2018 Volume 41 Issue 8 Pages 1228-1236
The objective of this study was to investigate the effect of dibenzoylmethane (DBM) on monocyte-to-macrophage differentiation, the inflammatory response, and the resulting signaling in human monocytes and murine macrophage. DBM effectively inhibited the monocyte-to-macrophage differentiation induced by phorbol 12-myristate 13-acetate (PMA) through a reduction in adhesion of THP-1 cells. Cluster of differentiation molecule β (CD11β) and CD36, which are surface markers of macrophage differentiation, were downregulated by 80 and 74%, respectively. DBM also significantly inhibited lipopolysaccharide (LPS)-induced nitrite (NO) production through the downregulation of inducible oxide synthase (iNOS) in RAW264.7 cells. The abundance of cyclooxygenase-2 (COX-2), a pro-inflammatory protein, was also effectively decreased by DBM in a dose-dependent manner. DBM (50 µM) reduced the levels of COX-2 and iNOS by 81 and 78%, respectively. DBM significantly inhibited the translocation of nuclear factor kappa-light-chain-enhancer of activated B cells (NF-κB), an inflammatory transcription factor, into the nucleus. DBM-mediated increase of NF-κB translocation resulted from the DBM-induced suppression of the phosphorylation of nuclear factor of kappa light polypeptide gene enhancer in B-cells inhibitor, alpha (IκBα). In contrast, DBM effectively increased the expression of nuclear factor E2-related factor 2 (Nrf2) and its target protein, hemeoxygenase-1 (HO-1). Nrf2 translocation into the nucleus was also significantly enhanced by DBM. Furthermore, DBM effectively inhibited the expression of pro-inflammatory cytokines such as tumor necrosis factor alpha (TNF-α), interleukin-1 beta (IL-1β), IL-6, and monocyte chemoattractant protein-1 (MCP-1). These results indicated that the DBM-mediated differential regulation of NF-κB and Nrf2, which are major transcription factors involved in inflammation, inhibited the expression of inflammatory cytokines.
Inflammation is a highly adaptive immune response that protects the body from infection and damage.1,2) It is natural physiological response that plays a crucial role in the removal of harmful pathogens and the repair of damaged tissues.1,3) Despite the beneficial effects in the protection of the body, inappropriately prolonged or unresolved inflammation can lead to chronic inflammation. Chronic inflammation is known to be low-grade and systemic response that stealthily damages a wide range of tissues, including healthy cells, over a long period.4) The chronic inflammatory response is accompanied by the excessive or prolonged release of pro-inflammatory cytokines and mediators from immune cells and the chronic activation of the immune system.4) This dysregulation of pro-inflammatory cytokines in chronic inflammation has been implicated with various severe diseases, such as cardiovascular disease, cancer, diabetes, and obesity.3–6) Thus, the control of pro-inflammatory cytokines is a major aim in the prevention of chronic inflammation-mediated diseases. The production or release of pro-inflammatory cytokines is related to the activation of nuclear factor kappa-light-chain-enhancer of activated B cells (NF-κB), a transcription factor, which stimulates the gene expression of pro-inflammatory cytokines such as interleukins, inducible nitric oxide synthase (iNOS), and cyclooxygenase-2 (COX-2).7) The NF-κB-induced activation of pro-inflammatory mediators requires the degradation of nuclear factor of kappa light polypeptide gene enhancer in B-cells inhibitor, alpha (IκBα), an upstream regulator, which binds to NF-κB to form a complex in the toll-like receptor (TLR) signaling pathway.8,9) In recent years, nuclear factor E2-related factor 2 (Nrf2) signaling has emerged as a mechanistic system for the regulation of inflammation by exhibiting an anti-inflammatory response in opposition to NF-κB activation.10) The activation of Nrf2 and its target genes has been known to suppress the expression of various pro-inflammatory cytokines.10–12)
Atherosclerosis is a hazardous inflammatory disease.13) During its pathogenesis, circulating monocytes adhere to endothelial cells, the vessel wall, and differentiate into macrophages. This monocyte-to-macrophage differentiation is a typical event in atherosclerosis and contributes to the inflammatory environment.14,15) Macrophages actively participate in the inflammatory immune response and mediate the recruitment of inflammatory cytokines to form plaques in the artery wall.14)
Nonsteroidal anti-inflammatory drugs (NSAIDs) are commonly used to control inflammatory responses.16) However, the long-term use of NSAIDs causes side effects such as rashes, high blood pressure, intestinal bleeding, and kidney problems.17,18) Anti-inflammatory agents from natural plant sources have been proposed to offer an alternative solution.17,18)
Dibenzoylmethane (DBM) is a phenolic compound found in licorice, the root of Glycyrrhiza glabra, which belongs to a herbaceous legume native to Europe and Asia.19) Licorice root has been included in Chinese traditional treatments for various diseases, such as cold and liver disease, for thousands of years.20) In addition, it has been used as a food additive owing to its sweetness.21) DBM is a minor component derived from licorice, for which various biological activities have been reported, such as anti-mutagenesis, anti-tumorigenesis, and anti-estrogenic activity.22–24) Several studies have described the anti-inflammatory properties of DBM or its derivatives.25,26) However, most of them have simply reported partial effects or data on the anti-inflammatory capacity of DBM and did not investigate the responsible mechanisms. Systematic approaches on the anti-inflammatory effect of DBM have been rarely considered; the effect on monocyte-to-macrophage differentiation remains to be investigated. Thus, this study addressed the inhibitory effect of DBM on inflammatory responses and monocyte-to-macrophage differentiation in RAW264.7 and THP-1 cells, and showed that DBM regulated NF-κB and Nrf2 to produce anti-inflammatory responses.
Dibenzoylmethane (DBM) was purchased from Fluka (Fluka/Sigma-Aldrich, Germany). Dulbecco’s modified Eagle’s medium (DMEM) and insulin were purchased from Gibco (Gaithersburg, MD, U.S.A.). Fetal bovine serum (FBS), penicillin-streptomycin (PS), and phosphate-buffered saline (PBS) were obtained from Hyclone (Logan, UT, U.S.A.). Escherichia coli lipopolysaccharide (LPS) was purchased from Sigma-Aldrich Chemical Co. (St. Louis, MO, U.S.A.). 3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) was purchased from Amresco Inc. (Solon, OH, U.S.A.). Trypsin–ethylenediaminetetraacetic acid (EDTA) Solution (0.25%) was obtained from WELGNE Inc. (Daegu, Korea). Trypan Blue Solution (0.4%) was purchased from Thermo Fisher Scientific (Waltham, MA, U.S.A.). Griess reagent was purchased from Sigma-Aldrich Chemical Co. Antibodies for the cluster of differentiation molecule β (CD11β), CD36, iNOS, COX-2, IκBα, p-IκBα, and NF-κB proteins were purchased from Cell Signaling Technology, Inc. (Danvers, MA, U.S.A.). Nrf2, Keap1, heme oxygenase-1 (HO-1), Lamin B, and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) were obtained from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA, U.S.A.). All other chemicals were purchased from Sigma-Aldrich Chemical Co.
Cell CultureRAW264.7 cells was cultured in DMEM supplemented with 1.5 g/L sodium bicarbonate, 1% PS, and 10% FBS. The cells were seeded at a density of 2×106 cells/well in a 6-well plate. When the confluence reached 80%, DBM or LPS (1 µg/mL) was applied. THP-1 human monocytes were cultured in RPMI-1640 supplemented with 10% FBS and 1% PS. Differentiation was induced by the addition of 50 ng/mL phorbol 12-myristate 13-acetate (PMA) for 24 h. DBM was generally pretreated 2 h before stimulation of LPS or PMA to investigate protective effect of DBM (Fig. 1). Both cell lines were maintained at 37°C in an atmosphere with 5% CO2 and 95% humidity. The morphological activation and differentiation of the two cell lines was observed by using a microscope (Leica DM2500, Wetzlar, Germany).
THP-1 cells were treated with various concentrations of Dibenzoylmethane for 2 h prior to 50 ng/mL PMA stimulation after which the cells were incubated for 48 h. Cells were washed three times with PBS to remove non-adherent cells. Adherent cells were observed under microscope and were detached with 0.25% Trypsin–EDTA Solution. Cell suspension was diluted 1 : 1 using a 0.4% Trypan Blue solution. Cell count was performed using hemocytometer and microscope.
Cell Viability AssayTo analyze cell viability, an MTT assay was performed. Between 1×104 and 1×105 cells/well of RAW264.7 or LPS-treated RAW264.7 cells were seeded in a 96-well plate. After incubation for 24 h, the cells were treated with DBM for 24 h. The medium was removed and MTT reagent (0.5 mg/mL) in serum-free media was added for 45 or 60 min at 37°C. MTT formazan was then solubilized in dimethyl sulfoxide (DMSO). The absorbance was detected at 550 nm (Spectra Max M3; Molecular Device, Sunnyvale, CA, U.S.A.). THP-1 cells or PMA-treated tHP-1 cells were seeded at 2×104 cells/well in 96-well plates. The cells were incubated with DBM or vehicle (DMSO) for 48 h and then incubated with 12-(4-iodophenyl)-3-(4-nitrophenyl)-5-(2,4-disulfophenyl)-2H-tetrazolium (WST-1) reagent at 37°C for 1 h; after this, the formed formazan dye was solubilized and the color absorption determined at 450 nm (Spectra Max M3; Molecular Devices).
NO AssayBetween 2×106 and 2.5×106 cells/well (RAW264.7 cells) were seeded in 6-well plates. After incubation for 24 h, DBM was administered 2 h prior to the addition of LPS, which was incubated for a further 24 h; subsequently, 100 µL of media was transferred to 96-well plate and 100 µL of Griess reagent was added. The plate was incubated at 25°C in the dark for 5 min. The absorbance at 550 nm was measured by using a spectrophotometer (Spectra Max M3; Molecular Devices).
Protein Determination and Western BlottingThe cells were washed with PBS and collected by scraping with lysis buffer containing protease/phosphatase inhibitors. The cell was disrupted by pipetting on ice for 10 min. The protein lysates were centrifuged at 13000 rpm for 5 min in 4°C. The supernatant (protein extract) was determined by using the bicinchoninic acid (BCA) assay (Thermo Fisher Scientific) in accordance with the manufacturer’s protocol. Protein extracts (50 µg) were separated on 10% or 12% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to polyvinylidene difluoride (PVDF) membranes overnight. The membrane was incubated with blocking buffer (5% non-fat dried milk) for 30 min. The primary antibodies, appropriately diluted in blocking buffer, were probed overnight and washed twice with TBST buffer. Subsequently, the membrane was incubated with secondary antibody (1 : 5000) for 2 h. After multiple washes with TBST buffer, enhanced chemiluminescence (ECL) solution was applied and the bands were visualized by using an LAS 4000 imager (FUJIFILM, NY, U.S.A.). The quantification of the protein bands was computed by using ImageJ program (NIH, National Institutes of Health, Bethesda, MD, U.S.A.).
Separation of Cytosolic and Nuclear FractionsThe cultured RAW264.7 cells were washed twice with PBS, scraped, and resuspended in buffer [10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) (pH 7.9), 50 mM NaCl, 0.5 M sucrose, 0.1 mM EDTA, 1 mM dithiothreitol, 10 mM tetrasodium pyrophosphate, 100 mM NaF, 17.5 mM β-glycophosphate, 1 mM phenylmethylsulfonyl fluoride (PMSF), 4 µg/mL aprotinin, and 2 µg/mL pepstatin A]. The cell lysates were disrupted by pipetting on ice for 10 min and centrifuged at 1500 rpm for 10 min. The cytoplasmic fraction was separated as the supernatant, and stored at −80°C. The pellet was resuspended in buffer [10 mM HEPES, 10 mM KCl, 0.1 mM EDTA, 0.1 mM ethylene glycol bis(2-aminoethyl ether)-N,N,N′,N′-tetraacetic acid (EGTA), 1 mM PMSF, 4 µg/mL aprotinin, and 2 µg/mL pepstatin A]. The suspension was centrifuged at 1500 rpm for 5 min and the supernatant was removed. The pellet was vortexed for 10 min in buffer [0.1% NP-40, 10 mM HEPES, 10 mM KCl, 0.1 mM EDTA, 0.1 mM EGTA, 1 mM PMSF, 4 µg/mL aprotinin, and 2 µg/mL pepstatin A] and centrifuged at 14000 rpm for 10 min in 4°C. The obtained supernatant was the nuclear fraction and stored at −80°C.
Real-Time PCRRAW264.7 cells, treated with or without DBM, were washed with PBS. TRIzol reagent (Invitrogen, Carlsbad, CA, U.S.A.) was added to extract total RNA from the cells in accordance with the manufacturer’s protocol. Total RNA was determined by Nanodrop (Thermo Scientific). cDNA was synthesized from extracted total RNA (1 µg) by using a LaboPass cDNA synthesis kit (Cosmogenetech Co., Ltd., Seoul, Korea). Transcribed cDNA and targeted primers (Table 1) were mixed with SYBR Green PCR Master Mix (Applied Biosystems). PCR was performed in triplicate for each pair of primers by using AriaMx Real-Time PCR system (Agilent Technologies).
| Name | Forward (5′ to 3′) | Reverse (5′ to 3′) |
|---|---|---|
| TNF-α | AGCCCCCAGTCTGTATCCTT | CTCCCTTTGCAGAACTCAGG |
| IL-1β | ACTCATTGTGGCTGTGGAGA | TTGTTCATCTCGGAGCCTGT |
| IL-6 | CTGGGGATGTCTGTAGCTCA | CTGTGAAGTCTCCTCTCCGG |
| MCP-1 | GCTGCTACTCATTCACCAGC | CTTCTTGGCCTTGGTTTCCC |
| GAPDH | CTGCGACTTCAACAGCAACT | GAGTTGGGATAGGGCCTCTC |
Statistical analyzes were performed using the Statistical Package for Social Sciences version 12.0 (SPSS Inc., Chicago, IL, U.S.A.). Differences among groups were evaluated by one-way ANOVA and Tukey’s multiple range test. All data are expressed as means±standard error (S.E.M.). Student’s t-test was also used to analyze differences, where appropriate, and a p<0.05 was taken to indicate significance.
In THP-1 and RAW264.7 cells, 100 µM DBM did not affect the cell viability, regardless of PMA addition, but 200 µM DBM showed significant cell cytotoxicity (a 35% decrease in cell viability) (Fig. 2B). In RAW264.7 cells, LPS treatment led to a small reduction of cell viability in the absence of DBM treatment, with a 15% decrease compared with the control (Fig. 2C). The co-treatment of DBM and LPS resulted in a slightly higher cell viability than that of the control, demonstrating that DBM was not cytotoxic below 100 µM (Fig. 2C). However, similar to THP-1 cells, 200 µM DBM was cytotoxic, with a 24% reduction in cell viability. Therefore, subsequent experiments were performed with a DBM concentration of less than 100 µM.
THP-1 or RAW264.7 cells were treated with DBM (A) or vehicle (DMSO) for 24 or 48 h. The cell viability was examined by a WST-1 assay (B) and an MTT assay (C) for THP-1 and RAW264.7 cells, respectively. All assays were performed in triplicate and the results are presented as the mean and standard error of the mean (mean±S.E.M.). Different letters indicate statistically significant differences among the groups (p<0.05). Uppercase and lowercase letters represent statistical expressions for without PMA/LPS and with PMA/LPS, respectively. Symbols indicate significant differences at * p<0.05 between without PMA/LPS and with PMA/LPS by Student’s t-tests. n.s: not significnat.
To investigate the effect of DBM on THP-1 differentiation, cell adhesion and the protein abundance of CD11β/CD36 were examined. PMA, a monocyte-to-macrophage differentiation inducer, significantly increased the adherence of THP-1 cells to the plate compared with the control group (Fig. 3A). The adherence of THP-1 cells in the presence of PMA was five times greater than that of the control (Fig. 3B). However, the PMA-induced adhesion of THP-1 was decreased by DBM in a dose-dependent manner. At 50 µM DBM, adhesion was decreased by more than 50% compared with the group treated with only PMA (Fig. 3B). This result showed that DBM effectively inhibited the cell adhesion of THP-1, which is a feature of monocyte-to-macrophage differentiation. In addition, the PMA induction of CD11β and CD36, which are monocyte-to-macrophage differentiation protein markers, was decreased by DBM treatment in a dose-dependent manner (Fig. 3C). The high dose of DBM (50 µM) reduced the level of CD11β and CD36 proteins by 80% and 74%, respectively (Fig. 3C). These results showed that DBM effectively inhibited monocyte-to-macrophage differentiation of THP-1 cells.
THP-1 cell differentiation was induced by treatment with PMA. Adherent cells after PMA treatment were observed (A) and counted (B) by using a microscope. Protein lysates were prepared for Western blotting for the indicated antibodies of differentiation markers and GAPDH (C). The protein expression of CD11β and CD36 was quantified by using ImageJ software (D). The values are presented as means and standard error of the mean (mean±S.E.M.). Different letters indicate statistically significant differences among groups (p<0.05). CON: no treatment, PMA: only PMA treated. Three replicates of each experiment were performed.
LPS treatment markedly increased the NO production by over 10-fold increase compared with the untreated group (CON) (Fig. 4A). This result showed that LPS effectively induced inflammatory responses in RAW264.7 cells. DBM treatment significantly suppressed LPS-induced NO production in a dose-dependent manner (Fig. 4A). The high dose of DBM suppressed LPS-induced NO production by over 70% compared with the LPS group (Fig. 4A). These results showed that DBM effectively inhibited NO production, part of the inflammatory response, in RAW264.7 cells. This DBM-mediated inhibition of NO production was correlated with the morphological observation of cells (Fig. 4B). LPS caused a change in cell shape, from round to spiky, but this was suppressed by DBM treatment (Fig. 4B). The protein levels of iNOS, a NO synthesis enzyme, and COX-2, an additional pro-inflammatory marker, were determined after DBM treatment. iNOS and COX-2 proteins were almost undetectable in LPS-untreated RAW264.7 cells. However, LPS treatment effectively induced the expression of iNOS and COX-2 (Fig. 4C), confirming that inflammatory responses were induced by LPS treatment. At 50 µM, DBM decreased the LPS-induced increase in both iNOS and COX-2 protein expression by approximately 80% compared with the LPS-treated group (LPS) (Fig. 4D). These results indicated that DBM suppressed the inflammatory response through the downregulation of iNOS and COX-2.
Cells were treated with DBM or vehicle (DMSO) for 24 h in the presence or absence of LPS. NO production in the medium was detected by using Griess reagent (A). The morphological change of cells was observed by using a microscope (B). The protein expression of COX-2 and iNOS was analyzed by Western blotting (C) and quantified by using ImageJ software (D). The values are presented as means and standard error of the mean (mean±S.E.M.). Different letters indicate statistically significant differences among groups (p<0.05). CON: no treatment, PMA: only PMA treated. Three replicates of each experiment were performed.
The effect of DBM was examined on the phosphorylation of IκBα and the translocation of NF-κB p65 to nucleus. In the normal state, IκBα, a regulator of NF-κB translocation, is weakly detected in the phosphorylated form, but LPS treatment markedly induced the phosphorylation of IκBα (Fig. 5A). The LPS-induced phosphorylation of IκBα was dose-dependently decreased by DBM; 50 µM DBM suppressed the phosphorylation of IκBα by more than 70% (Fig. 5B). In addition, DBM effectively suppressed the LPS-induced translocation of NF-κB into the nucleus (Fig. 5C). The high dose of DBM blocked NF-κB translocation into the nucleus by 60% compared with the LPS group (Fig. 5D). In contrast, cytosolic NF-κB p65 in LPS treatment group was 27% lower than in the untreated control group (CON), but DBM (50 µM) increased cytosolic NF-κB p65 by 40% compared with LPS group (Fig. 5D).
Cells were treated with DBM 2 h before LPS treatment for 5–15 min. The effect of DBM (0–50 µM) on the abundance of p-IκBα, IκBα, and cytosolic/nuclear NF-κB p65 was analyzed by using Western blotting (A and C) and quantified by ImageJ software (B and D). The mass of nuclear protein analyzed was 300 µg. Different letters indicate a statistically significant difference among groups (p<0.05). CON: no treatment, LPS: only LPS treated. Three replicates of each experiment were performed.
LPS treatment caused a large reduction in the protein expression of Nrf2 and its target molecule, HO-1, compared with the untreated control (CON), but the LPS-induced reduction of their levels was suppressed by DBM treatment (Fig. 6A). The Nrf2 protein abundance was twice as high after treatment with 50 µM DMB than in the LPS group (Fig. 6B); similarly, HO-1 abundance was also increased by five-fold. However, Keap1, a negative regulator of Nrf2 nuclear translocation, in a high dose of DBM, was 72% lower after 50 µM DBM treatment than in the LPS group (Fig. 6B). In addition, DBM effectively promoted the translocation of Nrf2 into nucleus (Fig. 6C). The translocation of Nrf2 in the 50 µM DBM group was more than four-fold higher than in LPS group, whereas cytosolic Nrf2 content was more than 50% lower (Fig. 6D). After LPS treatment, most of the Nrf2 proteins were in the cytosolic fraction, but this was decreased by DBM treatment (Fig. 6C). These results showed that DBM activates the Nrf2/Keap1 pathway and the DBM-mediated increase in HO-1 resulted from the translocation of activated Nrf2.
RAW264.7 cells were treated with DBM for 24 h in the presence or absence of LPS. Proteins from each group were extracted and 50 µg of protein analyzed was by Western blotting using antibodies against total Nrf2, Keap1, HO-1, and GAPDH (A). Protein expression was quantified by using ImageJ (B). Nuclear and cytosolic fractions were prepared as described in the Methods and subjected to Western blotting with antibodies against Nrf2 and Lamin B (nuclear loading control) proteins (C). The mass of nuclear and cytosolic proteins used was 300 µg and 50 µg, respectively. Protein expression was quantified by using ImageJ software (D). The values are presented as the mean and standard error of the mean (mean±S.E.M.). Different letters indicate statistically significant differences among the groups (p<0.05). CON: no treatment, LPS: only LPS treatment. Three replicates of each experiment were performed.
The expression of pro-inflammatory cytokines was examined after DBM treatment. LPS treatment markedly increased the mRNA expression of cytokines (Fig. 7), which indicated the induction of the pro-inflammatory response. This upregulation of cytokines was reduced in a dose-dependent manner by DBM treatment. After treatment with the high dose of DBM, tumor necrosis factor alpha (TNF-α), IL-1β, IL-6, and monocyte chemoattractant protein-1 (MCP-1) were downregulated by 64, 52, 75, and 40%, respectively, compared with the LPS group. This result showed that DBM effectively inhibited the gene expression of pro-inflammatory cytokines (Fig. 7).
RAW264.7 cells were treated with DBM or vehicle (DMSO) in the presence or absence of LPS for 24 h. Total RNA was extracted from the cells and the mRNA expression of the indicated cytokines were analyzed by real-time PCR; expression was normalized to GAPDH. The values are presented as the mean and standard error of the mean (mean±S.E.M.). Different letters indicate statistically significant difference among the groups (p<0.05). CON: no treatment, LPS: only LPS treatment. Three replicates of each experiment were performed.
Monocytes have been recognized to play an important role in the development of atherosclerosis.27,28) Circulating monocytes differentiate into tissue macrophages after exposure to various regulatory signals and continue to be accumulated, forming plaque in arterial lesions with adherent properties. The activation of monocyte-to-macrophage has been known to promote inflammation and atherosclerosis.27) Therefore, the regulation of the monocyte-to-macrophage differentiation can offer a crucial protection from inflammatory diseases such as atherosclerosis. Our results demonstrated that DBM had the potential to suppress atherosclerogenesis through the inhibition of PMA-induced THP-1 differentiation and adherence with the downregulation of CD11β and CD36, which are surface markers of monocyte-to-macrophage differentiation (Fig. 3). DBM also suppressed inflammatory responses. Nitric oxide (NO), an inflammatory response, has a critical role in inflammation status. It exerts an inhibitory function on inflammation in normal physiological conditions, although the chronic excessive production of NO acts as a pro-inflammatory agent.29,30) The excessive exposure of NO, both endogenously and exogenously, affects the carcinogenic process and leads to genomic alterations.30) The overproduction of nitric oxide (NO) mediated by iNOS is known to induce inflammatory reactions and also can lead to negative cellular physiologies including mutagenesis, DNA damage, and the formation of N-nitrosoamine.29,30) Cyclooxygenase-2 (COX-2) can also generate prostaglandins, including PGE2, which are pro-inflammatory substances that lead to inflammation.31) DBM was observed to suppress iNOS and COX-2 protein expression in a dose-dependent manner (Fig. 4), which indicated that DBM effectively inhibited the inflammatory response. It was also thought to inhibit PGE2 production through the downregulation of COX-2, but this should be examined in future studies. Various natural compounds have been reported to control these inflammatory mediators: flavonoids, including quercetin, genistein, and kaempferol, have been reported as natural COX-2 inhibitors.17,32,33) A recent study showed that a component of olive oil, oleuropein, suppressed the expression of iNOS and COX-2.34) Resveratrol and curcumin are also known to be effective inhibitors of these inflammatory mediators in several cell lines.35) The inflammatory response accompanies systematic activation of many signaling pathways. Nuclear factor-kappa-light-chain-enhancer of activated B cells (NF-κB) has long been recognized as a major component in pro-inflammatory signaling, responsible for the expression and activation of various inflammatory cytokines.36) In basal conditions, NF-κB is controlled by IκBα, which binds to NF-κB to form a complex in the cytoplasm; in other words, IκBα prevents the nuclear transport of NF-κB. However, NF-κB is detached from the complex with IκBα and moved to the nucleus by the stimulation of signaling molecules including LPS, inflammatory cytokines, TNF-α, or oxidative and fluid mechanical stress.36) Simultaneously, IκBα is phosphorylated and degraded by ubiquitination in the proteasome. DBM effectively inhibited the phosphorylation of IκBα and suppressed the transport of NF-κB into the nucleus (Fig. 5). This inhibition of NF-κB was considered to lead to the DBM-mediated suppression of the inflammatory responses, including the production of inflammatory cytokines. Many studies have reported that anti-inflammatory phytochemicals regulate the transactivation of NF-κB into nucleus. Yoon et al. reported the inhibitory effect of apple extract on NF-κB activation.37) A component of green tea, epigallocatechin-3-gallate (EGCG), was also shown to inhibit the inflammatory response via the suppression of NF-κB activation.38)
The Nrf2/Keap1 system is another important regulator of inflammation. When this system is exposed to certain signals, such as reactive oxygen species or electrophiles, Keap1 is quickly degraded by proteasome-mediated ubiquitination, which leads to the dissociation of Nrf2 from Keap1. Free Nrf2 can then translocate to the nucleus and bind to antioxidant response elements (ARE) of its target genes to promote their expression.39) The activation of Nrf2 and its target molecules, such as HO-1, has been considered as an intracellular protective regulator against oxidative stress and inflammatory responses.40) The Nrf2/Keap1 pathway is proposed to regulate inflammation through two possible mechanisms. One mechanism is that Nrf2-targeted molecules, which are mostly anti-oxidants, suppress the inflammatory responses through the regulation of intracellular reactive oxygen species (ROS) production. HO-1 is a representative Nrf2-targeted molecule exerting anti-inflammatory activity,40) through the regulation of ROS production, which is known to induce inflammation.41) This study showed that DBM mediated the regulation of Nrf2/Keap1 pathway via the upregulation of Nrf2 and its target gene product HO-1, the downregulation of Keap1 (Fig. 6). Accordingly, the DBM-mediated regulation of ROS through the Nrf2/Keap1 pathway is considered to contribute to the anti-inflammatory response. In addition, ROS generation has also been a mechanism associated with the monocyte-to-macrophage differentiation.42–44) Many studies showed that ROS is critically produced during macrophage differentiation, but ROS removal or elimination blocks the macrophage differentiation.42–44) Based on our previous study showing that DBM suppressed intracellular ROS generations,45) DBM-mediated inhibition of differentiation of monocyte to macrophage is also considered to be due to the ROS regulation. The other possible mechanism for Nrf2-related anti-inflammation is based on the crosstalk of Nrf2 and NF-κB. Nrf2 activation has been reported to interfere with the interaction of NF-κB and its target genes in the nucleus and subsequently suppress inflammatory responses.10) In particular, HO-1, the target gene product of Nrf2, has inhibited the NF-κB-mediated transcription of inflammatory adhesion molecules.46) This potential mechanism was also supported by our data, which showed that DBM increased Nrf2 and decreased nuclear NF-κB (Figs. 5, 6). Thus, DBM-mediated Nrf2 activation may be a potential mechanism through which the NF-κB-mediated upregulation of inflammatory cytokines is blocked (Fig. 7). However, the detailed effect of DBM on crosstalk the Nrf2 and NF-κB requires further study.
This study was performed using 50 µM of DBM in cell line. The anti-inflammatory effect of DBM needs to be confirmed via in vivo study. In simple arithmetical figure, 50 µM used in this cell line study corresponds to 11.2 mg/kg which is considered to be low concentration in the animal. However, this value may not be directly applicable to animal due to the complexity of animal body. Even then, the DBM amount used in animal study has been recognized to be much higher than that of above arithmetic level of DBM in vivo. A recent study used DBM of 100 mg/kg in animal study,47) and another study used a diet supplemented with 1% DBM in in vivo study.48) These reports indicate that wide range of dosage in DBM could be accepted in animal study.
The biological effects of DBM in animal have been generally studied on chemical-induced tumor formation.48,49) Lin et al. reported that DBM effectively suppressed TPA-induced tumor promotion in ear edema of mouse,49) and Cheung et al. showed that DBM had a chemopreventive effect in an azoxymethane (AOM)-initiated and dextran sodium sulfate (DSS)-promoted colon cancer mouse model.48) A recent study showed that DBM derivatives inhibited LPS-induced NO production in microglial cells.26) However, systematic analyses of the inflammatory response, including signaling, have been limited. The current study has addressed the association of DBM and inflammatory signaling, including Nrf2 and NF-κB, as an anti-inflammatory mechanism, and suggested a possible role of DBM in protection against atherosclerosis via the inhibition of monocyte-to-macrophage differentiation. This study supports the potential of DBM as an anti-inflammatory agent.
This work was supported by the National Research Foundation of Korea (NRF), a Grant funded by the Korea government (the Ministry of Education) (NRF-2015R1D1A1A01059729) (2017).
The authors declare no conflict of interest.
