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Division of Food Science and Biotechnology, Graduate School of Agriculture, Kyoto University, Kyoto 606-8502, Japan
4To whom correspondence should be addressed. E-mail: ohigashi{at}kais.kyoto-u.ac.jp.
ABSTRACT
In the present study, we explored the suppressive activities of 1'-acetoxychavicol acetate (ACA), auraptene, nobiletin, and zerumbone toward LPS-induced cyclooxygenase (COX)-2 mRNA expression in mouse macrophages and the underlying molecular mechanisms. Pretreatment of RAW264.7 cells with LPS led to the activation of mitogen-activated protein kinase (MAPK)s [p38, extracellular signal-regulated kinase (ERK)1/2, c-Jun NH2-terminal kinase (JNK)1/2] and Akt, together with degradation of the inhibitor of nuclear factor-
B (IB)-
protein and nuclear translocation of nuclear factor (NF)-B p65, and the resultant activation of activator protein (AP)-1, NF-B, and cAMP-responsive element-binding protein (CREB) transcription factors. ACA abrogated ERK1/2 and JNK1/2, but not p38 MAPK, as well as the activation of those transcription factors. Although it allowed LPS-triggered phosphorylation of those MAPKs and NF-B nuclear translocation, nobiletin suppressed the activation of AP-1, NF-B, and CREB. Zerumbone had no effect on those transcription factors, though it attenuated COX-2 mRNA expression, suggesting that it disrupts the stabilization of COX-2 mRNA. Conversely, zerumbone significantly accelerated spontaneous COX-2 mRNA decay, the potency of which was comparable with that of SB203580, an inhibitor of p38 MAPK, whose activation has key roles in the proinflammatory mRNA stabilization processes. Because SB203580 but not zerumbone suppressed LPS-induced p38 MAPK activation, the molecular targets of zerumbone may be MAPK-activated protein kinase-2 or located downstream. However, auraptene suppressed the expression of COX-2 protein but not mRNA, implying that it targets translation. We propose that these phytochemicals are promising chemopreventive agents for inflammation-associated carcinogenesis. Their use in combination may enhance their efficacy because of their different modes of action.
KEY WORDS: • cyclooxygenase-2 • cancer prevention • anti-inflammation • macrophage • dietary factor
Inflammation is a pathophysiological phenomenon that is involved in numerous diseases, including the development of neoplasms. Stromal activation of inflammatory cells induces dormant tumor cells to grow and progress into malignant tumors. Upon stimulation, those cells produce and secrete a cocktail of biochemical mediators, including reactive oxygen and nitrogen species and proinflammatory chemokines and cytokines, as well as eicosanoids such as prostaglandins (PGs).5 Cyclooxygenase (COX; PGH2 synthase) donates 2 oxygen molecules to arachidonic acid to form PGG2 by peroxidation, which in turn is reduced to PGH2. Nonsteroidal anti-inflammatory drugs such as aspirin have received considerable attention for their ability not only to mitigate inflammatory responses but also potentially to prevent cancer incidence in the human colon (1). Conversely, numerous animal experiments have demonstrated the cancer preventive efficacy of COX inhibitors. The COX enzyme consists of at least two isoforms, COX-1 and COX-2. In contrast to COX-1, COX-2 protein is only slightly expressed in most normal mammalian tissues in response to physical, chemical, and biological stimuli, including UV light exposure, dioxin, and lipopolysaccharide insult. Recently, COX-2 has received the attention of numerous researchers regarding the relation of its expression to pathogenesis. In particular, ample evidence exists of the involvement of COX-2 expression in carcinogenesis in many different target organs (2).
We screened Japanese (3) and subtropical vegetables and fruits (4–6) for their anti–tumor-promoting activities and identified some active constituents [reviewed in Nakamura et al. (7)]. It is notable that 1'-acetoxychavicol acetate (ACA, from Alpinia galanga, Zingiberaceae) and zerumbone (from Zingiber zerumbet, Zingiberaceae) as well as auraptene and nobiletin (from citrus fruits), which are readily available from natural sources and/or chemically synthesized, prevented chemical carcinogenesis in several organs (8–16), including the colon, in mouse and rat experiments. The mechanisms of action of these agents are not fully understood, though important findings based on the results of cellular and animal experiments have accumulated. Those mechanisms include the induction of phase II enzymes and apoptosis, attenuation of reactive oxygen and nitrogen species from inflammatory cells, regulation of proinflammatory cytokines, and inhibition of mutagenesis. In addition, these dietary factors have shown a marked ability to suppress COX-2 expression both in vitro and in vivo (8,10,17,18), though their molecular mechanisms remain largely unknown. In the present study, we investigated the mechanisms by which ACA, auraptene, nobiletin, and zerumbone attenuate LPS-induced COX-2 mRNA expression in RAW264.7 mouse macrophages.
Materials and methods
Reagents and cells. ACA (19), auraptene (20), nobiletin (12), and zerumbone (5) were purified as previously reported. DMEM and fetal bovine serum (FBS) were purchased from Invitrogen. LPS (Escherichia coli serotype 0127, B8) was purchased from Difco Labs. All other chemicals were purchased from Wako Pure Chemical Industries unless specified otherwise. RAW264.7 murine macrophages were purchased from the American Type Culture Collection.
RT-PCR. RAW264.7 cells (1 x 106) were grown to confluence in 5 mL DMEM with 10% FBS on 6-well plates, then incubated in an atmosphere containing 5% CO2 at 37°C for 13 h. The cells were washed with PBS twice, after which the medium was replaced with FBS- and phenol-red-free medium (2.5 mL) containing samples dissolved in 25 µL dimethylsulfoxide (DMSO). After 30 min of preincubation, the cells were treated with LPS (100 µg/L). After a 6-h incubation, the cells were lysed, and total RNA was extracted using kits (RNeasy® minikit and QIAshredder®, Qiagen). One microgram of total RNA was reverse transcribed using an RNA PCR Kit® (Takara) with an oligo dT-adaptor primer, as recommended by the supplier. Then, PCR assays were performed using a thermal cycler (PTC-0100; MJ Research) with hypoxanthine phosphoribosyl transferase (HPRT) and COX-2 using primers synthesized by Proligo with 1 µL of a cDNA preparation, 45 µL of Platinum® PCR SuperMix (Invitrogen), and 2 µL of each primer (1 µmol/L) as follows: HPRT (5'-gTAATgATCAgTCAACggggAC-3' and 5'-CCAgCAAgCTTgcAACCTTAACCA-3'), 20 cycles at 94°C for 2 min, 58°C for 2 min, and 72°C for 2 min; and COX-2 (5'-gCATTCTTTgCCCAgCACTT-3' and 5'-AgACCAggCACCAgACCAAAgA-3'), 20 cycles at 94°C for 30 s, 59°C for 30 s, and 72°C for 30 s. The PCR products were separated on 2% NuSieve® 3:1 agarose (BioWhittaker Molecular Applications), and each band was visualized using 0.01% SYBR Gold® stain (Molecular Probes). The amplified products were photographed with a digital camera, and the band intensities were analyzed using NIH Image software. For each target gene, we calculated the relative expression levels of mRNA in the samples as the ratio of the amount of each gene mRNA to that of HPRT mRNA. The number of PCR cycles was optimized so that each band intensity increased proportionally as the amount of cDNA increased. Each experiment was done at least 3 times.
Western blotting.
RAW264.7 cells (2 x 106) were grown to confluence in 4 mL DMEM with 10% FBS on a 60-mm dish, then incubated in an atmosphere containing 5% CO2 at 37°C for 13 h. The cells were washed with PBS twice, after which the medium was exchanged with FBS- and phenol-red-free mediium (10 mL) containing samples dissolved in 20 µL DMSO. After 30 min of preincubation, the cells were treated with LPS (100 µg/L) for various times. After being washed with PBS twice, the cells were separated into nuclear and cytosol fractions using a kit (Bio Vision; Research Products). Protein concentrations were determined using a DC Protein Assay kit (Bio-Rad), with 
-globulin employed as the standard. Next, 10 or 20 µg of proteins (for the nuclear and cytosol fractions, respectively) were separated on 10% polyacrylamide gels and electrophoretically transferred onto polyvinylidene difluoride membranes (Millipore). After blocking, the membranes were incubated with rabbit anti-proteinase inhibitor (Pi)-extracellular signal-regulated kinase (ERK)1/2, rabbit anti-ERK1/2 (Promega), rabbit anti-Pi-c-Jun NH2-terminal kinase (JNK)1/2 antibody, rabbit anti-JNK1/2 antibody, rabbit anti-Pi-p38 mitogen-activated protein kinase (MAPK) antibody, rabbit anti-p38 antibody (Cell Signaling), rabbit anti-Pi-Ser473 Akt1/2/3 antibodies, rabbit anti-Akt1/2/3 antibody, rabbit anti–inhibitor of nuclear factor B (IB)- antibody, rabbit anti-nuclear factor (NF)-B p65 antibody, rabbit anti-MAPK-activated protein kinase (MAPKAPK)-2 (Santa Cruz Biotechnology), and goat anti-ß-actin antibody (Biochemical Technologies) (1:1000 dilution each) and then with the corresponding secondary antibodies [horseradish peroxidase (HRP)-conjugated anti-rabbit IgG, 1:1000 dilution (Dako); or HRP-conjugated anti-goat IgG, 1:1000 dilution (Dako)]. The blots were developed using an ECL detection kit (Amersham Life Science). Each experiment was done at least 3 times.
Reporter assays.
The following reporter assays were performed using a Mercury® Pathway Profiling System (Clontech Laboratories) with some modifications. RAW264.7 cells (3 x 105 cells/mL) were preincubated on a 24-well plate for 12 h. Next, 625 µL OPTI-MEM® (Invitrogen) and 37.5 µL LipofectAMINE Reagent® (Invitrogen) were mixed in a tube, after which 4 µg of either pNF-B-, pAP-1, or pcAMP-responsive element-binding protein (CREB)-luciferase vector, provided in the kit, and 4 µg of pRL-TK vector (Promega), which served as the internal standard, were added and allowed to stand at room temperature for 30 min, after which an additional 5 mL OPTI-MEM was added. After the cells were washed with Hanks’ buffer twice, 250 µL transfection mixture was added to each well, and the cells were incubated at 37°C for 6 h. After being washed, the cells were incubated in 1 mL DMEM with 10% FBS for 12 h. The cells were washed again with Hanks’ buffer twice, then exposed to the vehicle (0.5% DMSO, v:v) or samples dissolved in DMSO in serum-free DMEM for 30 min. After stimulation of the cells with LPS (100 µg/L) for 12 h, luciferase activity in the cell lysate was determined using a Dual-Luciferase Reporter Assay Kit® (Promega).
COX-2 mRNA decay. RAW264.7 cells (2 x 106) were grown to confluence in 3 mL DMEM with 10% FBS on 6-well plates, then incubated in an atmosphere containing 5% CO2 at 37°C for 13 h. The cells were washed with PBS twice, after which the medium was exchanged with FBS- and phenol-red-free medium (3 mL) containing LPS (100 µg/L) and incubated for 4.5 h. Then the cells were treated with actinomycin D (1 µg/mL final concentration) for 30 min, followed by treatment with the vehicle or sample for 6 h. RT-PCR was performed as described above.
In vitro kinase assay.
A cell-free kinase assay was performed using a Kinase-Glo® Luminescent Kinase Assay (Promega). To a disposable culture tube (12 x 75 mm; Iwaki), 1.5 µL recombinant MAPKAPK-2 (PanVera), 0.2 µL recombinant active p38 (PanVera), and 23.3 µL kinase buffer (25 mmol/L HEPES, pH 7.6; 2 mmol/L dithiothreitol; 20 mmol/L MgCl2; 0.1 mmol/L NaVO4) were added. After 1 µL of each sample and 25 µL ATP solution (4 µmol/L) were added, the reaction mixture was incubated at 28°C for 3 h. Thereafter, 50 µL Kinase-Glo Reagent (Promega) was added, and the culture was incubated at room temperature for 15 min before the determination of chemiluminescence.
Statistical analysis. Each experiment was done 3 times unless specified otherwise, with values shown as means ± SD. The significance of differences between groups in each assay was assessed using a Student’s t-test (2-sided) that assumed unequal variance.
Results
Effects on COX-2 mRNA expression. Treatment of RAW264.7 macrophages with LPS for 6 h caused a marked induction of COX-2 mRNA as detected by RT-PCR. The concentration of each dietary factor (Fig. 1A) was nonlethal up to the highest concentration tested. Both zerumbone and ACA (20 µmol/L each) abolished LPS-induced COX-2 expression, whereas nobiletin, but not auraptene, moderately attenuated that expression (Fig. 1B).
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B system.
LPS stimulation induced the time-dependent phosphorylation of p38 MAPK from 0 to 60 min, whereas that of JNK1/2 and ERK1/2 peaked at 30 min after the LPS challenge. Activation of p38 was not attenuated by zerumbone, ACA, or nobiletin (Fig. 2A). However, ACA substantially blocked both JNK1/2 and ERK1/2 activation, whereas both zerumbone and nobiletin allowed LPS-induced activation of those protein kinases. Furthermore, the phosphorylation of Akt (protein kinase B) at Ser473 was detectable in nontreated cells and enhanced by LPS in a time-dependent manner (Fig. 2B). None of the 3 compounds demonstrated any inhibition. LPS-induced NF-B activation is mediated through the degradation of IB, a suppressive partner of NF-B, leading to the nuclear translocation of NF-B for transactivation of numerous proinflammatory and antiapoptotic genes. As shown in Figure 2B, IB protein was scarcely seen at 30 min after the LPS challenge and was restored within 60 min. Conversely, NF-B p65 was significantly translocated into the nucleus from 30 to 60 min. ACA did not allow those LPS-induced biochemical processes to occur, whereas both zerumbone and nobiletin were virtually inactive, except that the nuclear protein level of p65 in the nobiletin-treated cells was comparable with that in the nontreated cells at 60 min.
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B, AP-1, and CREB.
We transfected the NF-B-, AP-1-, and CREB-luciferase vectors to examine the effects of dietary factors on those transcriptional activities (Fig. 3). LPS treatment markedly elevated the activities of the transcription factors by 2.1- to 6.8-fold. It is notable that ACA and nobiletin significantly suppressed the 3 transcription factors, whereas zerumbone had no effect.
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The dietary factors examined in the present study were shown to be cancer preventive in mouse and rat models (8–16). Although the mechanisms of action have not been fully elucidated, some of our previous findings in regard to their biological and biochemical activities may be helpful. For example, oral feeding of ACA and auraptene led to a marked elevation of phase II drug metabolizing enzymes, including glutathione S-transferase (GST) and NAD(P)H quinone oxidoreductase (NQO1), in the colon and liver of rats (14,22). Similarly, zerumbone induced the activation of transcription factor Nrf2 in RL34 rat hepatocytes for upregulating the antioxidative and self-protecting enzymes -glutamylcysteine synthetase, glutathione peroxidase (GPx), and hemeoxygenase-1 (23). These results are consistent with our other in vivo observations that a topical application of zerumbone enhanced or induced mRNA expression by the manganese superoxide dismutase gene as well as the GPx, GST-P1, and NQO1 genes without affecting the expression levels of cytochrome P450 1A1 and 1B1 mRNA in mouse epidermis (8). Furthermore, the antioxidative and antinitrosative properties of that compound, which are related to the attenuation of endotoxin- or phorbol ester–induced superoxide anion and nitric oxide generation in inflammatory cells, were described in other cell culture and animal model studies (7,12,17–20).
ACA, auraptene, nobiletin, and zerumbone were found to suppress COX-2 protein production in macrophages, whereas oral and topical administration of zerumbone suppressed phorbol ester– and azoxymethane-induced COX-2 protein expression in mouse skin (6) and rat colon (10), respectively, though the molecular mechanisms were not fully elucidated. The present results provide some preliminary but useful insights into the molecular mechanisms of these dietary factors (Fig. 5). Auraptene may disrupt the translation step of COX-2 mRNA, because it was reported to suppress expression of the protein in LPS-treated RAW264.7 cells (18), whereas it did not suppress mRNA expression in our experiment (Fig. 1). In addition, we recently found that auraptene suppresses the expression of protein but not mRNA of matrix metalloproteinase-7 in HT-29 human colon cancer cells (K. Kawabata et al., unpublished data, 2005).
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B, and CREB, as shown previously for the involvement of ERK1/2 and JNK1/2 with those transcriptional factors. For example, ERK activation is involved in the phosphorylation of p65, thereby activating NF-B (24); a similar observation was made for RAW264.7 cells (25). Furthermore, the importance of Ser536 as the phosphorylation site of p65 was proposed (26). Chen et al. (27) reported that ERK activation led to NF-B activation that was related to COX-2 expression, and other lines of evidence exist for crosstalk activity between the ERK and CREB pathways (28–30); the effects of ACA on protein kinase A activation should be explored in the future. Ample evidence shows that activation of the JNK pathway leads to AP-1 transcriptional activation, because phosphorylated c-Jun, one of the direct targets of JNK, is a component of the AP-1 complex (31), suggesting that ACA suppresses AP-1 activation by blocking JNK activation, as shown in the present study (Fig. 5). The direct targets of ACA, which are probably located upstream of ERK/JNK, should be investigated in the future.
Transcription coactivators, including CREB-binding protein and its homologue p300, are known to play major roles in various stimuli-induced or -repressed intracellular signaling pathways (32) and were shown to potentiate the transcriptional activities of AP-1 (33), NF-B (34), and CREB (35). In addition, ASC-2 has emerged as a novel coactivator participating in the transcriptional activation of NF-B and AP-1 (36). Thus, nobiletin may disrupt the binding of those coactivators with their corresponding partner transcription factors, because we found that it suppressed NF-B, AP-1, and CREB activation, whereas it was virtually inactive in suppression of the activation of MAPKs and the Akt-NF-B pathway (Figs. 2, 3, and 5).
The mRNA expression of many, if not all, proinflammatory genes is regulated by posttranscriptional mechanisms (37). Those genes, including the COX-2 gene, contain highly conserved AU-rich elements in the proximal regions of the 3'-untranslated region (23). Some binding proteins for AU-rich elements were identified and shown to highly affect COX-2 mRNA stability in RAW264.7 macrophages (38). Of importance, the binding abilities of those proteins are regulated by the p38 MAPK pathway (39). We found that zerumbone (20 µmol/L) abrogated LPS-induced COX-2 mRNA expression (Fig. 1B), whereas it did not show any suppression of the transcriptional activation of NF-B, AP-1, and CREB. It is likely that CCAAT/enhancer binding protein ß and 
also may not be suppressed, because their production depends on NF-B transcriptional activity (40). This raises the possibility that zerumbone suppresses COX-2 mRNA expression via a posttranscriptional mechanism. In the present study, zerumbone accelerated spontaneous degradation of COX-2 mRNA (Fig. 4A), was virtually inactive for suppressing p38 MAPK activation (Fig. 2), and slightly attenuated MK2 phosphorylation (Fig. 4B) whereas our in vitro kinase assay revealed that it was definitely inactive regarding inhibition of p38 MAPK activity (Fig. 4C). Based on these findings, we hypothesize that zerumbone inhibits MK2 activity or targets unidentified molecules located downstream of MK2 (Fig. 5).
In conclusion, ACA targets both JNK1/2 and ERK1/2, and nobiletin may interfere with coactivators, such as CREB-binding protein/p300, that suppress the transactivation of NF-B, AP-1, and CREB. Zerumbone allowed LPS-induced MAPKs/Akt activation and transcriptional activation of those transcription factors, whereas it abrogated COX-2 mRNA induction. Thus, zerumbone may target MK-2 or downstream molecules for destabilizing mRNA. In addition, auraptene may disturb the translation process of COX-2 mRNA. because it suppressed COX-2 protein but not mRNA production. These results indicate that the dietary factors studied here have distinct molecular mechanisms for COX-2 protein suppression and that their use in combination may lead to synergistic results with higher efficacy and lower toxicity.
FOOTNOTES
1 Published in a supplement to The Journal of Nutrition. Presented as part of the International Research Conference on Food, Nutrition, and Cancer held in Washington, DC, July 14–15, 2005. This conference was organized by the American Institute for Cancer Research and the World Cancer Research Fund International and sponsored by (in alphabetical order) California Avocado Commission; California Walnut Commission; Campbell Soup Company; The Cranberry Institute; Danisco USA, Inc.; The Hormel Institute; National Fisheries Institute; The Solae Company; and United Soybean Board. Guest editors for this symposium were Vay Liang W. Go, Ritva R. Butrum, and Helen A. Norman. Guest Editor Disclosure: R. R. Butrum and H. Norman are employed by conference sponsor American Institute for Cancer Research; V.L.W. Go, no relationships to disclose. 
2 Author Disclosure: No relationships to disclose.
3 Supported by grants-in-aid from the Ministry of Agriculture, Forestry, and Fisheries Food Research Project, Integrated Research on Safety and Physiological Function of Food.
5 Abbreviations used: ACA, 1'-acetoxychavicol acetate; AP-1, activator protein-1; COX, cyclooxygenase; CREB, cAMP-responsive element-binding protein; DMSO, dimethylsulfoxide; ERK, extracellular signal-regulated kinase; FBS, fetal bovine serum; GPx, glutathione peroxidase; GST, glutathione S-transferase; HPRT, hypoxanthine phosphoribosyl transferase; HRP, horseradish peroxidase; IB, inhibitor of NF-B; JNK, c-Jun NH2-terminal kinase; LPS, lipopolysaccharide; MAPK, mitogen-activated protein kinase; MAPKAPK-2, MAPK-activated protein kinase-2; NF-B, nuclear factor B; NQO1, NAD(P)H quinone oxidoreductase; PG, prostaglandin; Pi, proteinase inhibitor.
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