Biflorin, Isolated from the Flower Buds of - ACS Publications

Mar 15, 2016 - ABSTRACT: Two chromone C-glucosides, biflorin (1) and isobiflorin (2), were isolated from the flower buds of Syzygium aromaticum L...
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Biflorin, Isolated from the Flower Buds of Syzygium aromaticum L., Suppresses LPS-Induced Inflammatory Mediators via STAT1 Inactivation in Macrophages and Protects Mice from Endotoxin Shock Hwi-Ho Lee,†,‡ Ji-Sun Shin,†,§,⊥ Woo-Seok Lee,†,‡ Byeol Ryu,‡ Dae Sik Jang,*,‡ and Kyung-Tae Lee*,†,‡ †

Department of Pharmaceutical Biochemistry, College of Pharmacy, ‡Department of Life and Nanopharmaceutical Sciences, College of Pharmacy, §Reactive Oxygen Species Medical Research Center College of Pharmacy, and ⊥Department of Physiology, School of Medicine, Kyung Hee University, Seoul 130-701, Republic of Korea S Supporting Information *

ABSTRACT: Two chromone C-glucosides, biflorin (1) and isobiflorin (2), were isolated from the flower buds of Syzygium aromaticum L. (Myrtaceae). Here, inhibitory effects of 1 and 2 on lipopolysaccharide (LPS)-induced production of nitric oxide (NO) and prostaglandin E2 (PGE2) in RAW 264.7 macrophages were evaluated, and 1 (IC50 = 51.7 and 37.1 μM, respectively) was more potent than 2 (IC50 > 60 and 46.0 μM). The suppression of NO and PGE2 production by 1 correlated with inhibition of iNOS and COX-2 protein expression. Compound 1 reduced inducible NO synthase (iNOS) and cyclooxygenase-2 (COX-2) mRNA expression via inhibition of their promoter activities. Compound 1 inhibited the LPS-induced production and mRNA expression of tumor necrosis factor-α (TNF-α) and interleukin (IL)-6. Furthermore, 1 reduced p-STAT1 and p-p38 expression but did not affect the activity of nuclear factor κ light-chain enhancer of activated B cells (NF-κB) or activator protein 1 (AP-1). In a mouse model of LPS-induced endotoxemia, 1 reduced the mRNA levels of iNOS, COX-2, and TNF-α, and the phosphorylation-mediated activation of the signal transducer and activator of transcription 1 (STAT1), consequently improving the survival rates of mice. Compound 1 showed a significant anti-inflammatory effect on carrageenan-induced paw edema and croton-oil-induced ear edema in rats. The collective data indicate that the suppression of pro-inflammatory gene expression via p38 mitogen-activated protein kinase and STAT1 inactivation may be a mechanism for the anti-inflammatory activity of 1.

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achieved by orchestrating chemokine production and leukocyte apoptosis.8 Macrophages are the major cellular source of these factors, which, in turn, participate in the mediation of acute phase responses to injury.5,9 Therefore, reducing activation signals in activated macrophages has been suggested as a therapeutic strategy for various inflammatory diseases.10 Stimulation of Tolllike receptor 4 (TLR4) on macrophages by lipopolysaccharide (LPS) promotes the recruitment of myeloid differentiation factor 88/IL-1 receptor-associated kinase (IRAK)/TNF receptorassociated factor 6 (TRAF6) to the receptor’s intracellular complex and results in the activation of transforming growth factor-β-activated kinase 1 (TAK1) in macrophages.11 Activated TAK1 subsequently induces the activation of transcription factors such as nuclear factor-κB (NF-κB) and activator protein-1 (AP-1), signal transducers and activators of transcriptions (STATs), and interferon response factors (IRFs).12,13 These

nflammation is a damaging component of many human diseases, including arteriosclerosis, inflammatory bowel disease (IBD), arthritis, infectious diseases, and cancer.1 The pathogenesis of inflammation is a complex process that is regulated by cytokine networks and many pro-inflammatory products such as nitric oxide (NO), prostaglandin E2 (PGE2), tumor necrosis factor-α (TNF-α), and interleukin (IL)-6. NO is produced enzymatically by inducible nitric oxide synthase (iNOS) at inflammatory sites and acts as a cytotoxic agent during immune and inflammatory responses.2 Large amounts of PGE2 are generated during the inflammatory process due to the increased expression of cyclooxygenase-2 (COX-2) and play an important role in the regulation of inflammatory responses such as fever, pain hypersensitivity, and edema.3,4 Cytokines like TNF-α, IL-1β, and IL-6 have been reported to be crucial mediators of inflammation during acute response to injury.5 TNF-α is induced by a wide range of pathogenic stimuli and performs a key role during the orchestration of inflammatory responses.6,7 IL-6 is a pivotal pro-inflammatory cytokine and a crucial checkpoint regulator of neutrophil trafficking, which is © XXXX American Chemical Society and American Society of Pharmacognosy

Received: July 10, 2015

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activated transcription factors then regulate various proinflammatory mediator and cytokine genes. The Janus kinase (JAK)-STAT cascade is an essential inflammatory signaling pathway that mediates immune responses. TLR4 activation brings about the phosphorylation of receptor-associated JAK, which, in turn, leads to STAT phosphorylation.14 Upon phosphorylation, STATs are released from the receptor complex and form homo- or heterodimers with each other, after which they translocate to the nucleus where they bind to the conserved DNA sequences in the promoter regions of target genes and activate transcription of those genes.15 Genetic ablation of STAT1 protected LPS-induced lethality, suggesting that STAT1 may play a key role in TLR4-induced inflammation.16 A similar role may be in humans expressing defective STAT1 mutants, where the hosts have increased susceptibility to viral and bacterial infections.17 Cloves are dried flower buds of Syzygium aromaticum (L.) Merr. & Perry (Myrtaceae). They are especially renowned as a relief for toothaches and for mouth and throat inflammation.18 Pharmacological studies with flower buds of S. aromaticum have shown antibacterial,19 antifungal,20 antioxidant,21 antinociceptive, and anti-inflammatory activities,22 along with immunomodulatory effects.23 In macrophages, clove extract inhibited LPSinduced production of NO, PGE2, IL-6, IL-1β, and IL-8.24−26 To date, a number of bioactive compounds have been isolated from the flower bud of S. aromaticum such as eugenol, tannins, sesquiterpenes, flavonoids, and chromones.27 Eugenol is a major volatile constituent comprising 72−90% of the essential oils extracted from cloves. The diverse pharmacological activities of eugenol include antimicrobial,28 anti-inflammatory,29 antioxidant,30 and anticancer activities.31,32 Biflorin (5,7-dihydroxy-2methylchromone-6-C-β-D-glucopyranoside, 1) and isobiflorin (5,7-dihydroxy-2-methylchromone-8-C-β-D-glucopyranoside, 2) are two major chromone C-glucosides in cloves (Figure 1).33

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RESULTS AND DISCUSSION

To compare the effects of biflorin (1) and isobiflorin (2) on LPSinduced NO and PGE2 production in RAW 264.7 macrophages, cells were treated with/without 1 or 2 (15, 30, or 60 μM) for 1 h and then treated with LPS (10 ng/mL) for 24 h. As shown in Figure 2A,B, 2 had relatively mild inhibitory effects on NO and PGE2 production, in contrast to the higher inhibitory effects of 1 (IC50 value for NO was 51.7 μM for 1 vs >60 μM for 2; for PGE2, 37.1 μM for 1 vs 46.0 μM for 2). L-NIL (20 μM) and NS-398 (3 μM) were used as positive controls to block the production of NO and PGE2, respectively. Based on these observations, we suggest that 1 is likely to be more effective than 2 as an antiinflammatory agent. Compound 1 is a 6-C-glucoside, and 2 is an 8-C-glucoside. Our results suggest that the position of the sugar moiety in the chromone backbone is an important factor influencing the anti-inflammatory activities in macrophages. This insight may help to identify other novel chromone C-glucoside candidates for anti-inflammatory properties and provide additional information on the structure−activity relationships relevant to such properties. The cytotoxicity of 1 (0−400 μM) on RAW 264.7 macrophages over 24 h was examined using the MTT assay. Compound 1 gradually decreased the cell viability from 100 μM (IC50 = 140 μM) and did not affect cell viability at 60 μM in the presence or absence of LPS. Therefore, the suppressive effects of 1 on NO and PGE2 production could not be attributable to nonspecific cytotoxicity. Next, we investigated whether inhibition of NO and PGE2 production by 1 was associated with reductions in the expression of iNOS or COX-2. As shown in Figure 2C, the protein levels of iNOS and COX-2 were markedly increased in response to LPS in RAW 264.7 macrophages, whereas 1 inhibited this effect in a concentration-dependent manner. To further elucidate the mechanism of 1-induced changes in iNOS and COX-2 protein levels, we investigated the effect of 1 on LPS-induced iNOS and COX-2 mRNA expressions using qRT-PCR. Our results showed that 1 significantly reduced the levels of LPS-induced iNOS and COX-2 mRNA (Figure 2D,E). We estimated the effects of 1 on the LPS-induced gene transcriptional levels of iNOS and COX-2 by measuring their promoter activities. Compound 1 significantly inhibited the iNOS and COX-2 promoter activities (Figure 2F), indicating that 1 might regulate LPS-induced iNOS and COX-2 at the level of gene transcription. Since TNF-α and IL-6 are important inflammatory cytokines secreted by macrophages,35 we examined the effect of 1 on LPSinduced TNF-α and IL-6 protein and mRNA expression by EIA and qRT-PCR, respectively. Pretreatment with 1 considerably reduced LPS-induced TNF-α and IL-6 production (75.0 and 51.3% at 60 μM) (Figure 3A,D) and their mRNA expressions (Figure 3B,E), indicating that 1 might regulate LPS-induced proinflammatory mediators at the level of gene transcription. Using a promoter assay, we revealed that 1 down-regulates TNF-α expression at the gene transcriptional level (Figure 3C). To determine whether the anti-inflammatory effects of 1 observed in RAW 264.7 macrophages also occur in primary cells, we examined the effect of 1 on the LPS-induced production of NO, PGE2, TNF-α, and IL-6 in peritoneal macrophages isolated from C57BL/6 mice. In these cells, 1 was found to inhibit the LPS-induced production of NO (55.9% at 60 μM), PGE2 (42.2% at 60 μM), and pro-inflammatory cytokines (TNF-α, 77.5% at 60 μM; IL-6, 61.1% at 60 μM) (Figure 4). In addition, the cytotoxic effects of 1 were evaluated using the MTT assay. Compound 1

Figure 1. Chemical structures of biflorin (1) and isobiflorin (2).

Compound 1 has also been found in plants other than S. aromaticum. Compound 1 was first isolated from the roots of Pancratium biflorum (Amaryllidaceae), and its structure was confirmed as a new polyoxygenated chromone-C-glucoside.34 Although the 1 isolated from S. aromaticum flower buds has been reported to possess an antibacterial effect,19 no report has been issued on its anti-inflammatory activities in activated macrophages or in murine inflammation models. As part of our ongoing screening program to evaluate the anti-inflammatory potential of natural compounds, we isolated 1 and 2 from cloves and compared their inhibitory effects on LPS-induced production of NO and PGE2 in macrophages. The object of this study is to evaluate the anti-inflammatory properties of 1 and to reveal the underlying molecular mechanisms in activated macrophages as models of inflammatory response. For establishing the in vivo anti-inflammatory characteristics of 1, the models of LPS-induced sepsis, carrageenan-induced paw edema, and croton-oil-induced ear edema were used. B

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Figure 2. Effects of 1 or 2 on LPS-induced NO and PGE2 production, iNOS and COX-2 expression, and iNOS and COX-2 promoter activities in RAW 264.7 macrophages. (A,B) Following pretreatment with 1 or 2 (15, 30, or 60 μM) for 1 h, cells were treated with LPS (10 ng/mL) for 24 h. NO and PGE2 production in the culture media were quantified using the Griess reaction assay and an EIA kit, respectively. Negative controls were not treated with LPS or test compound. Positive controls were L-NIL (20 μM) and NS-398 (3 μM) for NO and PGE2 production, respectively. (C) Lysates were prepared from cells pretreated with/without 1 (15, 30, or 60 μM) for 1 h and then with LPS (10 ng/mL) for 24 h. Total cellular proteins were resolved by SDS-PAGE, transferred to PVDF membranes, and detected with specific iNOS and COX-2 antibodies. β-Actin was used as an internal control. (D,E) Total RNA was prepared for qRT-PCR analysis of iNOS and COX-2 in cells stimulated with LPS (10 ng/mL) with/without 1 (60 μM) for 2, 4, 6, and 12 h. The mRNA levels of iNOS and COX-2 were determined using gene-specific primers as described in the Experimental Section. (F) Cells were transiently transfected with a pGL3-iNOS or a pGL3-COX-2 promoter vector, and a phRL-TK vector was used as an internal control. Cells were treated with/without 1 (60 μM) for 1 h and then stimulated with LPS (10 ng/mL) for 6 and 12 h. Cells were then harvested, and luciferase activity levels were determined as described in the Experimental Section. Data are presented as mean ± SD of three independent experiments. #p < 0.05 vs the control group; *p < 0.05, **p < 0.01, ***p < 0.001 vs LPS-stimulated cells.

did not affect the cell viability up to 60 μM for 24 h (cell viability = 90.36% at 60 μM) (Figure 4A). NF-κB, AP-1, and STAT1/3 are pivotal transcription factors regulating the expression of pro-inflammatory proteins and

cytokines as up-regulated by LPS.12,36 To investigate whether the inhibitory effects of 1 on these pro-inflammatory mediators involve these transcription factors, we examined the effect of 1 on LPS-induced NF-κB- and AP-1-dependent reporter gene C

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Figure 3. Effects of 1 on LPS-induced TNF-α and IL-6 protein production, mRNA expression, and TNF-α promoter activity in RAW 264.7 macrophages. (A,D) Cells were pretreated with/without 1 (15, 30, or 60 μM) for 1 h and then stimulated with LPS (10 ng/mL) for 24 h. Negative controls were not treated with LPS or 1. TNF-α and IL-6 levels were determined by using enzyme immunoassay kits. (C) Cells were transiently transfected with pTNF-α-luc reporter; phRL-TK vector was used as an internal control. Cells were pretreated with/without 1 (60 μM) for 1 h and then stimulated with LPS (10 ng/mL) for 6 and 12 h. Cells were then harvested, and luciferase activity levels were determined as described in the Experimental Section. (B,E) Total RNA was prepared for qRT-PCR analysis of TNF-α and IL-6 in cells stimulated with LPS (10 ng/mL) with/without 1 (60 μM) for 2, 4, 6, and 12 h. The mRNA levels of TNF-α and IL-6 were determined using gene-specific primers as described in the Experimental Section. Negative controls were not treated with LPS or 1. Data are presented as mean ± SD of three independent experiments. #p < 0.05 vs the control group; **p < 0.01, ***p < 0.001 vs LPS-stimulated cells.

suppressed in response to interferon-γ (IFN-γ) by decreasing the histone acetylation of promoter.39 Compound 1 markedly reduced LPS-induced STAT1 phosphorylation at both Y701 and S727 in macrophages (Figure 5C,D), suggesting that 1 inhibits STAT1 activation. Since certain naphthoquinones manifest their anticancer and anti-inflammatory properties via inactivation of STAT3,40,41 we also tested the effects of 1 on LPS-induced STAT3 phosphorylation. As shown Figure 5C,D, compound 1 slightly inhibited the LPS-induced phosphorylation of STAT3 (Y705) in LPS-induced RAW 264.7 and peritoneal macrophages. Because

activities. Analysis of reporter gene expression using pNF-κB-luc or pAP-1-luc demonstrated that 1 did not have any inhibitory effects on LPS-induced NF-κB and AP-1 luciferase activity in RAW 264.7 macrophages (Figure 5A,B). STAT1 activation is achieved by tyrosine and serine phosphorylation at particular residues.37 Phosphorylation at Y701 by receptor tyrosine kinases causes the STAT1 dimers to accumulate in the nucleus and bind DNA. For full transcriptional activity and biological function, STAT1 must also be phosphorylated on S727.38 In macrophages expressing mutant STAT1S727A, STAT1-regulated expression of genes is markedly D

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Figure 4. Effects of 1 on LPS-induced NO, PGE2, TNF-α, and IL-6 production in peritoneal macrophages. (A−D) Cells were isolated from 5% thioglycolate-elicited C57BL/6 mice. Following pretreatment with/without 1 (15, 30, or 60 μM) for 1 h, cells were treated with LPS (10 μg/mL) for 48 h. Levels of NO, PGE2, TNF-α, and IL-6 in culture media were quantified using the Griess reaction and EIA kits. Negative controls were not treated with LPS or 1. L-NIL (20 μM) and NS-398 (3 μM) were used as the positive controls for NO and PGE2 production, respectively. Experiments were repeated three times, and similar results were obtained. Data are presented as mean ± SD of three independent experiments. #p < 0.05 vs the control group; **p < 0.01, ***p < 0.001 vs LPS-stimulated cells.

The demonstrated ability of 1 to attenuate LPS-induced iNOS, COX-2, and pro-inflammatory cytokine levels in macrophages prompted an assessment of the anti-inflammatory effects of 1 in an animal model of sepsis. Sepsis is characterized by a severe inflammatory response to infection with multiple physiologic and immunologic abnormalities.46,47 As LPS is the most frequent cause of sepsis, it has been shown that LPS-stimulated progressive production of pro-inflammatory mediators, including NO, IL-6, TNF-α, and IL-1β, plays an important role in multiple organ failure and death during endotoxemia.48 Because LPS administration to mice induces systemic inflammation that mimics many of the initial clinical features of sepsis, the LPSinduced sepsis model is useful for evaluating the effects of candidate anti-inflammatory compounds.49 LPS administration (25 mg/kg, i.p.) markedly increased mRNA levels of iNOS, COX-2, and TNF-α in liver tissue, while pretreatment with 1 (5 or 10 mg/kg, i.p.) 1 h before LPS administration significantly decreased the mRNA levels of iNOS, COX-2, and TNF-α (Figure 7A). In addition, 1 markedly reduced the LPS-induced phosphorylation of STAT1 at Y701 and S727 (Figure 7B), suggesting that 1 inhibited STAT1 activation in septic mice as well as macrophages. STAT1 rapidly undergoes phosphorylation in a model of LPS hypersensitivity in vivo, and importantly, genetic ablation of STAT1 protects against LPS-induced lethality, suggesting that STAT1 promotes inflammatory responses in LPS-induced endotoxemia.16 As shown in Figure 7C, LPS administration resulted in 100% mortality at 48 h postinjection, but pretreatment with 1 (5 or 10

phosphorylation of STATs is required for their transcriptional activities,15 our results demonstrated that STAT1 rather than STAT3 might be the main target for anti-inflammatory activity of 1. Several mitogen-activated protein kinases (MAPKs), including p38 MAPK, c-Jun N-terminal kinase (JNK), and extracellular signal-regulated kinase (ERK) 1/2 are involved in the signal transduction pathways that lead to up-regulation of inflammatory mediators. Since these pathways play critical roles in the transcriptional activation of STATs,42 we investigated whether the inhibitions of inflammatory responses by 1 are mediated through the MAPK pathway in LPS-induced RAW 264.7 macrophages. As shown in Figure 6, LPS significantly increased phosphorylation of all MAPKs at 30 min. Compound 1 pretreatment suppressed the LPS-induced phosphorylation of p38 MAPK in a concentration-dependent manner. However, 1 did not affect the phosphorylation of ERK1/2 or JNK. These results were consistent with studies reporting that p38 MAPK was necessary for S727 phosphorylation of STAT1.43,44 It was reported that LPS increases phosphorylation of STAT1 at S727, independently of phosphorylation of Y701, and that p38 MAPK inhibitor (SB203580 and SB202190) abolished the LPS-induced S727 phosphorylation of STAT1 along with induction of iNOS.43,45 Compound 1-mediated inhibition of STAT1 and the p38 MAPK pathway provides an insight into the mechanism by which 1 inhibits LPS-induced pro-inflammatory mediator release. E

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Figure 5. Effects of 1 on LPS-induced NF-κB and AP-1 activity and STAT1/3 activation in RAW 264.7 macrophages and peritoneal macrophages. (A,B) Cells were transiently transfected with pNF-κB-luc reporter or pAP-1-luc reporter; phRL-TK vector was used as an internal control. Cells were pretreated with/without the indicated concentrations of 1 for 1 h and then stimulated with LPS (10 ng/mL) for 12 h. Cells were then harvested, and luciferase activity levels were determined as described in the Experimental Section. (C) RAW 264.7 macrophages were pretreated with/without 1 (15, 30, or 60 μM) for 1 h and then stimulated with LPS (10 ng/mL) for p-STAT1 (120 min for Y701 and 15 min for S727) or p-STAT3 (120 min for Y705). (D) Peritoneal macrophages were pretreated with/without 1 (15, 30, or 60 μM) for 1 h and then stimulated with LPS (10 μg/mL) for p-STAT1 (120 min for Y701 and 30 min for S727) or p-STAT3 (60 min for Y705). Total cellular proteins were resolved by SDS-PAGE, transferred to PVDF membranes, and detected using specific p-STAT1 (Y701 and S727), p-STAT3 (Y705), and STAT1 and STAT3 antibodies. β-Actin was used as an internal control. The experiment was repeated three times, and similar results were obtained. Data are presented as mean ± SD of three independent experiments. #p < 0.05 vs the control group; *p < 0.05, **p < 0.01, ***p < 0.001 vs LPS-stimulated cells.

results suggest that 1 has anti-inflammatory potency in various inflammatory animal models. In summary, our findings show that biflorin inhibits the LPSinduced production of NO, PGE2, and inflammatory cytokines by blocking the activation of STAT1 and p38 MAPK in macrophages. More importantly, biflorin decreases the expression of inflammatory mediators in liver tissue in vivo and protected mice from LPS-induced lethality. Our in vivo results demonstrate that biflorin inhibits the carrageenan-induced or croton-oil-induced edema in rat models of acute inflammation. Thus, our findings indicate that biflorin can be considered for evaluation as a potential treatment option for inflammatory diseases.

mg/kg, i.p.) reduced this lethality to 60 or 40%, respectively, at 96 h postinjection. These results indicate that 1 was protective against LPS-induced endotoxemia and that its effect might be via STAT1 inactivation. To verify whether 1 could ameliorate acute inflammatory symptoms in vivo, we employed a rat model of acute inflammatory paw edema induced by carrageenan. As expected, pretreatment with 1 (5, 10, or 15 mg/kg, i.p.) 1 h before carrageenan injection inhibited carrageenan-induced paw swelling. At 3 h, 1 reduced paw edema by 26, 48, and 60% (5, 10, and 15 mg/kg, i.p., respectively) compared to the carrageenan-only group. The positive control, ibuprofen (100 mg/kg, p.o.), decreased the edema by 68% at 3 h (Figure 8A). In order to further evaluate the anti-inflammatory activity of 1 in vivo, croton-oil-induced ear edema was studied (Figure 8B). Pretreatment with 1 (5, 10, or 15 mg/kg, i.p.) significantly reduced the ear thickness caused by croton oil application. The inhibition rates by 1 (5, 10, or 15 mg/kg, i.p.) were 33.36, 44.52, or 74.67%, respectively, compared to the croton-oil-only group. In addition, the positive control, ibuprofen (100 mg/kg, p.o.), effectively inhibited ear edema (78.19%). To examine histopathological changes during ear inflammation, cross sections of ear discs were stained with hematoxylin and eosin. Intense inflammatory changes, such as the development of ear edema and inflammatory cell infiltration, were caused by croton oil. Pretreatment with 1 (10 mg/kg, i.p.) significantly inhibited croton-oil-induced inflammatory changes (Figure 8C). Edema is the typical feature of inflammation not only in systemic inflammation but also in local inflammation.50 Therefore, these

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EXPERIMENTAL SECTION

Plant Material. The flower buds of S. aromaticum L. were obtained from a domestic Korean market (Kyungdong Crude Drugs Market, Seoul, South Korea) in June 2013. The moisture content of the plant material was 8.60% by oven-drying method at 105 °C for 7 h. The origin of the herbal material was identified by Prof. Dae Sik Jang, and a voucher specimen (SYAR1-2013) has been deposited in the Laboratory of Natural Product Medicine, College of Pharmacy, Kyung Hee University, Republic of Korea. Extraction and Isolation. The extraction scheme for 1 and 2 is shown in Supporting Information Figure S1. The dried and powdered plant material (2.1 kg) was extracted twice with 20 L of 70% EtOH in a 60 °C water bath for 2 h, and the solvent was evaporated in vacuo at 40 °C. The 70% EtOH extract (625.0 g) was suspended in H2O (2 L) and successively extracted with n-hexane (3 × 2 L), EtOAc (3 × 2 L), and BuOH (3 × 2 L) to give n-hexane- (303.1 g), EtOAc- (132.0 g), BuOHF

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(EIA) kits for PGE2, TNF-α, and IL-6 were obtained from R&D Systems (Minneapolis, MN, USA). Random oligonucleotide primers and MMLV reverse transcriptase were purchased from Promega (Madison, WI, USA). The dNTP mix and SYBR green ex Taq were obtained from TaKaRa (Shiga, Japan). iNOS, COX-2, TNF-α, IL-6, and β-actin oligonucleotide primers were purchased from Bioneer (Seoul, Korea). 3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), sulfanilamide, aprotinin, leupeptin, phenylmethylsulfonyl fluoride (PMSF), DL-dithiothreitol (DTT), L-N6-(1-iminoethyl)lysine (L-NIL), N-(2-cyclohexyloxy-4-nitrophenyl)methanesulfonamide (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. The RAW 264.7 macrophage cell line was obtained from the Korea Cell Line Bank (Seoul, Korea). These cells were grown at 37 °C in DMEM supplemented with 10% FBS, penicillin (100 units/mL), and streptomycin sulfate (100 μg/ mL) in a humidified atmosphere of 5% CO2. Murine peritoneal macrophages were elicited by i.p. injection of 3 mL of 4% thioglycolate into the peritoneal cavity of 6−10 week old C57BL/6 male mice. After 4 days, cells in the peritoneal exudates were obtained by lavage with icecold DMEM. The cells were washed twice, resuspended in DMEM (supplemented with 10% FBS, 100 units/mL penicillin, and 100 μg/mL streptomycin sulfate) and were seeded in sterile disposable culture plates. Cells were incubated with the tested samples at increasing concentrations (15, 30, or 60 μM) or an appropriate positive control for 1 h and then stimulated with LPS (RAW 264.7 macrophages: 10 ng/mL or peritoneal macrophages; 10 μg/mL) for the indicated time. MTT Assay. Cell viability was assessed by the MTT assay. RAW 264.7 macrophage cells were seeded at 1 × 105 cells/mL in 96-well plates containing 100 μL of DMEM medium with 10% FBS and incubated overnight. After overnight incubation, 1 or 2 was added and the plates were incubated for 24 h. The cells were then incubated with a MTT solution [(3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide, 5 mg/mL stock solution in PBS] for 4 h at 37 °C under 5% CO2. The medium was discarded, and the formazan blue that formed in the cells was dissolved in 200 μL of DMSO. Absorbance of each well was measured at 540 nm using a microplate reader (Molecular Devices Inc., CA, USA). Nitrite Determination. Nitrite levels in culture media were determined using the Griess assay51 and presumed to reflect NO levels. Cells were plated at 2 × 105 cells/mL (RAW 264.7 macrophages) or 5 × 105 cells/mL (peritoneal macrophages) in 24-well plates and incubated overnight. Following treatment with various concentrations of 1 or 2 for 1 h, cells were treated with LPS (10 ng/mL; RAW 264.7 macrophages or 10 μg/mL; peritoneal macrophages) for 24 h. The iNOS inhibitor L-NIL served as a positive control. Cell culture medium (100 μL) was mixed with 100 μL of Griess reagent [equal volumes of 1% (w/v) sulfanilamide in 5% (v/v) phosphoric acid and 0.1% (w/v) naphthylethylenediamineHCl], incubated at room temperature for 10 min, and then the absorbance at 540 nm was determined in a microplate reader (Molecular Devices Inc., Sunnyvale, CA, USA). The amount of nitrite in the samples was determined by using NaNO2 as a standard. PGE2, TNF-α, and IL-6 Assays. Cells were plated at 2 × 105 cells/ mL (RAW 264.7 macrophages) or 5 × 105 cells/mL (peritoneal macrophages) in 24-well plates and incubated overnight. Following treatment with various concentrations of 1 or 2 for 1 h, cells were treated with LPS (10 ng/mL; RAW 264.7 macrophages or 10 μg/mL; peritoneal macrophages) for 24 h. Dilutions of the cell culture medium were assayed for PGE2, TNF-α, and IL-6. PGE2 levels using a colorimetric competitive enzyme-linked immunosorbent assay (ELISA) kit (Enzo Life Science, NY, USA) according to the manufacturer’s instructions. TNF-α and IL-6 levels in cell culture medium were quantified by ELISA using mouse DuoSet kit (R&D Systems, MN, USA) according to the manufacturer’s instructions. Western Blot Analysis. Protein extracts were isolated from the liver tissue of septic mice and macrophages using PRO-PREP protein extraction solution (Intron Biotechnology, Seoul, Korea) supplemented with protease inhibitor cocktail (Sigma, MO, USA) and incubated for 30 min at 4 °C. Debris was removed by microcentrifugation, followed by

Figure 6. Effects of 1 on LPS-induced MAPKs activation in RAW 264.7 macrophages. Lysates were prepared from cells pretreated with/without 1 (15, 30, or 60 μM) for 1 h and then with LPS (10 ng/mL) for 30 min. Total cellular proteins were resolved by SDS-PAGE, transferred onto PVDF membranes, and detected using specific p-p38, p38, p-JNK, JNK, p-ERK, and ERK antibodies. β-Actin was used as an internal control. The experiment was repeated three times, and similar results were obtained. Data are presented as mean ± SD of three independent experiments. #p < 0.05 vs the control group; ***p < 0.001 vs LPSstimulated cells. (93.0 g), and water-soluble extracts (95.7 g). The BuOH-soluble fraction (93.0 g) was chromatographed over Diaion HP-20 (ϕ 7.7 × 67 cm) as stationary phase eluting with H2O−MeOH gradient [1:0 (2.0 L), 4:1 (2.0 L), 3:2 (4.0 L), 2:3 (3.0 L), 1:4 (2.0 L), 0:1 (2.0 L)] to afford nine pooled fractions (B01−B09). Fraction B04 [eluted with H2O− MeOH (2:3 v/v), 6.4 g] was further fractionated using Sephadex LH-20 (ϕ 4.5 × 55 cm) with 60% acetone to give five subfractions (B04-1− B04-5). Compound 1 (931.6 mg) was purified from fraction B04-4 (3.24 g) by recrystallization in MeOH. Fraction B03 [eluted with H2O− MeOH (3:2 v/v), 24.5 g] was chromatographed over Diaion HP-20 (ϕ 4.5 × 55 cm) eluting with H2O−MeOH gradient (3:7 v/v) to afford six subfractions (B03-1−B03-6). Compound 2 (667.8 mg) was isolated from fraction B03-4 (6.50 g) by recrystallization in MeOH. The chemical structures of 1 and 2 were identified to be biflorin and isobiflorin, respectively, by 1H and 13C NMR spectroscopic data analysis and by comparison with published values.33 The purity of both 1 and 2 used in these studies was higher than 97% from the peak area in the LC/ MS chromatogram. Chemicals. Dulbecco’s modified Eagle’s medium (DMEM), fetal bovine serum (FBS), and penicillin and streptomycin were obtained from Life Technologies Inc. (Grand Island, NY, USA). iNOS, COX-2, p38, p-JNK, p-ERK, JNK, ERK, STAT1, STAT3, and β-actin monoclonal antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA). p-p38, p-STAT1(Y701), p-STAT1(S727), and p-STAT3(Y705) monoclonal antibodies were purchased from Cell Signaling Technologies (Beverly, MA, USA). HPR-conjugated antimouse, anti-rabbit, and anti-goat IGg were purchased from Jackson Immunoresearch (West Grove, PA, USA). The enzyme immunoassay G

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Figure 7. Effects of 1 on LPS-induced endotoxin model. (A,B) Eight mice per group were injected with vehicle only or 1 (10 mg/kg, i.p.) and 1 h prior to LPS injection (25 mg/kg, i.p.). Six hours after LPS injection with/without 1, liver tissue was collected to determine mRNA and protein expression. Total cellular proteins were resolved by SDS-PAGE, transferred to PVDF membranes, and detected with specific antibodies. Data are presented as mean ± SD (n = 8). #p < 0.05 vs the control group; ***p < 0.001 vs LPS-stimulated mice. (C) Male C57BL/6 mice (6 weeks) were injected with 1 (5 or 10 mg/kg, i.p.) or vehicle 1 h before LPS injection (25 mg/kg, i.p.). Survival of mice was monitored every 12 h following LPS injection (n = 5). quick freezing of the supernatants. The protein concentration was determined using the Bio-Rad protein assay reagent according to the manufacturer’s instructions. Cellular protein extracts were mixed with 5× SDS sample buffer, boiled for 5 min, and then were separated by electrophoresis on a 10−12% SDS polyacrylamide gel and were electroblotted onto a PVDF membrane. The membrane was incubated for 1 h with blocking solution (5% skim milk) at room temperature, followed by incubation overnight with a primary antibody at 4 °C (1:1000). Blots were washed three times with Tween 20/Tris-buffered saline (T/TBS) and incubated with a 1:2000 dilution of horseradish peroxidase conjugated secondary antibody for 2 h at room temperature. Blots were again washed three times with T/TBS and developed using an ECL chemiluminescence substrate (Santa Cruz, CA, USA). Autoradiographs were obtained by LAS-4000 luminescent image analyzer (Fujifilm, Tokyo, Japan). Biorad Quantity One Software was used for the densitometric analysis. Quantitative Real-Time RT-PCR (qRT-PCR). Total RNA was isolated from the liver tissue of septic mice and macrophages using by Easy Bluekits (Intron Biotechnology, Seoul, Korea). RNA (1 μg) was reverse-transcribed (RT) using MuLV reverse transcriptase, 1 mM deoxyribonucleotide triphosphate (dNTP), and 0.5 mg/mL random

primer. PCR amplification was performed using the incorporation of SYBR green using SYBR Premix Ex Taq (Takara, Shiga, Japan). The PCR primers used in this study are listed below and were purchased from Bioneer (Seoul, Korea): for iNOS designed from mouse, CAT GCT ACT GGA GGT GGG TG (forward) and CAT TGA TCT CCG TGA CAG CC (reverse); for COX-2 designed from mouse, GGA GAG ACT ATC AAG ATA GT (forward) and ATG GTC AGT AGA CTT TTA CA (reverse); for TNF-α designed from mouse, AGC ACA GAA AGC ATG ATC CG (forward) and CTG ATG AGA GGG AGG CCA TT (reverse); for IL-6, GAG GAT ACC ACT CCC AAC AGA CC (forward) and AAG TGC ATC ATC GTT GTT CAT ACA (reverse). The oligonucleotide primers for β-actin used as a house-keeping gene designed from mouse were ATC ACT ATT GGC AAC GAG CG (forward) and TCA GCA ATG CCT GGG TAC AT (reverse). Steadystate mRNA levels were determined by real-time qPCR using the Takara thermal cycler device. A dissociation curve analysis of iNOS, COX-2, TNF-α, IL-6, and β-actin showed a single peak for each. Mean Ct values of genes of interest were calculated from triplicate measurements and normalized versus the mean Ct of β-actin. Plasmid, Transient Transfection, and Luciferase Assay. RAW 264.7 macrophages were cotransfected with pGL3-iNOS, pGL3-COXH

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Figure 8. Effects of 1 on carrageenan-induced paw edema and croton-oil-induced ear edema in rats. (A) 1 (5, 10, or 15 mg/kg, i.p.) was administered 1 h before carrageenan injection. Positive control animals were treated with ibuprofen (100 mg/kg, p.o.). The paw volume was measured at 1, 3, and 5 h after carrageenan injection. Values are the mean ± SD (n = 7); *p < 0.05, **p < 0.01, ***p < 0.001 vs the carrageenan-only group. (B) Ear edema was induced by croton oil as described in the Experimental Section. 1 (5, 10, or 15 mg/kg, i.p.) was injected 1 h before croton oil, and 6 mm ear punch was measured for thickness 3 h after croton oil application to assess edema. Values are the mean ± SD (n = 7); *p < 0.05, **p < 0.01, ***p < 0.001 vs the croton-oilonly group. (C) Histological analysis of croton-induced ear edema. Representative H&E sections of ear tissues were obtained from vehicle-treated control rat left ear (control), croton-oil-only rat right ear, 1 (10 mg/kg, i.p.) plus croton-oil-treated rat right ear, and ibuprofen (100 mg/kg, p.o.) plus croton-oil-treated rat right ear. In Vivo Paw Edema Model. The initial hind paw volume of the SD rats was determined volumetrically. Rats were treated with 1 (5 or 10 mg/kg, i.p.). After 1 h, a 1% solution of carrageenan in saline (0.1 mL per rat) was injected subcutaneously into right hind paws. Paw volumes were measured at 1, 3, and 5 h after injections, and edema volumes were measured using a plethymometer. Ibuprofen (100 mg/kg, p.o.), an antiinflammatory drug, was used as a positive control. In all experiments, we used seven animals per group. In Vivo Ear Edema Model. SD male rats were pretreated with 1 (5, 10, or 15 mg/kg, i.p.). After 1 h, ear edema was induced on the inner surface of the right ear by topical application of croton oil (5% solution in 100 μL of acetone). The left ear was used as a control and received the same amount of the vehicle (acetone). Three hours after croton oil application, rats were sacrificed using ethyl ether, and both ear tissues were collected using 6 mm punching. Ear punch biopsies were immediately measured for thickness to assess ear edema. In all experiments, we used seven animals per group. Ear biopsies were fixed in 4% paraformaldehyde overnight and embedded in paraffin. Ear biopsy sections were stained with hematoxylin and eosin (H&E) at the Seoul Medical Science Institute (SCL Co. Ltd., Seoul, Korea). Statistical Analysis. Results are presented as the mean ± SD of triplicate experiments. In the animal study, the data were expressed as the mean ± SD (n = 7−8). Statistically significant values were compared using ANOVA and Dunnett’s post-hoc test, and p values of less than 0.05 were considered statistically significant.

2, pTNF-α (Affymetrix, CA, USA), pNF-κB-Luc, or pAp-1-Luc (Clontech, Shiga, Japan) plasmid plus the phRL-TK plasmid (Promega, WI, USA) using Lipofectamine LTX (Invitrogen, CA, USA) as instructed by the manufacturers. After 6 h of transfection, cells were pretreated with 1 for 1 h and then stimulated with LPS (10 ng/mL) for 6 or 12 h. Each well was washed with cold PBS, and cells were lysed and the luciferase activity was determined using the Promega luciferase assay system (WI, USA). Animals. C57BL/6 male mice (5 weeks) and Sprague−Dawley (SD) male rats (5 weeks) were obtained from the Orient Bio Inc. (Seongnamsi, Korea) and maintained under constant conditions (temperature: 20− 25 °C, humidity: 40−60%, light/dark cycle: 12 h). At 12 h before the experiment, only water was provided. All procedures were conducted in accordance with university guidelines and approved by the ethical committee for Animal Care and the Use of Laboratory Animals, College of Pharmacy, Kyung Hee University. In Vivo Sepsis Model. Male C57BL/6 mice, 6−8 weeks old, were treated with 1 (5 or 10 mg/kg, i.p.). After 1 h, septic shock was induced by intraperitoneally injecting LPS (Salmonella enterica, 25 mg/kg). Survival was monitored for 96 h after LPS administration (n = 5). Liver tissues (n = 8) were obtained 6 h after LPS injection from each mouse and immediately frozen (−80 °C). The mRNA levels of iNOS, COX-2, and TNF-α were quantitated by qRT-PCR, and protein expression of pSTAT1, STAT1, p-STAT3, and STAT3 were detected by Western blot analysis. In all experiments, we used seven animals per group. I

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(22) Tanko, Y.; Mohammed, A.; Okasha, M. A.; Umah, A. H.; Magaji, R. A. Afr. J. Tradit., Complementary Altern. Med. 2008, 5, 209−12. (23) Dibazar, S. P.; Fateh, S.; Daneshmandi, S. J. Immunotoxicol. 2015, 12, 124−31. (24) Hong, C. H.; Hur, S. K.; Oh, O. J.; Kim, S. S.; Nam, K. A.; Lee, S. K. J. Ethnopharmacol. 2002, 83, 153−9. (25) Lee, G. I.; Ha, J. Y.; Min, K. R.; Nakagawa, H.; Tsurufuji, S.; Chang, I. M.; Kim, Y. Planta Med. 1995, 61, 26−30. (26) Bachiega, T. F.; de Sousa, J. P.; Bastos, J. K.; Sforcin, J. M. J. Pharm. Pharmacol. 2012, 64, 610−6. (27) Bensky, D.; Clavey, S.; StogerE.Chinese Herbal Medicine: Materia Medica; Eastland Press, 2004. (28) Ali, S. M.; Khan, A. A.; Ahmed, I.; Musaddiq, M.; Ahmed, K. S.; Polasa, H.; Rao, L. V.; Habibullah, C. M.; Sechi, L. A.; Ahmed, N. Ann. Clin. Microbiol. Antimicrob. 2005, 4, 20. (29) Kim, S. S.; Oh, O. J.; Min, H. Y.; Park, E. J.; Kim, Y.; Park, H. J.; Nam Han, Y.; Lee, S. K. Life Sci. 2003, 73, 337−48. (30) Ito, M.; Murakami, K.; Yoshino, M. Food Chem. Toxicol. 2005, 43, 461−6. (31) Al-Sharif, I.; Remmal, A.; Aboussekhra, A. BMC Cancer 2013, 13, 600. (32) Sarkar, A.; Bhattacharjee, S.; Mandal, D. P. Asian Pac. J. Cancer Prev. 2015, 16, 6753−9. (33) Zhang, Y.; Chen, Y. Phytochemistry 1997, 45, 401−403. (34) Yang, S. J.; Ryu, J. H.; Jang, D. S.; Yang, L.; Han, H. K. J. Pharm. Biomed. Anal. 2015, 115, 272−6. (35) Scheinecker, C.; Redlich, K.; Smolen, J. S. Immunity 2008, 28, 440−4. (36) Darnell, J. E., Jr.; Kerr, I. M.; Stark, G. R. Science 1994, 264, 1415− 21. (37) Kovarik, P.; Stoiber, D.; Novy, M.; Decker, T. EMBO J. 1998, 17, 3660−8. (38) Sadzak, I.; Schiff, M.; Gattermeier, I.; Glinitzer, R.; Sauer, I.; Saalmuller, A.; Yang, E.; Schaljo, B.; Kovarik, P. Proc. Natl. Acad. Sci. U. S. A. 2008, 105, 8944−9. (39) Varinou, L.; Ramsauer, K.; Karaghiosoff, M.; Kolbe, T.; Pfeffer, K.; Muller, M.; Decker, T. Immunity 2003, 19, 793−802. (40) Andujar, I.; Rios, J. L.; Giner, R. M.; Miguel Cerda, J.; Recio, M. C. Evid. Based Complement. Alternat. Med. 2012, 2012, 271606. (41) Aziz, M. H.; Dreckschmidt, N. E.; Verma, A. K. Cancer Res. 2008, 68, 9024−32. (42) Lim, C. P.; Cao, X. J. Biol. Chem. 1999, 274, 31055−61. (43) Kovarik, P.; Stoiber, D.; Eyers, P. A.; Menghini, R.; Neininger, A.; Gaestel, M.; Cohen, P.; Decker, T. Proc. Natl. Acad. Sci. U. S. A. 1999, 96, 13956−61. (44) Turkson, J.; Bowman, T.; Adnane, J.; Zhang, Y.; Djeu, J. Y.; Sekharam, M.; Frank, D. A.; Holzman, L. B.; Wu, J.; Sebti, S.; Jove, R. Mol. Cell. Biol. 1999, 19, 7519−28. (45) Huang, H.; Rose, J. L.; Hoyt, D. G. Mol. Pharmacol. 2004, 66, 302−11. (46) Ando, H.; Takamura, T.; Ota, T.; Nagai, Y.; Kobayashi, K. J. Pharmacol. Exp. Ther. 2000, 294, 1043−6. (47) Matsukawa, A.; Takeda, K.; Kudo, S.; Maeda, T.; Kagayama, M.; Akira, S. J. Immunol. 2003, 171, 6198−205. (48) Yang, G.; Ham, I.; Choi, H. Y. Food Chem. Toxicol. 2013, 58, 124− 32. (49) Doi, K.; Leelahavanichkul, A.; Yuen, P. S.; Star, R. A. J. Clin. Invest. 2009, 119, 2868−78. (50) Zhang, X.; Li, J.; Chen, C.; Ci, X.; Yu, Q.; Deng, X. Fundam. Clin. Pharmacol. 2011, 25, 700−7. (51) Griess, J. P. Philos. Trans. R. Soc. 1864, 154, 667−731.

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jnatprod.5b00609. Figure S1 (PDF)

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AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]. Tel: +82-2-9610719. Fax: +82-29663885. *E-mail: [email protected]. Tel.: +82-2-9610860. Fax: +82-29663885. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This research was supported by Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Science, ICT and future Planning (No. NRF-2015R1A2A2A01003459) and by the Ministry of Education (NRF-2013R1A1A2004398).

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REFERENCES

(1) Yamamoto, Y.; Gaynor, R. B. Trends Biochem. Sci. 2004, 29, 72−9. (2) Ryu, S.; Shin, J. S.; Cho, Y. W.; Kim, H. K.; Paik, S. H.; Lee, J. H.; Chi, Y. H.; Kim, J. H.; Lee, K. T. Biol. Pharm. Bull. 2013, 36, 467−74. (3) Boniface, K.; Bak-Jensen, K. S.; Li, Y.; Blumenschein, W. M.; McGeachy, M. J.; McClanahan, T. K.; McKenzie, B. S.; Kastelein, R. A.; Cua, D. J.; de Waal Malefyt, R. J. Exp. Med. 2009, 206, 535−48. (4) Murakami, M.; Kudo, I. Prog. Lipid Res. 2004, 43, 3−35. (5) Bendtzen, K. Immunol. Lett. 1988, 19, 183−91. (6) Mumm, J. B.; Oft, M. Oncogene 2008, 27, 5913−9. (7) Nambu, A.; Nakae, S.; Iwakura, Y. Int. Immunol. 2006, 18, 701−12. (8) Hodge, D. R.; Hurt, E. M.; Farrar, W. L. Eur. J. Cancer 2005, 41, 2502−12. (9) Shi, Q.; Cao, J.; Fang, L.; Zhao, H.; Liu, Z.; Ran, J.; Zheng, X.; Li, X.; Zhou, Y.; Ge, D.; Zhang, H.; Wang, L.; Ran, Y.; Fu, J. Int. Immunopharmacol. 2014, 20, 298−306. (10) Duffield, J. S. Clin. Sci. 2003, 104, 27−38. (11) Wang, C.; Deng, L.; Hong, M.; Akkaraju, G. R.; Inoue, J.; Chen, Z. J. Nature 2001, 412, 346−51. (12) Sakurai, H.; Nishi, A.; Sato, N.; Mizukami, J.; Miyoshi, H.; Sugita, T. Biochem. Biophys. Res. Commun. 2002, 297, 1277−81. (13) Lee, J.; Mira-Arbibe, L.; Ulevitch, R. J. J. Leukoc. Biol. 2000, 68, 909−15. (14) Qi, Z.; Yin, F.; Lu, L.; Shen, L.; Qi, S.; Lan, L.; Luo, L.; Yin, Z. Inflammation Res. 2013, 62, 845−55. (15) Schindler, C.; Levy, D. E.; Decker, T. J. Biol. Chem. 2007, 282, 20059−63. (16) Luu, K.; Greenhill, C. J.; Majoros, A.; Decker, T.; Jenkins, B. J.; Mansell, A. Immunol. Cell Biol. 2014, 92, 761−9. (17) Chapgier, A.; Boisson-Dupuis, S.; Jouanguy, E.; Vogt, G.; Feinberg, J.; Prochnicka-Chalufour, A.; Casrouge, A.; Yang, K.; Soudais, C.; Fieschi, C.; Santos, O. F.; Bustamante, J.; Picard, C.; de Beaucoudrey, L.; Emile, J. F.; Arkwright, P. D.; Schreiber, R. D.; Rolinck-Werninghaus, C.; Rosen-Wolff, A.; Magdorf, K.; Roesler, J.; Casanova, J. L. PLoS Genet. 2006, 2, e131. (18) Debjit, B.; Sampath, K.; Akhilesh, Y.; Shweta, S.; Shravan, P.; Amit Sankar, D. J. Pharmacogn. Phytochem. 2012, 1, 13−22. (19) Cai, L.; Wu, C. D. J. Nat. Prod. 1996, 59, 987−90. (20) Correa, A. P.; Daroit, D. J.; Velho, R. V.; Brandelli, A. Braz. J. Microbiol. 2011, 42, 1479−84. (21) Nm, J.; Rm, R.; G, G.; Maliakel, B.; D, S.; Im, K. Food Funct. 2015, 6, 3373−82. J

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