2020 Volume 45 Issue 7 Pages 401-409
Dihydropyrazines (DHPs), including 3-hydro-2,2,5,6-tetramethylpyrazine (DHP-3), are glycation products that are spontaneously generated in vivo and ingested via food. DHPs generate various radicals and reactive oxygen species (ROS), which can induce the expression of several antioxidant genes in HepG2 cells. However, detailed information on DHP-response pathways remains elusive. To address this issue, we investigated the effects of DHP-3 on the nuclear factor-κB (NF-κB) pathway, a ROS-sensitive signaling pathway. In lipopolysaccharide-stimulated (LPS-stimulated) HepG2 cells, DHP-3 decreased phosphorylation levels of inhibitor of NF-κB (IκB) and NF-κB p65, and nuclear translocation of NF-κB p65. In addition, DHP-3 reduced the expression of Toll-like receptor 4 (TLR4) and the adaptor protein myeloid differentiation primary response gene 88 (MyD88). Moreover, DHP-3 suppressed the mRNA expression of tumor necrosis factor-alpha (TNFα), and interleukin-1 beta (IL-1β). Taken together, these results suggest that DHP-3 acts as a negative regulator of the TLR4-MyD88-mediated NF-κB signaling pathway.
Dihydropyrazines (DHPs) are glycation products generated by the Maillard reaction, which begins with nonenzymatic condensation between the carbonyl group of a reduced sugar and an amino group (Glomb and Monnier, 1995). Glycation products are constantly generated and accumulated in a living body. For example, glycated hemoglobin HbA1c is a well-known endogenous glycation product (Kennedy et al., 1978), which is detected in sera of healthy subjects. Glycation products are also unintentionally ingested exogenously glycation via food. Recent studies have shown that excess accumulation of these glycation products in a body leads to chronic diseases such as diabetes (Kawasaki et al., 2002; Goldin et al., 2006), atherosclerosis (Wang et al., 2012), and Alzheimer’s and Parkinson’s disease (Li and Zhang, 2012). Therefore, it is important to reveal the biological effect of glycation products on human health. DHP skeleton was generated from a dimer of D-glucosamine (Kashige et al., 1995) and 5-aminolevulinic acid (Teixeira et al., 2001), and the skeleton was involved in DNA strand-breaking activity. Methyl-substituted DHPs can generate hydroxyl and carbon-centered radicals in electro spin resonance spectroscopy (Yamaguchi et al., 2012), and DNA strand-cleaving against plasmid pBR322 (Yamaguchi et al., 1996; Kashige et al., 2000). DHPs also exert cytotoxicity in HepG2 cells (Ishida et al., 2012).
The transcription factor nuclear factor-κB (NF-κB) plays important roles in diverse cellular processes such as the innate immune response, embryogenesis, organ development, cell proliferation, apoptosis, and stress responses to various external stimuli. In its inactivated state, NF-κB, of which the p50/p65 heterodimer is the major form, is retained in the cytoplasm through binding with its inhibitor of NF-κB (IκB). Upon stimulation by various activating signals, IκB is phosphorylated and degraded through a ubiquitin-dependent proteolysis, following which NF-κB translocates into the nucleus and activates the target genes. The phosphorylation of NF-κB p65 is also required for the activation and nuclear translocation of NF-κB (Hayden and Ghosh, 2004). NF-κB signaling is both activated and inactivated by reactive oxygen species (ROS). Toll-like receptor 4 (TLR4) exhibits immunostimulating activity against lipopolysaccharide (LPS), a membrane glycolipid of Gram-negative bacteria (Akira and Takeda, 2004). Additionally, TLR4 is correlated not only with infectious diseases but also with obesity and cancer metastasis (Suganami et al., 2007; Hiratsuka et al., 2008). TLR4 activation has two signaling pathways, the myeloid differentiation primary response gene 88-dependent (MyD88-dependent) and -independent pathways (Zughaier et al., 2005). The MyD88-dependent signaling pathway depends on Toll-interleukin 1 receptor adaptor protein MyD88. LPS induces the interaction of TLR4 with MyD88 and subsequently activates downstream signaling pathways, including the NF-κB and mitogen-activated protein kinase (MAPK) signaling pathways. The production of pro-inflammatory mediators is strongly affected by MAPKs, including p38, c-Jun N-terminal kinase and extracellular signal-regulated protein kinase (Qian et al., 2015).
Recently, the relationship between DHP and stress response pathways has been revealed. For example, 3-hydro-2,2,5,6-tetramethyipyrazine (DHP-3) (Fig. 1) activates the nuclear factor erythroid 2-related factor 2-antioxidant (Nrf2-antioxidant) responsive element (ARE) pathway, and induces the mRNA and protein productions levels of heme oxygenase-1 (HO-1) and glutamate cysteine ligase catalytic subunit (Ishida et al., 2014). Moreover, DHP-3 also activates the metal-responsive transcription factor 1 pathway, and induces the mRNA and protein productions levels of zinc transporter-1 (Ishida et al., 2015). However, our current understanding is insufficient as to whether DHP-3 may also have an effect on other ROS-responsive signaling pathways. To address this, we have investigated the effect of DHP-3 on the TLR4-MyD88-mediated NF-κB signaling pathway.
Chemical structure of 3-hydro-2,2,5,6-tetramethylpyrazine (DHP-3) in this study.
DHP-3 was synthesized via the condensation of diketones and diamines using the method described by Yamaguchi et al. (1996). DHP-3 stock solution was prepared in dimethyl sulfoxide at 0.1 M, and was stored at -40°C until use.
HepG2 (JCRB1054) was obtained from the Human Science Research Resources Bank (Osaka, Japan). The Dulbecco’s Modified Eagle’s Medium (DMEM) was purchased from Wako (Osaka, Japan). The Cell Count Kit-8 (CCK-8) and the Cytotoxicity Lactate dehydrogenase (LDH) Assay Kit-WST were obtained from Dojindo Molecular Technologies, Inc. (Kumamoto, Japan). LPS was purchased from Sigma-Aldrich (St. Louis, MO, USA). Antibodies against IκB, NF-κB p65, TLR4, MyD88 and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) were obtained from Santa Cruz Biotechnology (Dallas, TX, USA). Phospho-IκBα Ser-32/36 rabbit polyclonal antibody and phospho-NF-κB p65 Ser-536 (93H1) rabbit monoclonal antibody were purchased from Enzo Life Sciences (Farmingdale, NY, USA) and Cell Signaling (Danvers, MA, USA), respectively. Goat anti-mouse immunoglobulin G (IgG) peroxidase antibody, goat anti-rabbit IgG peroxidase antibody and goat anti-mouse IgG fluorescein isothiocyanate-conjugated (FITC-conjugated) antibody were obtained from Sigma-Aldrich. Unless otherwise stated, all the other reagents and chemicals were of the highest grade commercially available.
HepG2 cells were cultured in DMEM supplemented with 10% (v/v) fetal bovine serum (FBS), in a humidified atmosphere with 5% CO2 at 37°C. HepG2 cells were grown in 60-mm culture dishes to 80%-90% confluence before being subjected to various treatments.
Cell viability was determined using the CCK-8 Assay Kit according to the manufacturer’s instructions. Briefly, HepG2 cells were seeded at a density of 5000 cells/well in a 96-well plate. After 24 hr, the culture medium was replaced with various concentrations of DHP-3 containing medium without FBS. After 24 hr, WST-8 dye solution was added to each well. Cells were incubated for 30-60 min in a CO2 incubator to develop color. The absorbance was measured at a wavelength of 450 nm using a microtiter plate reader. The relative cell viability (%) was expressed as a percentage relative to the untreated control cells.
LDH release was determined using the Cytotoxicity LDH Assay Kit-WST according to the manufacturer’s instructions. Briefly, HepG2 cells were seeded at a density of 5000 cells/well in a 96-well plate. After 24 hr, the culture medium was replaced with various concentrations of DHP-3 containing medium without FBS. After 24 hr, LDH release was measured photometrically at 492 nm using a kit. The maximum LDH release was measured after the addition of the lysis buffer attached to the kit. The percentage of LDH release was calculated by comparing the absorbance to the maximum LDH release of the control cells.
After HepG2 cells were treated with 1 μg/mL LPS in serum-free medium, they were exposed to various concentrations of DHP-3 (50-400 µM). Cells were scraped and collected by centrifugation, and cell pellets were lysed with the sample buffer containing 50 mM Tris-HCl (pH 6.8), 2% w/v sodium dodecyl sulfate (SDS), 10% glycerol, 5% 2-mercaptoethanol, and 0.1% w/v bromophenol blue. Cell lysates were sonicated in an ice bath for 15 sec, heated for 4 min at 100°C, separated by 10% SDS-PAGE and transferred onto a polyvinylidene difluoride (PVDF) membrane in a Tris/glycine transfer buffer. The membrane was blocked with PVDF Blocking Reagent (Toyobo, Tokyo, Japan), and then incubated with primary antibodies. After washing with phosphate buffered saline-Tween20 (PBS-Tween20), horseradish peroxidase–conjugated secondary antibodies goat anti-mouse IgG and goat anti-rabbit IgG were applied. The blots were developed using a western blotting detection reagent (Thermo Fisher Scientific, Waltham, MA, USA), and the band intensities were estimated using the VersaDoc™ Imaging System (Bio-Rad Laboratories, Hercules, CA, USA) and the iBright system (Thermo Fisher Scientific).
HepG2 cells were plated on a Lab-Tek® II Chamber SlideTM (Nalge Nunc International, Rochester, NY, USA) with 1.0 × 105 cells/mL/well, and cultured in DMEM with 10% FBS until 70%-80% confluent. After 1 μg/mL LPS stimulation for 6 hr, the cells were exposed to 200 µM DHP-3 for 1 hr. The cells were washed in PBS, fixed in 4% paraformaldehyde on ice for 10 min, methanol permeabilization at -20°C for 10 min, washed in PBS, and blocked in PBS with 1% bovine serum albumin at room temperature for 30 min. After blocking and washing with PBS, the cells were incubated with PBS + 1% anti-NF-κB p65 monoclonal antibody at room temperature for 1 hr. Then anti-mouse IgG FITC-conjugated antibody was applied. Cells were washed in PBS, and the coverslip was mounted and viewed under a TE-2000U inverted fluorescence microscope (Nikon, Tokyo, Japan).
HepG2 cell lysates were reverse transcribed to synthesize cDNA using the Power SYBR® Green Cells-to-CT™ (Life Technologies, Carlsbad, CA, USA) according to the manufacturer’s instructions. q-PCR was performed with each primer using a StepOnePlus™ Real-Time PCR System (Life Technologies) according to the manufacturer’s instructions. Primers are indicated in Table 1. Thermal cycling conditions were applied as follows: 95°C for 10 min, followed by 40 cycles of 15 sec at 95°C and 1 min at 60°C. The results were normalized by GAPDH expression, and fold change (2-ΔΔCt) was compared with that of the control group.
F, forward; R, reverse
Statistical calculations of the data were performed using an unpaired Student’s t-test.
Previous studies have demonstrated, albeit briefly, the cell viability in DHP-3-exposed HepG2 cells (Ishida et al., 2012). To evaluate the cytotoxicity more precisely, HepG2 cells were exposed to various concentrations of DHP-3. The cell viability and cytotoxicity of DHP-3-exposed cells were assessed using a CCK-8 assay and LDH assay, respectively. Cell viability gradually decreased in a dose-dependent manner (Fig. 2A). Correspondingly, a dose-dependent increase in LDH release was observed upon addition of DHP-3 (Fig. 2B). These assays showed that DHP-3 had a severe effect on cell cytotoxicity at concentrations of 500 µM or larger, and that concentrations of 20-200 µM had a sublethal effect. Therefore, 50-400 µM exposure was chosen for most of the subsequent experiments.
Cytotoxic effect of DHP-3 in HepG2 cells. (A) Cell viability was determined by Cell Count Kit-8 (CCK-8) assay after exposure to DHP-3 (20-1000 µM) for 24 hr. (B) Lactate dehydrogenase (LDH) release was evaluated after exposure to DHP-3 (20-1000 µM) for 24 hr. The control was culture medium containing dimethyl sulfoxide (DMSO). Values represent the mean ± S.D. of 3 samples. (*p < 0.05, **p < 0.01, *** p < 0.001 indicate significant differences from the control group).
We investigated whether DHP-3 affected activation of the NF-κB signaling pathway, which is regulated by ROS. As shown in Fig. 3A and C, IκB was phosphorylated by LPS-stimulation. After LPS-stimulation, cells were exposed to each DHP-3 concentration (50, 100, 200, 400 µM) for 1 hr, during which the phosphorylation levels of IκB significantly decreased (Fig. 3A-C).
Effect of DHP-3 on activation of the nuclear factor-κB (NF-κB) signaling pathway in lipopolysaccharide-stimulated (LPS-stimulated) HepG2 cells. After 1 μg/mL LPS-stimulation for 6 hr, the HepG2 cells were exposed to various concentrations of DHP-3 (50, 100, 200, 400 µM) for 1 hr. The control was culture medium containing DMSO and LPS. (A) Protein levels of inhibitor of NF-κB (IκB), phospho-IκB (p-IκB), NF-κB p65 and phospho-NF-κB p65 (p-NF-κB p65) were examined by immunoblotting analysis. (B, D) The band intensities of total IκB and NF-κB p65 were normalized to the expression levels of glyceraldehyde-3-phosphate dehydrogenase (GAPDH). (C, E) The relative ratios of p-IκB/IκB and p-NF-κB p65/NF-κB p65 were present. Values represent the mean ± S.D. of 3 samples. (*p < 0.05, **p < 0.01 indicate significant differences from the LPS-stimulation without DHP-3 group.)
The IκB phosphorylation decrease indicated that the subsequent NF-κB inactivation was induced by DHP-3. Additionally, the effect of DHP-3 on NF-κB activity was investigated using immunoblotting and immunofluorescence assays. Exposure of HepG2 cells to these DHP-3 concentrations (50, 100, 200, 400 µM) for 1 hr decreased the phosphorylation of NF-κB in response to LPS (Fig. 3A, D and E). DHP-3 markedly suppressed the nuclear translocation of NF-κB in the immunofluorescence assay (Fig. 4).
Effect of DHP-3 on the nuclear translocation of NF-κB in LPS-stimulated HepG2 cells. After LPS-stimulation for 6 hr, the HepG2 cells were exposed to 200 µM DHP-3 for 1 hr. HepG2 cells were immunostained with anti-NF-κB p65 monoclonal antibody, and labeled with fluorescein isothiocyanate-conjugated (FITC-conjugated) secondary antibody. Nuclei were stained with 4’,6-diamidino-2-phenylindole (DAPI). Green fluorescence from FITC and blue fluorescence from DAPI were monitored using a confocal laser scanning microscope. The control was culture medium containing DMSO.
To explore the underlying inhibitory mechanisms by DHP-3 against the NF-κB signaling pathway in LPS-stimulated HepG2 cells, we examined the expression of TLR4 and MyD88. TLR4 regulates LPS-stimulated inflammatory mediator expression through the NF-κB and MAPK signaling pathways, and LPS stimulates protein expressions of TLR4 and MyD88 (Hsiao et al., 2015; Tseng et al., 2019). As shown in Fig. 5A and B, after LPS-stimulation, the LPS-stimulated TLR4 protein levels were suppressed by 50 µM DHP-3. Correspondingly, the LPS-stimulated protein expression levels of MyD88 were suppressed by 200 µM DHP-3 (Fig 5A and C). These data suggested that DHP-3 inhibited the activation of the TLR4-MyD88 pathway via NF-κB signaling in LPS-stimulated HepG2 cells.
Effect of DHP-3 on Toll-like receptor 4 (TLR4) and adapter protein myeloid differentiation primary response gene 88 (MyD88) expression in LPS-stimulated HepG2 cells. After 1 μg/mL LPS-stimulation for 6 hr, the HepG2 cells were exposed to various concentrations of DHP-3 (50, 100, 200, 400 µM) for 1 hr. The control was culture medium containing DMSO and LPS. (A) Protein levels of TLR4 and MyD88 were detected by immunoblotting analysis. (B, C) The band intensities of TLR4 and MyD88 were normalized to the expression levels of GAPDH. Values represent the mean ± S.D. of 3 samples. (*p < 0.05, **p < 0.01 indicate significant differences from the LPS-stimulation without DHP-3 group).
As DHP-3 suppressed activation of the TLR4-NF-κB pathway, we examined whether pro-inflammatory cytokines of TLR4-NF-κB downstream were suppressed by DHP-3. Two hundred micro molar DHP-3 exposure was shown to suppress the expression of tumor necrosis factor-alpha (TNFα) and interleukin-1 beta (IL-1β) mRNA after LPS-stimulation (Fig. 6A and B). Correspondingly, LPS-posttreated cells also inhibited the mRNA expression of TNF-α and IL-1β after exposure to 200 µM DHP-3 (Fig. 6A and B). These results indicated that DHP effectively suppressed the pro-inflammatory response regardless of pretreatment and exposure conditions.
DHP-3 inhibits the production of pro-inflammatory cytokines in LPS-stimulated HepG2 cells. The mRNA levels of tumor necrosis factor-alpha (TNFα) (A) and interleukin-1 beta (IL-1β) (B) were analyzed using quantitative polymerase chain reaction after normalization to GAPDH mRNA. After 1 μg/mL LPS-stimulation for 6 hr, the HepG2 cells were exposed to 200 µM DHP-3 for 1 hr (pretreatment LPS). After 200 µM DHP-3 exposure for 1 hr, the HepG2 cells were stimulated by 1 μg/mL LPS for 6 hr (posttreatment LPS). Values represent the mean ± S.D. of 3 samples. (#p < 0.05, ##p < 0.01 indicate significant differences from the untreated group. **p < 0.01 indicate significant differences from the LPS-stimulation without DHP-3 group).
In this study, we demonstrated the effect of the low-molecular-weight glycation product DHP-3 on the NF-κB signaling pathway. In LPS-stimulated HepG2 cells, DHP-3 inhibited the phosphorylation of IκB and NF-κB, and suppressed the nuclear translocation of NF-κB. DHP-3 also reduced the expression of TLR4 and the TLR adaptor protein MyD88. Furthermore, DHP-3 repressed transcription of pro-inflammatory cytokines TNF-α and IL-1β. We previously demonstrated that, with exposure l mM DHP-3 to HepG2 cells, an apparent increase in intracellular hydrogen peroxide concentration is observed. In addition, with exposure to HepG2 cells at least 0.5 mM DHP-3, increases in HO-1 and Nrf2 protein levels are observed, and ARE-luciferase reporter activity is significantly increased (Ishida et al., 2014). However, the effect of DHP-3 on the NF-κB signaling pathway was observed with lower concentration than that of the Nrf2-ARE signal pathway. Therefore, it seems likely that DHP-3 has more effect on the NF-κB signaling pathway than that of the latter.
ROS can induce cellular toxicity by reacting with biomolecules, such as proteins, lipids, and nucleic acids. The role of ROS in NF-κB activation by inflammatory cytokines and LPS has been the focus of several studies, and has revealed distinct mechanisms for various stimuli (Gloire et al., 2006). The NF-κB signaling pathway was activated by H2O2 whereas this activation was blocked by the anti-oxidant N-acetyl cysteine (Schreck et al., 1991), although H2O2 may act as a direct inducer, but rather, as an indirect modulator of the NF-κB signaling pathway (Oliveira-Marques et al., 2009). NF-κB can be modified under the influence of ROS. Cys-62 in the Rel homology domain of the p50 subunit tends to be oxidized by ROS. Consequently, ROS reduce the ability of NF-κB binding to DNA (Gambhir et al., 2015). Phosphorylation of Ser-536 of the p65 subunit is induced by N-acetyl cysteine, and the shift lead to increase in the ability of NF-κB binding to DNA (Liu et al., 2008). TLR4, which is able to combine adaptor molecules including MyD88, can activate the NF-κB signaling pathway. LPS binding to TLR4 sequentially causes production of pro-inflammatory cytokines, such as TNF-α, IL-1β, interleukin-6, interleukin-8, and interleukin-12, which induce pathogenesis of liver diseases (Wang et al., 2013). Moreover, in regard to control of TLR4 gene expression, LPS stimulates NF-κB binding in the TLR4 gene promoter, and NF-κB inhibitor attenuates TLR4 gene expression (Wan et al., 2016). Overall, our observations raise the possibility that DHP-3 can suppress the TLR4-MyD88-mediated NF-κB signaling pathway by suppression of NF-κB, yet future experiments will test this possibility.
This study is the first to demonstrate the effects of DHP-3 on LPS-stimulated HepG2 cells through inhibiting the activation of the TLR4-MyD88-mediated NF-κB signaling pathway. Further studies are required to explore the effects of DHP-3 in an animal model of hepatoma.
The authors declare that there is no conflict of interest.