers a strategy for modulating the potentially harmful pro‐inflammatory activity of macrophages (Kim et al. 2010a).

Phase II detoxifying and antioxidant enzymes, such as heme oxygenase‐1 (HO‐1), glutathione‐S‐transferase (GST), NAD(P)H quinine oxidoreductase 1 (NQO1), glutamate–cysteine ligase catalytic subunit (GCLC), and glutamate–cysteine ligase, modifier subunit (GCLM), catalyse a series of reactions essential for cellular defence and survival (Park et al. 2009). HO‐1 is an inducible enzyme that catalyses the rate‐limiting step in the oxidative degradation of cytotoxic free haeme into carbon monoxide (CO), biliverdin and free iron (Ryter et al. 2006). The Fe released from the protoporphyrin IX ring of haeme is then stored by the ferritin H chain (FtH). Biliverdin is converted by biliverdin reductase into the antioxidant bilirubin. Three end products of heme catabolism, that is biliverdin/bilirubin, CO and Fe/FtH provide a host defence mechanism against oxidative injury and inflammatory activities in cells and tissues (Gozzelino et al. 2010). When expressed in innate immune cells, HO‐1 exerts anti‐inflammatory effects that limit the damaging consequences of inflammation and immunity. In activated macrophages, HO‐1 expression inhibits the production of pro‐inflammatory mediators like NO (Suh et al. 2006). Moreover, an increasing number of therapeutic agents have been reported to induce HO‐1 expression and exert their anti‐inflammatory effects by inducing HO‐1. These studies support the beneficial effects of HO‐1 and its potential as a therapeutic target in inflammatory diseases (Malorni et al. 2007). HO‐1 expression is primarily induced by the binding of redox dependent transcription factors, such as activator protein‐1 (AP‐1), nuclear factor‐kappa B (NF‐κB) and NF‐E2‐related factor‐2 (Nrf2), to their promoter regions (Paine et al. 2010). In particular, Nrf2, which belongs to the cap'n'collar subfamily of basic leucine zipper (CNC‐bZIP) proteins is maintained in an inactive state by association with Kelch‐like ECH associating protein 1 (Keap1) under normal conditions. Upon oxidative stimulation, Nrf2 dissociates from Keap1, translocates to the nucleus and binds to the antioxidant response element (ARE) of HO‐1 promoter (Kim et al. 2010b). Furthermore, recent evidence suggests phosphatidylinositol 3‐kinase (PI3K)/Akt, mitogen‐activated protein kinases (MAPKs), and reactive oxygen species (ROS) act as intermediates in the signalling pathway responsible for Nrf2‐induced HO‐1 expression (Kim et al. 2010a; Paine et al. 2010).

Amomum tsao‐ko Crevost et Lemarié (Zingiberaceae) is widely distributed in the south‐west regions of China and Korea, and the dried fruit of this herb is commonly used as a traditional medicine for the treatment of throat infections, malaria, abdominal pain, stomach disorders, dyspepsia, nausea, vomiting and diarrhoea (Zhongyao 1977). Studies have demonstrated the anti‐HBV (Li et al. 1999), anti‐microbial (Yang et al. 2008), cytotoxic, apoptotic and antioxidant (Yang et al. 2010) activities of A. tsao‐ko. In addition, anti‐fungal, antioxidant, cytotoxic, anti‐proliferative and anti‐inflammatory constituents have been reported in A. tsao‐ko (Martin et al. 2000; Lee et al. 2008; Yang et al. 2010). For example, epicatechin and tsaokoin isolated from this herb were found to inhibit LPS‐induced NO production in BV2 microglial cells (Lee et al. 2008). Recently, it was demonstrated that ethanol extract of the fruits A. tsao‐ko exhibited anti‐inflammatory activities via Nrf2/HO‐1 induction in LPS‐induced RAW 264.7 macrophages (Li et al. 2014). However, the molecular mechanisms involved in Nrf2/HO‐1 induction and any anti‐inflammatory effects in animal model have not been evaluated yet. Accordingly, in this study, we investigated the molecular mechanisms responsible for the Nrf2/HO‐1 induction by the methanol extract of the fruits of A. tsao‐ko (AOM) in LPS‐induced RAW 264.7 macrophages and its anti‐inflammatory properties in a murine model of sepsis.

Materials and methods

Plant material and preparation of the methanol extract

The fruits of A. tsao‐ko Crevost et Lemarié were delivered by the Department of Oriental Pharmacy, Kyung Hee Medical Center and identified by Prof. Nam‐In Back (Kyung Hee University, Suwon, Korea). A voucher specimen (KHUOPS‐08‐51‐1) was deposited at the herbarium of the College of Pharmacy, Kyung Hee University (Seoul, Korea). The plant material (250 g) was extracted two times with 70% aqueous methanol (1 l) under reflux. The methanol extract was filtered and evaporated under reduced pressure to give a solid AOM (25.66 g), which was then stored at −20°C until required. To set a standard of AOM contents, total phenols and flavonoids contents were 157.02 and 33.01 mg/g, respectively (Tunalier et al. 2007). Epicatechin and tsaokoin of AOM were analysed and found to be present at concentrations of 20.4 ± 3.2 mg/g and 11.0 ± 1.1 mg/g by LC‐MS/MS, respectively.

Chemicals

Dulbecco's modified eagle's medium (DMEM), foetal bovine serum, penicillin and streptomycin were obtained from Life Technologies Inc. (Grand Island, NY, USA). iNOS, HO‐1, Nrf2, p‐c‐Jun N‐terminal kinase 1/2 (pJNK1/2), JNK1/2, p‐extracellular signal‐regulated kinase (pERK), ERK, poly(ADP ribose)polymerase (PARP), β‐actin monoclonal antibodies and peroxidase‐conjugated secondary antibody were purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA, USA). Random oligonucleotide primers and M‐MLV reverse transcriptase were purchased from Promega (Madison, WI, USA). SYBR green ex Taq were obtained from TaKaRa (Shiga, Japan). iNOS, GCLC, GCLM, NQO‐1 and β‐actin oligonucleotide primers were purchased from Bioneer (Seoul, Korea). 3‐(4,5‐Dimethylthiazol‐2‐yl)‐2,5‐diphenyl tertazolium bromide (MTT), sulphanilamide, aprotinin, leupeptin, phenylmethylsulfonyl fluoride (PMSF), DL‐Dithiothreitol (DTT), NS‐398, LPS (Escherichia coli, serotype 0111:B4), LPS (Salmonella enterica, serotype enteritidis), Triton X‐100, and all other chemicals were purchased from Sigma Chemical Co. (St Louis, MO, USA).

Cell culture and sample treatment

RAW 264.7 murine macrophage cell line was obtained from Korean Cell Line Bank (Seoul, Korea). Cells were grown at 37°C in DMEM medium supplemented with 10% FBS, penicillin (100 units/ml), and streptomycin sulphate (100 μg/ml) in a humidified 5% CO2 atmosphere. Cells were incubated with AOM at concentrations of 5, 10 and 20 μg/ml, or with an appropriated positive control, and then stimulated with LPS (1 μg/ml) for the indicated time. All samples dissolved in DMSO were added to the medium in serial dilution (the final DMSO concentration in all assays did not exceed 0.05% and did not affect the cells).

Nitrite determination

Nitrite levels in culture media were determined using the Griess reaction assay and presumed to reflect NO levels (Griess 1864).

Western blot analysis

Protein extracts from AOM‐treated cells were prepared as described previously (Shin et al. 2011). Cellular protein from treated and untreated cell extracts was electroblotted onto a PVDF membrane following separation on a 10‐12% SDS‐PAGE. The immunoblot was incubated for 1 h with blocking solution (5% skim milk), followed by incubation overnight with a primary antibody at 4°C. Blots were washed three times with Tween‐20/Tris‐buffered saline and incubated with a 1:1000 dilution of horseradish peroxidase‐conjugated secondary antibody for 2 h at room temperature. Blots were again washed three times with Tween‐20/Tris‐buffered saline, and then developed by enhanced chemiluminescence (GE healthcare, Milwaukee, WI, USA).

Real‐time quantitative RT‐PCR (qRT‐PCR)

Total cellular RNA was isolated by Easy Blue® kits (Intron Biotechnology, Seoul, Korea). For each sample, 500 ng of RNA was reverse‐transcribed (RT) using TOPscript™ RT DryMIX (Enzynomics™, Seoul, Korea) and 0.5 μg/μl Random primers (Promega). The PCR primers used for SYBR Green real‐time RT‐PCR are listed below and were purchased from Bioneer: sense strand for mouse iNOS, 5′‐AAT GGC AAC ATC AGG TCG GCC ATC ACT‐3′, anti‐sense strand for mouse iNOS, 5′‐GCT GTG TGT CAC AGA AGT CTC GAA CTC‐3′; sense strand for mouse HO‐1, 5′‐GCT TGT TGT TGC GCT CAT TCT CC‐3′, anti‐sense strand HO‐1, 5′‐GCC ACC AAG GAG GTA CAC ATA‐3′; sense strand GCLC, 5′‐ TCC GGC ATC GGA GAG GAG A‐3′, anti‐sense strand GCLC, 5′‐ AGC AGT TGC CCA TCC CGA AT‐3′; sense strand GCLM, 5′‐ GGG AAC CTG CTC AAC TGG GG‐3′, anti‐sense strand GCLM, 5′‐ CTG CAT GGG ACA TGG TGC ATT‐3′; sense strand NQO‐1, 5′‐ TGG CCG ATT CAG AGT GGC AT‐3′, anti‐sense strand NQO‐1, 5′‐ AAA CAG GCT GCT TGG AGC AAA A‐3′; sense strand for mouse β‐actin, 5′‐TCA TGA AGT GTG ACG TTG ACA TCC GT‐3′, anti‐sense strand for mouse β‐actin, 5′‐CCT AGA AGC ATT TGC GGT GCA CGA TG‐3′. Real‐time RT‐PCR was performed using Thermal Cycler Dice Real‐Time PCR System (Takara). A dissociation curve analysis of iNOS, HO‐1, GCLM, GCLM, NQO and β‐actin showed a single peak for each. The mean C t of the gene of interest was calculated from triplicate measurements and normalized with the mean C t of a control gene, β‐actin.

Nuclear extraction and electrophoretic mobility shift assay

RAW 264.7 macrophages were plated in 100‐mm dishes (1 × 106 cells/ml), and treated with AOM (20 μg/ml) for indicated time, washed once with PBS, scraped into 1 ml of cold PBS, and pelleted by centrifugation. Nuclear extracts were prepared as described previously (Shin et al. 2011). Nuclear extracts (5 μg) were mixed with double‐stranded Nrf2 oligonucleotide, 5′‐AACCATGACACAGCATAAA‐3′ and 5′‐TTTATGCTGTGTCATGGTT‐5′, respectively, and end‐labelled by biotin. DNA‐binding activities are measured using LightShftChemiluminescent electrophoretic mobility shift assay (EMSA) kit (Thermo Fisher Scientific, Waltham, MA, USA).

siRNA transfection

HO‐1 siRNA and control siRNA were purchased from Santa Cruz Biotechnology, Inc. siRNA was transfected into RAW 264.7 macrophages according to the manufacturer's protocol using the transfection reagent, Amaxa cell line nucleofector kit (Köln, Lonza, Germany). The cells were incubated with 400 nM of HO‐1 siRNA or control siRNA for 24 h and cells were washed and pretreated with or without AOM, following LPS stimulation.

Measurement of reactive oxygen species

Intracellular accumulation of ROS was determined using the fluorescent probes 2′, 7′‐dichlorodihydrofluorescein diacetate (H2DCFDA). Cells were seeded on a 6‐well plate for 24 h and treated with 20 μM H2DCFDA for 30 min prior to treatment with AOM 20 μg/ml. Then, cells were resuspended in PBS and analysed using flow cytometry (Beckman Coulter Inc, Brea, CA, USA). A total of 5000 events were acquired for analysis using cell quest software (Beckman Coulter Inc., Brea, CA, USA).

Animals

C57BL/6 male mice weighing 20–25 g were purchased from the Orient Bio Inc. (Seongnam‐si, Korea) and maintained under constant conditions (temperature: 20 ± 2°C, humidity: 40‐60%, light/dark cycle: 12 h). At 24 h before the experiment, only water was provided. All procedures were conducted under university guideline of ethical committee for Animal Care and Use of the Kyung Hee University according to an animal protocol (KHP‐2012‐05‐2).

Septic shock in mice

Male C57BL/6 mice, 6–8 weeks old, were administered AOM (10, 20 or 50 mg/kg, p.o.). After 1 h, septic shock was induced by injecting LPS (Salmonella enterica, 25 mg/kg, i.p). AOM was first dissolved in 5% EtOH and 5% Cremophor and diluted with saline. Survival rates were monitored over 48 h. Blood was collected (using the retrobulbar plexus route under sodium pentobarbital anaesthesia) 6 h after LPS injection, and allowed to clot at room temperature. Serum was separated by centrifugation, and stored at −80°C until NO analysis. Liver samples were obtained 6 h after LPS challenge and immediately frozen (−80°C) for the determination of HO‐1 and iNOS mRNA.

Statistical analysis

The data are expressed as the mean ± SD of at least three experiments performed using different cell preparations in vitro. Statistically significant values were compared using a one‐way anova followed by Dunnett's post hoc test and P‐values less than 0.05 were considered statistically significant. In animal study, the data were expressed as the mean ± SD (n = 8). In animal survival study (n = 5), the Kaplan–Meier method was used to estimate survival as a function of time, and differences in survival pattern among experimental groups were analysed by the log‐rank test. Statistical significance was defined as P < 0.05. If overall comparison turned out to be significant, pairwise comparisons were performed using Bonferroni correction.

Results

AOM attenuated LPS‐induced NO production and iNOS expression and induced HO‐1 expression in RAW 264.7 macrophages

To examine the inhibitory effects of AOM on LPS‐induced NO production and iNOS expression, cells were pretreated with AOM (5, 10, or 20 μg/ml) for 1 h and then treated with LPS (1 μg/ml) for 24 h. In accordance with previous study (Li et al. 2014), AOM concentration‐dependently inhibited LPS‐induced NO production and iNOS expression (Figure 1a,b). Next, to determine whether AOM regulates iNOS expression at mRNA, we treated cells with AOM in the presence of LPS, and assessed iNOS mRNA levels by qRT‐PCR at 6 h. AOM treatment significantly inhibited iNOS mRNA expression (Figure 1c). To determine whether AOM induces the expression of HO‐1, cells were treated with AOM (20 μg/ml) for 0, 2, 4, 8, 12 or 24 h. HO‐1 protein and mRNA levels were significantly increased by AOM and peaked at 8 h (protein) and 4 h (mRNA), respectively, after treatment (Figure 1d,e). These effects of AOM were not caused by nonspecific cytotoxicity, because AOM had no effect on cell viability as determined by MTT assay at concentrations from 6.25 to 50 μg/ml in the presence or absence of LPS (Figure 1f).

Figure 1.

Figure 1

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AOM suppressed LPS‐induced NO production and iNOS expression and induced HO‐1 expression in RAW 264.7 macrophages. (a) Following pretreatment with AOM (5, 10, or 20 μg/ml) for 1 h, cells were treated with LPS (1 μg/ml) for 24 h. Culture media were collected and subjected to a Griess assay to determine NO production levels. Controls were not treated with LPS or AOM. L‐NIL (20 μM) was used as a positive control. (b) Lysates were prepared from cells pretreated with/without indicated concentrations of AOM for 1 h and then treated with LPS (1 μg/ml) for 24 h. Total cellular proteins were resolved by SDS‐PAGE, transferred to PVDF membranes and detected using a specific iNOS antibody. (c) Total RNAs for qRT‐PCR were prepared from cells pretreated with/without the indicated concentrations of AOM for 1 h and then treated with LPS (1 μg/ml) for 4 h. iNOS levels were normalized against β‐actin. Data are presented as the means ± SDs of three independent experiments. # P < 0.05 vs. the non‐treated control group; ***P < 0.001 vs. LPS‐stimulated group. (d) Lysates were prepared from cells treated with AOM (20 μg/ml) for the indicated times. Total cellular proteins were resolved by SDS‐PAGE, transferred to PVDF membranes, and detected using a specific HO‐1 antibody. (e) Total RNAs for qRT‐PCR analysis were prepared from cells treated with AOM (20 μg/ml) for the indicated times. HO‐1 levels were normalized against β‐actin. (f) Cells were treated with AOM from 6.25 to 100 μg/ml for 24 h in the presence or absence of LPS (1 μg/ml). Cell viability were determined by MTT assay. Data are presented as the means ± SDs of three independent experiments. *P < 0.05, **P < 0.01 and ***P < 0.001 vs. non‐treated control group.

Involvement of HO‐1 in the inhibitions of LPS‐induced NO production and iNOS expression by AOM in RAW 264.7 macrophages

To determine whether AOM‐induced HO‐1 expression mediates its inhibitory effect on NO production, cells were pretreated with SnPP (a HO‐1 inhibitor) before administering AOM. Pretreatment with SnPP significantly prevented the inhibition of NO production by AOM (Figure 2a). Furthermore, the involvement of HO‐1 in the anti‐inflammatory effects of AOM was confirmed by transfecting cells with a specific HO‐1 siRNA, which substantially downregulated HO‐1 protein levels (Figure 2b). As shown in Figure 2c,d, HO‐1 knockdown markedly prevented the AOM‐mediated inhibition of NO production and iNOS expression, whereas transfection with control siRNA had no effect. These results indicate that the inhibitory effects of AOM on LPS‐induced NO production and iNOS expression were intimately linked with AOM‐induce expression of HO‐1.

Figure 2.

Figure 2

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The induction of HO‐1 mediated the AOM‐induced suppression of NO production and iNOS expression in RAW 264.7 macrophages. (a) Cells were pretreated with AOM (5, 10, or 20 μg/ml) in the presence or absence of SnPP (10 μM) for 1 h and then treated with LPS (1 μg/ml) for 24 h. (b) Cells were transfected with HO‐1 siRNA or control siRNA prior to AOM (20 μg/ml) treatment. HO‐1 deletion was determined by Western blotting using a specific HO‐1 siRNA. (c and d) After siRNA transfection, cells were pretreated with AOM (5, 10, or 20 μg/ml) for 1 h, and then treated with LPS (1 μg/ml) for 24 h. The culture medium and total protein extracts were then subjected to Griess reaction assay to determine NO levels (c) or to Western blotting to determine iNOS protein expression (d). Data are presented as the means ± SDs of three independent experiments. # P < 0.05 vs. the non‐treated control group; *P < 0.05, **P < 0.01, ***P < 0.001 vs. SnPP‐treated or HO‐1 siRNA‐transfected group.

AOM induced Nrf2 activation by increasing the DNA binding, expression and nuclear accumulation of Nrf2 in RAW 264.7 macrophages

Nrf2 is a key transcription factor of cytoprotective and detoxifying regulatory genes, including HO‐1 (Rushworth et al. 2008). To elucidate the mechanisms underlying HO‐1 induction by AOM, EMSA was used to examine the effect of AOM on the DNA binding of Nrf2. As shown in Figure 3a, Nrf2 appeared at 30 min after AOM treatment and peaked at 120 min. LPS (used as a positive control) also markedly increased this binding. AOM also increased the protein expression of Nrf2, which peaked at 60 min after the AOM treatment and then decreased. Nrf2 nuclear accumulation was detected within 30 min of AOM treatment and persistent to 240 min (Figure 3b). Furthermore, AOM concentration‐dependently increased the expression and nuclear accumulation of Nrf2 at 60 min post‐treatment (Figure 3c). In addition, AOM was found to concentration‐dependently induce the mRNA expressions of other Nrf2‐ARE regulated genes, such as, NQO1, GCLM and GCLC (Figure 3d–f), thus confirming the activation of Nrf2 in response to AOM.

Figure 3.

Figure 3

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AOM induced Nrf2 activation in RAW 264.7 macrophages. (a) Nuclear extracts were prepared from cells treated with AOM (20 μg/ml) for the indicated times or cells treated with LPS (1 μg/ml) for 1 h, and analysed for Nrf2 binding by EMSA. LPS was used as a positive control. The specificity of binding was examined by competition with 100‐fold unlabelled Nrf2 oligonucleotide (CP). (b and c) Cells were treated with AOM (20 μg/ml) for the indicated times (b) or with the indicated concentrations of AOM for 1 h (c). Total protein or nuclear protein extracts were resolved by SDS‐PAGE, transferred to PVDF membranes and detected with specific Nrf2 antibody. β‐Actin and PARP were used as internal controls. (d–f) Total RNAs for qRT‐PCR analysis were prepared from cells treated with indicated concentrations of AOM for 12 h (d) or 8 h (e) or 4 h (f). NQO‐1, GCLM and GCLC levels were normalized against β‐actin. Data are presented as the means ± SDs of three independent experiments. ***P < 0.001 vs. non‐treated control group.

AOM induced HO‐1 expression via the activation of MAPKs signalling

As MAPKs play crucial roles in the expression of HO‐1 (Papaiahgari et al. 2006), we examined the role of MAPKs signalling in AOM‐induced HO‐1 expression using specific pharmacological inhibitors. SB203580 (a p38 MAPK inhibitor), PD98059 (an ERK inhibitor) or SP600125 (a JNK inhibitor) were pretreated 1 h prior to AOM. As shown in Figure 4a, SP600125 and PD98059 inhibited AOM‐induced HO‐1 expression, but SB203580 did not. To confirm that AOM activates ERK and JNK, we investigated the phosphorylation statuses of ERK and JNK. AOM rapidly increased phosphorylated ERK and JNK levels, which peaked at 30 min (ERK) and 20 min (JNK) post‐treatment respectively (Figure 4b). These observations suggest that activations of the ERK and JNK signalling pathways are required for AOM‐induced HO‐1 expression.

Figure 4.

Figure 4

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AOM induced HO‐1 expression via ERK and JNK activation in RAW 264.7 macrophages. (a) Cells were pretreated with SB203580 (20 μM), PD98059 (20 μM) or SP600125 (10 μM) for 1 h and then treated with AOM (20 μM) for 8 h. (b) Lysates were prepared from cells treated with AOM (20 μg/ml) for the indicated times. Total protein extracts were resolved by SDS‐PAGE, transferred to PVDF membranes, and detected using specific HO‐1, pERK, ERK, pJNK and JNK antibodies.

ROS was involved in the AOM‐mediated expression of HO‐1 via MAPKs signalling

Next, we investigated whether AOM‐induced HO‐1 expression was triggered by ROS production. To accomplish this, we examined whether AOM induced intracellular ROS production using a H2DCFDA‐based flow cytometer detection method. Treatment with AOM at 20 μg/ml elevated intracellular ROS levels from 10 min and peaking at 120 min post‐treatment (Figure 5a). H2O2 was used as a positive control. When cells were pretreated with N‐acetyl‐l‐cysteine (NAC; a ROS scavenger), NAC significantly abolished AOM‐induced HO‐1 expression (Figure 5b). Western blotting also showed that the AOM‐induced phosphorylations of ERK and JNK were reduced by NAC pretreatment (Figure 5c), suggesting ROS acts upstream of the AOM‐induced activations of ERK and JNK.

Figure 5.

Figure 5

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AOM‐induced ROS production enhanced HO‐1 expression via ERK and JNK activation in RAW 264.7 macrophages. (a) Cells were incubated with H2 DCFDA (20 μM) for 30 min in the presence of AOM (20 μg/ml) or H2O2 (200 μM). Fluorescence intensities were measured by flow cytometry. H2O2 (200 μM) was used as a positive control for ROS production. (b and c) Cells were pretreated with NAC (5 mM) for 1 h and then treated with AOM (20 μM) for 8 h (HO‐1), 30 min (pERK) or 20 min (pJNK). Total protein extracts were resolved by SDS‐PAGE, transferred to PVDF membranes, and detected with specific HO‐1, pERK, ERK, pJNK and JNK antibodies.

AOM inhibited NO release and increased HO‐1 expression and survival rates in mice with LPS‐induced endotoxemia

The observation that AOM attenuated LPS‐induced NO release by inducing HO‐1 expression in macrophages encouraged us to explore its efficacy in an animal model of LPS‐induced endotoxemia. A LPS injection (25 mg/kg, i.p) remarkably increased NO serum levels, but the administration of AOM (10, 20, or 50 mg/kg, p.o) significantly decreased LPS‐induced NO release (Figure 6a). In liver tissues, LPS significantly increased iNOS mRNA expression, and this was inhibited dose‐dependently by the administration of AOM (Figure 6b). Furthermore, AOM significantly increased HO‐1 mRNA expression in the liver tissues of mice with LPS‐induced sepsis (Figure 6c). Finally, the survival rate of AOM‐administered mice (10, 20, or 50 mg/kg, p.o) was significantly greater than that of untreated mice after LPS treatment.

Figure 6.

Figure 6

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AOM reduced NO production and iNOS expression and increased HO‐1 expression and survival rates in LPS‐induced septic mice. (a) AOM (10, 20, or 50 mg/kg, p.o.) was administered to mice 1 h before injecting LPS (25 mg/kg, i.p.). Serum samples were collected 6 h after injecting LPS (25 mg/kg, i.p.). NO levels were determined using a Griess reaction assay. (b and c) Liver tissues were collected 6 h after injecting LPS (25 mg/kg, i.p.) and iNOS and HO‐1 expressions were assessed by qRT‐PCR. # P < 0.05 vs. non‐treated control group; *P < 0.05, **P < 0.01, ***P < 0.001 vs. LPS‐injected mice (n = 8). (d) Survival rates were observed over 48 h after LPS administration (n = 5) and percentage was shown;**P < 0.01 vs. LPS‐injected mice.

Discussion

HO‐1 has been recognized to exhibit important immunomodulatory and anti‐inflammatory effects (Paine et al. 2010). In a HO‐1 knockout mice model, animals were found to develop chronic inflammatory disease and to be highly vulnerable to the experimental sepsis induced by the classical pro‐inflammatory mediator endotoxin (Poss & Tonegawa 1997). Furthermore, phenotypic alterations in human cases of genetic HO‐1 deficiency are similar to those observed in HO‐1 knockout mice (Yachie et al. 1999). Pharmacological induction of HO‐1 or the administration of its end products (CO, biliverdin, and iron) can exert therapeutic effects in a variety of immune‐mediated inflammatory diseases, including sepsis, ischaemia–reperfusion injury, autoimmune neuro‐inflammation, myocardial infarction, diabetes and obesity (Soares & Bach 2009). A number of natural phytocompounds have been shown to be effective non‐stressful and non‐cytotoxic inducers of HO‐1 (Li Volti et al. 2008). Some polyphenols, such as curcumin, caffeic acid phenethyl ester, and resveratrol, can act as HO‐1 inducers and their abilities to induce HO‐1 are consistent with their anti‐inflammatory properties (Motawi et al. 2011; Zhong et al. 2013; Liu et al. 2014). It was reported by Li et al. (2014) that ethanol extract of A. tsao‐ko suppresses LPS‐induced inflammatory responses in RAW264.7 macrophages via Nrf2‐dependent HO‐1 expression. In this study, we attempted to explore the anti‐inflammatory effects of AOM and intensified molecular mechanisms underlying Nrf2/HO‐1 induction focusing on the ROS and MAPK in vitro and in vivo. AOM was found to significantly induce HO‐1 expression at the transcriptional level in RAW 264.7 macrophages. Furthermore, this induction of HO‐1 by AOM was correlated with the inhibition of LPS‐induced iNOS expression. In LPS‐induced RAW 264.7 macrophages, AOM significantly inhibited iNOS expression and NO production, and HO‐1 knockdown by siRNA or treatment with SnPP prevented the inhibitory effect of AOM on the NO production in LPS‐induced RAW 264.7 macrophages. These observations indicate that AOM inhibits iNOS expression and NO production by modulating the expression of HO‐1.

Because the promoter region of the HO‐1 gene contains multiple copies of ARE, many reports have suggested that HO‐1 induction is dependent on the activation of Nrf2. However, the mechanism by which Nrf2 is activated in response to its many inducers remains a matter of discussion. Certain stimuli promote the release of Nrf2 from its inhibitor Keap1, and thus, enable its nuclear translocation. Previous study reported that ethanol extract of A. tsao‐ko induced nuclear translocation of Nrf2 (Li et al. 2014). Similarly, AOM increased the nuclear accumulation of Nrf2 in this study. Interesting, we also found that AOM both upregulated the cellular protein expression of Nrf2. It was well‐known that increasing the cellular level of Nrf2 appears to be another mechanism regulating Nrf2 activation (Park et al. 2009). Although it is evident that Nrf2 protein induction is importantly required for ARE activation, the mechanism of cellular Nrf2 expression is controversial (Keum et al. 2008). Kwak et al. found that treatment with 3H‐1,2‐dithiole‐3‐thione increased Nrf2 mRNA expression in murine keratinocytes by providing the binding site of Nrf2 itself (Kwak et al. 2002). In contrast, other investigators have reported increases in Nrf2 protein levels are regulated by the attenuation of ubiquitin‐dependent 26S proteosomal degradation irrespective of Nrf2 mRNA expression (Nguyen et al. 2003; Keum et al. 2008). It remains to be determined whether the AOM‐induced nuclear accumulation of Nrf2 is mediated by the dissociation of Nrf2 from Keap1 or by an increase the total amount of Nrf2. Additional studies are needed to identify the mechanisms underlying the upregulation of cellular Nrf2 expression by AOM.

In the present study, AOM induced other Nrf2‐ARE regulated genes, namely NQO1, GCLM and GCLC, which demonstrates AOM is able to functionally activate Nrf2 in macrophages. NQO‐1 is a cytosolic antioxidant flavoprotein that catalyses the reduction of quinones to hydroquinones by utilizing NADH as an electron donor (Kim et al. 2013), whereas GCLC and GCLM are components of glutamate cysteine ligase, which synthesizes intracellular glutathione (GSH). Moreover, HO‐1, NQO‐1, GCLC and GCLM maintain redox homoeostasis and participate in intracellular anti‐inflammatory processes (Lu et al. 2014; Thapa et al. 2014). Therefore, it might be expected that the anti‐inflammatory activity of AOM is due to the enhancement of the Nrf2‐ARE regulated genes which are mediated cellular defence system.

In many previous studies, the MAPKs signalling system has been implicated in Nrf2 activation in responds to diverse stimuli, including oxidative stress (Yu et al. 2000). Sun et al. reported that inhibition of the catalytic activity of MAPKs, either through pharmaceutical inhibitors or genetic engineering, negatively modulates Nrf2 activation (Sun et al. 2009). Although the roles played by MAPKs in Nrf2 activation are not well understood, the phosphorylation of Nrf2 by MAPKs is presumed to facilitate the dissociation of Nrf2 from Keap1 and the subsequent nuclear translocation of Nrf2 (Surh 2003; Zipper & Mulcahy 2003). Nguyen et al. reported that the phosphorylation of Nrf2 by MAPKs (especially by ERK) increases the protein stability of Nrf2 via an ubiquitin‐dependent pathway (Nguyen et al. 2003), and it has also been reported that MAPKs signalling is activated by various Nrf2‐activating agents, such as phenethyl isothiocyanate, butylated hydroxyanisole and 3H‐1,2‐dithiole‐3‐thione (Cheung et al. 2013). In the present study, experiments with specific pharmacological inhibitors for p38 MAPK, JNK and ERK showed that JNK and ERK act upstream of AOM‐induced HO‐1 expression. Consistent with these findings, AOM significantly induced the phosphorylations of ERK and JNK. Interestingly, AOM also increased the phosphorylation of p38 MAPK (data not shown), but this activation was not involved in AOM‐induced HO‐1 expression in RAW 264.7 macrophages.

ROS has been implicated in multiple physiological and pathological processes as a secondary messenger in cell signalling (Kong et al. 2010). Several studies have demonstrated ROS generation plays a role in the modulation of HO‐1. Some HO‐1 inducers, such as taurine chloramine and tanshinone, generate ROS and have been reported to induce HO‐1 expression in macrophages (Chen et al. 2007; Kim et al. 2010b). Itoh et al. hypothesized that ROS may react with one or more of the multiple cysteine residues on Keap1 to cause a conformational change that liberates Nrf2 (Itoh et al. 1999). Furthermore, ROS play a role in the phosphorylations of MAPKs in various cells (Fubini & Hubbard 2003; Keshari et al. 2013). Riemann et al. reported that scavenging of ROS prevented MAPKs phosphorylation whereas direct application of ROS (H2O2) enhanced it (Riemann et al. 2011). The present study shows AOM generated ROS in macrophages and subsequently increased HO‐1 expression via the activations of ERK and JNK, and in line with these results, increased HO‐1 expression and the phosphorylations of ERK and JNK caused by AOM were abolished by exposure to NAC.

As Nrf2‐ARE regulated genes contribute to the potentiation of antioxidant defence capacity in cells, the modulation of Nrf2‐ARE signalling may have profound effects on redox‐sensitive inflammation‐regulating factors, such as NF‐κB and AP‐1. Although it remains unclear whether the inhibitions of the activities of NF‐κB and AP‐1 by Nrf2‐activating agent are indeed mediated via Nrf2 activation, treatments with Nrf2‐activating agents, such as dithiolethione, sulforaphane and curcumin, have been shown to inhibit the activations of NF‐κB and AP‐1 by inhibiting their translocations or DNA‐binding activities in various inflammatory cells (Kim et al. 2010c). In the present study, we investigated the effects of AOM on the LPS‐induced transcriptional activities of NF‐κB and AP‐1 using luciferase assays, but AOM did not affect these activities (Figure S1). These results suggest that AOM exerts its anti‐inflammatory activities through NF‐κB and AP‐1‐independent pathway. However, it was conflicting with previous result demonstrating that ethanol extract of A. tsao‐ko was found to inhibit the LPS‐induced NF‐κB activation (Li et al. 2014). We assumed that these opposite activity might be due to the difference in extracted constituents and extraction efficiency according to the polarity of the solvent (ethanol vs. methanol). Furthermore, the difference in action concentration between two extract supported our hypothesis. AOM was required a much lower concentration for NO inhibition and HO‐1 induction than ethanol extract (ethanol extract: 400 μg/ml vs. AOM: 20 μg/ml) in RAW 264.7 macrophages.

To verify the in vivo relevance of our in vitro results regarding the HO‐1‐mediated anti‐inflammatory effect of AOM, we evaluated its protective effects in a murine model of LPS‐induced sepsis. LPS is the most frequent cause of sepsis, and circulating NO, pro‐inflammatory cytokine, and chemokine levels are elevated under this condition. In fact, reductions of these elevated NO and pro‐inflammatory cytokine levels are an important therapeutic target in sepsis (Dinarello 1997; Van Amersfoort et al. 2003). In our murine model of LPS‐induced endotoxemia, AOM protected against LPS‐induced lethality and reduced NO serum levels. Furthermore, AOM significantly reduced liver iNOS expression, but induced HO‐1 expression. These results suggest that AOM‐induced HO‐1 expression is involved in the protection afforded by AOM in our murine model of sepsis.

In summary, we investigated the anti‐inflammatory effect of AOM and its association with the induction of HO‐1 in vitro and in vivo. It was found that the significant anti‐inflammatory effects of AOM were due to the inhibition of LPS‐induced NO release and iNOS expression via HO‐1 induction in RAW 264.7 macrophages and in our LPS‐induced murine model of sepsis. Furthermore, the induction of HO‐1 expression was mediated via ROS, and this ROS production lead to the activation of the ERK‐JNK/Nrf2 signalling pathway. Taken together, these findings suggest that the induction of HO‐1 expression underlies the anti‐inflammatory effect of AOM.

Ethical approval statement

All animal procedures were approved by university guideline of ethical committee for Animal Care and Use of the Kyung Hee University according to an animal protocol (KHP‐2012‐05‐2).

Conflict of interest

The authors have declared no conflict of interests.

Funding source

This research was supported by the Basic Science Research Program through the National Research Foundation of Korea (NRF), which is funded by the Ministry of Education, Science and Technology (No. 2013R1A1A2011043).

Supporting information

Data S1. Supplementary materials.

Figure S1. Effects of AOM on LPS‐induced transcriptional activation of NF‐κB and AP‐1 in RAW 264.7 macrophages (a and b) Cells were transiently transfected with pNF‐κB‐luc vector or pAP‐1‐luc vector; phRL‐TK‐vector was used as the internal control. Transfected cells were stimulated with LPS (1 μg/mL) in presence or absence of AOM (5, 10, or 20 μg/mL) for 18 h. Luciferase activities were determined using Promega luciferase assay system. Data are presented as the means ± SDs of three independent experiments. # P < 0.05 vs. the control group.

Click here for additional data file. (42.3KB, docx)

References

  1. Chen T.H., Hsu Y.T., Chen C.H., Kao S.H. & Lee H.M. (2007) Tanshinone IIA from Salvia miltiorrhiza induces heme oxygenase‐1 expression and inhibits lipopolysaccharide‐induced nitric oxide expression in RAW 264.7 cells. Mitochondrion 7, 101–105. [DOI] [PubMed] [Google Scholar]
  2. Cheung K.L., Lee J.H., Shu L., Kim J.H., Sacks D.B. & Kong A.N. (2013) The Ras GTPase‐activating‐like protein IQGAP1 mediates Nrf2 protein activation via the mitogen‐activated protein kinase/extracellular signal‐regulated kinase (ERK) kinase (MEK)‐ERK pathway. J. Biol. Chem. 288, 22378–22386. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Dinarello C.A. (1997) Proinflammatory and anti‐inflammatory cytokines as mediators in the pathogenesis of septic shock. Chest 112, 321S–329S. [DOI] [PubMed] [Google Scholar]
  4. Fubini B. & Hubbard A. (2003) Reactive oxygen species (ROS) and reactive nitrogen species (RNS) generation by silica in inflammation and fibrosis. Free Radic. Biol. Med. 34, 1507–1516. [DOI] [PubMed] [Google Scholar]
  5. Gozzelino R., Jeney V. & Soares M.P. (2010) Mechanisms of cell protection by heme oxygenase‐1. Annu. Rev. Pharmacol. Toxicol. 50, 323–354. [DOI] [PubMed] [Google Scholar]
  6. Griess J.P. (1864) On a new series of bodies in which nitrogen is substituted for hydrogen. Philos. Trans. R. Soc. Lond. 154, 667–731. [Google Scholar]
  7. Guzik T.J., Korbut R. & Adamek‐Guzik T. (2003) Nitric oxide and superoxide in inflammation and immune regulation. J. Physiol. Pharmacol. 54, 469–487. [PubMed] [Google Scholar]
  8. Itoh K., Wakabayashi N., Katoh Y. et al (1999) Keap1 represses nuclear activation of antioxidant responsive elements by Nrf2 through binding to the amino‐terminal Neh2 domain. Genes Dev. 13, 76–86. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Keshari R.S., Verma A., Barthwal M.K. & Dikshit M. (2013) Reactive oxygen species‐induced activation of ERK and p38 MAPK mediates PMA‐induced NETs release from human neutrophils. J. Cell. Biochem. 114, 532–540. [DOI] [PubMed] [Google Scholar]
  10. Keum Y.S., Chang P.P., Kwon K.H. et al (2008) 3‐Morpholinopropyl isothiocyanate is a novel synthetic isothiocyanate that strongly induces the antioxidant response element‐dependent Nrf2‐mediated detoxifying/antioxidant enzymes in vitro and in vivo. Carcinogenesis 29, 594–599. [DOI] [PubMed] [Google Scholar]
  11. Kim A.N., Jeon W.K., Lee J.J. & Kim B.C. (2010a) Up‐regulation of heme oxygenase‐1 expression through CaMKII‐ERK1/2‐Nrf2 signalling mediates the anti‐inflammatory effect of bisdemethoxycurcumin in LPS‐stimulated macrophages. Free Radic. Biol. Med. 49, 323–331. [DOI] [PubMed] [Google Scholar]
  12. Kim C., Jang J.S., Cho M.R., Agarawal S.R. & Cha Y.N. (2010b) Taurine chloramine induces heme oxygenase‐1 expression via Nrf2 activation in murine macrophages. Int. Immunopharmacol. 10, 440–446. [DOI] [PubMed] [Google Scholar]
  13. Kim J., Cha Y.N. & Surh Y.J. (2010c) A protective role of nuclear factor‐erythroid 2‐related factor‐2 (Nrf2) in inflammatory disorders. Mutat. Res. 690, 12–23. [DOI] [PubMed] [Google Scholar]
  14. Kim J.H., Jung S.H., Yang Y.I. et al (2013) Artemisia leaf extract induces apoptosis in human endometriotic cells through regulation of the p38 and NFkappaB pathways. J. Ethnopharmacol. 145, 767–775. [DOI] [PubMed] [Google Scholar]
  15. Kong X., Thimmulappa R., Kombairaju P. & Biswal S. (2010) NADPH oxidase‐dependent reactive oxygen species mediate amplified TLR4 signaling and sepsis‐induced mortality in Nrf2‐deficient mice. J. Immunol. 185, 569–577. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Kwak M.K., Itoh K., Yamamoto M. & Kensler T.W. (2002) Enhanced expression of the transcription factor Nrf2 by cancer chemopreventive agents: role of antioxidant response element‐like sequences in the nrf2 promoter. Mol. Cell. Biol. 22, 2883–2892. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Lee K.Y., Kim S.H., Sung S.H. & Kim Y.C. (2008) Inhibitory constituents of lipopolysaccharide‐induced nitric oxide production in BV2 microglia isolated from Amomum tsao‐ko. Planta Med. 74, 867–869. [DOI] [PubMed] [Google Scholar]
  18. Li Volti G., Sacerdoti D., Di Giacomo C. et al (2008) Natural heme oxygenase‐1 inducers in hepatobiliary function. World J. Gastroenterol. 14, 6122–6132. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Li W., Zou Z.Y., Zha G., Zheng M.S. & Huang X.T. (1999) Experimental research on anti‐HBV activity of 170 kinds of Chinese herbs. World Chinese J. Digestol. 7, 89–90. [Google Scholar]
  20. Li B., Choi H.J., Lee D.S. et al (2014) Amomum tsao‐ko suppresses lipopolysaccharide‐induced inflammatory responses in RAW264.7 macrophages via Nrf2‐dependent heme oxygenase‐1 expression. Am. J. Chin. Med. 42, 1229–1244. [DOI] [PubMed] [Google Scholar]
  21. Liu H., Ren J., Chen H. et al (2014) Resveratrol protects against cigarette smoke‐induced oxidative damage and pulmonary inflammation. J. Biochem. Mol. Toxicol. 28, 465–471. [DOI] [PubMed] [Google Scholar]
  22. Lu C.Y., Yang Y.C., Li C.C., Liu K.L., Lii C.K. & Chen H.W. (2014) Andrographolide inhibits TNFalpha‐induced ICAM‐1 expression via suppression of NADPH oxidase activation and induction of HO‐1 and GCLM expression through the PI3K/Akt/Nrf2 and PI3K/Akt/AP‐1 pathways in human endothelial cells. Biochem. Pharmacol. 91, 40–50. [DOI] [PubMed] [Google Scholar]
  23. Malorni W., Campesi I., Straface E., Vella S. & Franconi F. (2007) Redox features of the cell: a gender perspective. Antioxid. Redox Signal. 9, 1779–1801. [DOI] [PubMed] [Google Scholar]
  24. Martin T.S., Kikuzaki H., Hisamoto M. & Nakatani N. (2000) Constituents of Amomum tsao‐ko and their radical scavenging and antioxidant activities. J. Am. Oil Chem. Soc. 77, 667–673. [Google Scholar]
  25. Motawi T.K., Darwish H.A. & Abd El Tawab A.M. (2011) Effects of caffeic acid phenethyl ester on endotoxin‐induced cardiac stress in rats: a possible mechanism of protection. J. Biochem. Mol. Toxicol. 25, 84–94. [DOI] [PubMed] [Google Scholar]
  26. Nguyen T., Sherratt P.J., Huang H.C., Yang C.S. & Pickett C.B. (2003) Increased protein stability as a mechanism that enhances Nrf2‐mediated transcriptional activation of the antioxidant response element. Degradation of Nrf2 by the 26 S proteasome. J. Biol. Chem. 278, 4536–4541. [DOI] [PubMed] [Google Scholar]
  27. Paine A., Eiz‐Vesper B., Blasczyk R. & Immenschuh S. (2010) Signaling to heme oxygenase‐1 and its anti‐inflammatory therapeutic potential. Biochem. Pharmacol. 80, 1895–1903. [DOI] [PubMed] [Google Scholar]
  28. Papaiahgari S., Zhang Q., Kleeberger S.R., Cho H.Y. & Reddy S.P. (2006) Hyperoxia stimulates an Nrf2‐ARE transcriptional response via ROS‐EGFR‐PI3K‐Akt/ERK MAP kinase signaling in pulmonary epithelial cells. Antioxid. Redox Signal. 8, 43–52. [DOI] [PubMed] [Google Scholar]
  29. Park P.H., Kim H.S., Jin X.Y. et al (2009) KB‐34, a newly synthesized chalcone derivative, inhibits lipopolysaccharide‐stimulated nitric oxide production in RAW 264.7 macrophages via heme oxygenase‐1 induction and blockade of activator protein‐1. Eur. J. Pharmacol. 606, 215–224. [DOI] [PubMed] [Google Scholar]
  30. Poss K.D. & Tonegawa S. (1997) Reduced stress defense in heme oxygenase 1‐deficient cells. Proc. Natl. Acad. Sci. USA 94, 10925–10930. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Rankin J.A. (2004) Biological mediators of acute inflammation. AACN Clin. Issues 15, 3–17. [DOI] [PubMed] [Google Scholar]
  32. Riemann A., Schneider B., Ihling A. et al (2011) Acidic environment leads to ROS‐induced MAPK signaling in cancer cells. PLoS One 6, e22445. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Rushworth S.A., MacEwan D.J. & O'Connell M.A. (2008) Lipopolysaccharide‐induced expression of NAD(P)H:quinone oxidoreductase 1 and heme oxygenase‐1 protects against excessive inflammatory responses in human monocytes. J. Immunol. 181, 6730–6737. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Ryter S.W., Alam J. & Choi A.M. (2006) Heme oxygenase‐1/carbon monoxide: from basic science to therapeutic applications. Physiol. Rev. 86, 583–650. [DOI] [PubMed] [Google Scholar]
  35. Ryu S., Shin J.S., Cho Y.W. et al (2013) Fimasartan, anti‐hypertension drug, suppressed inducible nitric oxide synthase expressions via nuclear factor‐kappa B and activator protein‐1 inactivation. Biol. Pharm. Bull. 36, 467–474. [DOI] [PubMed] [Google Scholar]
  36. Shin J.S., Noh Y.S., Lee Y.S. et al (2011) Arvelexin from Brassica rapa suppresses NF‐kappaB‐regulated pro‐inflammatory gene expression by inhibiting activation of IkappaB kinase. Br. J. Pharmacol. 164, 145–158. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Soares M.P. & Bach F.H. (2009) Heme oxygenase‐1: from biology to therapeutic potential. Trends Mol. Med. 15, 50–58. [DOI] [PubMed] [Google Scholar]
  38. Suh G.Y., Jin Y., Yi A.K., Wang X.M. & Choi A.M. (2006) CCAAT/enhancer‐binding protein mediates carbon monoxide‐induced suppression of cyclooxygenase‐2. Am. J. Respir. Cell Mol. Biol. 35, 220–226. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Sun Z., Huang Z. & Zhang D.D. (2009) Phosphorylation of Nrf2 at multiple sites by MAP kinases has a limited contribution in modulating the Nrf2‐dependent antioxidant response. PLoS One 4, e6588. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Surh Y.J. (2003) Cancer chemoprevention with dietary phytochemicals. Nat. Rev. Cancer 3, 768–780. [DOI] [PubMed] [Google Scholar]
  41. Thapa D., Meng P., Bedolla R.G., Reddick R.L., Kumar A.P. & Ghosh R. (2014) NQO1 suppresses NF‐kappaB‐p300 interaction to regulate inflammatory mediators associated with prostate tumorigenesis. Cancer Res. 74, 5644–5655. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Tunalier Z., Kosar M., Kupeli E., Calis I. & Baser K.H. (2007) Antioxidant, anti‐inflammatory, anti‐nociceptive activities and composition of Lythrum salicaria L. extracts. J. Ethnopharmacol. 110, 539–547. [DOI] [PubMed] [Google Scholar]
  43. Van Amersfoort E.S., Van Berkel T.J. & Kuiper J. (2003) Receptors, mediators, and mechanisms involved in bacterial sepsis and septic shock. Clin. Microbiol. Rev. 16, 379–414. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Yachie A., Niida Y., Wada T. et al (1999) Oxidative stress causes enhanced endothelial cell injury in human heme oxygenase‐1 deficiency. J. Clin. Invest. 103, 129–135. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Yang Y., Yan R.W., Cai X.Q., Zheng Z.L. & Zou G.L. (2008) Chemical composition and antimicrobial activity of the essential oil of Amomum tsao‐ko. J. Sci. Food Agric. 88, 2111–2116. [Google Scholar]
  46. Yang Y., Yue Y., Runwei Y. & Guolin Z. (2010) Cytotoxic, apoptotic and antioxidant activity of the essential oil of Amomum tsao‐ko. Bioresour. Technol. 101, 4205–4211. [DOI] [PubMed] [Google Scholar]
  47. Yu R., Chen C., Mo Y.Y. et al (2000) Activation of mitogen‐activated protein kinase pathways induces antioxidant response element‐mediated gene expression via a Nrf2‐dependent mechanism. J. Biol. Chem. 275, 39907–39913. [DOI] [PubMed] [Google Scholar]
  48. Zhong Y., Liu T., Lai W., Tan Y., Tian D. & Guo Z. (2013) Heme oxygenase‐1‐mediated reactive oxygen species reduction is involved in the inhibitory effect of curcumin on lipopolysaccharide‐induced monocyte chemoattractant protein‐1 production in RAW264.7 macrophages. Mol. Med. Rep. 7, 242–246. [DOI] [PubMed] [Google Scholar]
  49. Zhongyao D. (1977) Dictionary of Chinese Materia Medical, Jiangsu New Medical College, Shanghai, Science and Technology Press of Shanghai, pp. 1574‐1575.
  50. Zipper L.M. & Mulcahy R.T. (2003) Erk activation is required for Nrf2 nuclear localization during pyrrolidine dithiocarbamate induction of glutamate cysteine ligase modulatory gene expression in HepG2 cells. Toxicol. Sci. 73, 124–134. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Data S1. Supplementary materials.

Figure S1. Effects of AOM on LPS‐induced transcriptional activation of NF‐κB and AP‐1 in RAW 264.7 macrophages (a and b) Cells were transiently transfected with pNF‐κB‐luc vector or pAP‐1‐luc vector; phRL‐TK‐vector was used as the internal control. Transfected cells were stimulated with LPS (1 μg/mL) in presence or absence of AOM (5, 10, or 20 μg/mL) for 18 h. Luciferase activities were determined using Promega luciferase assay system. Data are presented as the means ± SDs of three independent experiments. # P < 0.05 vs. the control group.

Click here for additional data file. (42.3KB, docx)

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