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Introduction

Reactive oxygen species (ROS) are biologically significant due to their role in cellular redox signaling (1). It has been reported that ROS may induce cellular damage and increase physiological dysfunction (2). In locations where ROS accumulate, oxidative damage occurs, which has been linked to a diverse range of neurodegenerative disorders, including Alzheimer's disease, Parkinson's disease, amyotrophic lateral sclerosis, prion diseases, hereditary ataxia, dentatorubral-pallidoluysian atrophy and Wilson's disease, as well as cancer and skin aging (3,4). Glutamate is the fundamental excitatory neurotransmitter, which is activated via N-methyl-D-aspartate receptors. Several important physiological functions are co-regulated by glutamate, and excessive concentrations of glutamate lead to the pathological effects caused by ROS. In addition, glutamate-associated neurotoxicity is implicated in numerous neuronal disorders (5). Hydrogen peroxide (H2O2) is the product of a non-radical two-electron reduction of oxygen, which has previously been implicated in redox signaling and oxidative stress (6). Heme consists of Fe2+ and protoporphyrin IX, and is a prosthetic group that is present in hemoglobin and myoglobin (7). Free heme can lead to oxidative damage; therefore, in order to degrade heme, cells produce the rate-limiting enzyme heme oxygenase (HO). HO-1, which is an inducible form of HO, degrades heme into three byproducts: Biliverdin, carbon monoxide (CO) and Fe2+. CO is generally considered to be toxic; however, recent studies have suggested that it exerts antiproliferative, anti-inflammatory and anti-apoptotic effects (8–10). Biliverdin is converted into bilirubin by biliverdin reductase, and numerous reports have suggested that bilirubin exerts antioxidant effects (11,12). Fe2+ is immediately converted to ferritin, which has a protective effect against heme synthesis (13). Nuclear factor erythroid-derived 2-related factor-2 (Nrf2) has been reported to act as a positive regulator of detoxification enzyme gene expression, and recent studies have demonstrated that Nrf2 translocation regulates the expression of hundreds of cytoprotective genes, which counteract endogenously- or exogenously-generated oxidative stress (14). Nrf2 is a major upstream donor that induces HO expression (15). Mitogen-activated protein kinases (MAPKs) are associated with the majority of signal transduction pathways, including those involved in cell differentiation, cell proliferation, cell survival and cell transformation (16). The MAPK family comprises extracellular signal-regulated kinase (ERK), c-Jun NH2-terminal kinase (JNK) and p38 MAPK. Previous studies have demonstrated that MAPK is activated by oxidative stress or other stimuli, and that phosphorylation of MAPK regulates the expression of diverse genes and proteins, including HO-1 (17,18).

KCHO-1 is a novel mixture comprised of 30% ethanol (EtOH) extracts obtained from nine natural products: Curcuma longa, Salvia miltiorrhiza, Gastrodia elata, Chaenomeles sinensis, Polygala tenuifolia, Paeonia japonica, Glycyrrhiza uralensis, Atractylodes japonica and processed Aconitum carmichaeli. These natural products are well known as traditional medicinal herbs, which are used as alternative therapies in Korea and China, and recent studies have reported the beneficial effects of these herbs (19–25). In our previous study, it was suggested that KCHO-1 exerted anti-inflammatory effects in BV2 microglia (26). Using an in vitro oxidative stress model, the present study aimed to explore the direct neuroprotective effects of KCHO-1, and to determine the possible underlying mechanisms.

Materials and methods

Reagents

Dulbecco's modified Eagle's medium (DMEM) and other tissue culture reagents were purchased from Gibco (Thermo Fisher Scientific, Inc., Waltham, MA, USA). The HO activity inhibitor tin protoporphyrin IX (SnPP IX) was obtained from Porphyrin Products (Frontier Scientific, Logan, UT, USA). Primary antibodies, including rabbit polyclonal anti-HO-1 (1:1,000 dilution; cat. no. sc-10789), rabbit polyclonal anti-Nrf2 (1:1,000 dilution; cat. no. sc-722), goat polyclonal anti-lamin B (1:1,000 dilution; cat. no. sc-6216) and goat polyclonal anti-actin (1:1,000 dilution; cat. no. sc-1616) were purchased from Santa Cruz Biotechnology, Inc. (Heidelberg, Germany). Rabbit polyclonal anti-phosphorylated-ERK (1:1,000 dilution; cat. no. 9101) and rabbit polyclonal anti-ERK (1:1,000 dilution; cat. no. 9102) antibodies were obtained from Cell Signaling Technology, Inc. (Danvers, MA, USA). Secondary horseradish peroxidase (HRP)-conjugated polyclonal goat anti-rabbit IgG (1:1,000 dilution; cat. no. sc-2004) and HRP-conjugated normal goat IgG (1:1,000 dilution; cat. no. sc-2741) were purchased from Santa Cruz Biotechnology, Inc. The HO-1 inducer cobalt protoporphyrin IX (CoPP) and all other chemicals used were obtained from Sigma-Aldrich (St. Louis, MO, USA).

Extract preparation

C. longa, C. sinensis, P. tenuifolia, P. japonica, G. uralensis and A. japonica were purchased from Won Kwang Herb Co., Ltd. (Jinan, South Korea) in August 2013. S. miltiorrhiza and G. elata were purchased from Dongkyung Pharm. Co., Ltd. (Boeun, South Korea). Processed A. carmichaeli was purchased from Hanpoong Pharm & Foods Co., Ltd. (Jeonju, South Korea). All voucher specimens were deposited at Hanpoong Pharm & Foods Co., Ltd. [C. longa (HP2013-10-01), S. miltiorrhiza (HP2013-10-02), G. elata (HP2013-10-03), C. sinensis (HP2013-10-04), P. tenuifolia (HP2013-10-05), P. japonica (HP2013-10-06), G. uralensis (HP2013-10-07), A. japonica (HP2013-10-08), and processed A. carmichaeli (HP2013-10-09)]. To prepare the extract, C. longa (4 kg), S. miltiorrhiza (4 kg), G. elata (4 kg), C. sinensis (2 kg), P. tenuifolia (2 kg), P. japonica (2 kg), G. uralensis (2 kg), A. japonica (2 kg) and processed A. carmichaeli (1 kg) were mixed, pulverized and extracted in 30% EtOH for 3 h at 84–90°C. Subsequently, the mixture was concentrated using a rotary evaporator and lyophilized.

High-performance liquid chromatography (HPLC) analysis

The sample was analyzed by reversed-phase HPLC using a Sykam HPLC (Sykam GmbH, Eresing, Germany), equipped with S7131 Reagent Organizer, S2100 Solvent Delivery system, S7511 Vacuum Degaser, S5200 Sample Injection and S3210 UV/Vis Detector. HPLC-grade acetonitrile was purchased from Burdick & Jackson® (Honeywell; Muskegon, MI, USA). Data processing was carried out using ChromStar DAD (GPC) software (Sykam GmbH). An Inertsil-ODS3 column (150×4.6 mm; particle size, 5 μm; GL Sciences Inc., Torrance, CA, USA) was used in the stationary phase. The mobile phase consisted of eluent A (0.1% formic acid in water with 10% acetonitrile) and eluent B (acetonitrile). The starting eluent was 100% A. The proportion of eluent B was increased linearly to 36% from 0 to 60 min, increased to 60% from 60 to 90 min, and increased to 100% from 90 to 110 min. The detector wavelength was set over a range of 190–700 nm and recorded at 254 nm. The flow rate was 1.0 ml/min, and the injection volume was 20 μl. Identification was based on comparison of retention time and ultraviolet (UV) spectra with commercial standards. For each compound, peak areas were determined as the wavelength providing maximal UV absorbance.

Cell culture and viability assay

The HT22 mouse hippocampal cells were provided by Dr. Inhee-Mook (Seoul National University, Seoul, South Korea). The cells were maintained in DMEM supplemented with 10% heat-inactivated fetal bovine serum, penicillin G (100 units/ml), streptomycin (100 mg/ml) and L-glutamine (2 mM), and were incubated at 37°C in a humidified atmosphere containing 5% CO2 and 95% air. For determination of cell viability, HT22 cells (1×105 cells/well in 24-well plates) were incubated with glutamate (0.5–20 mM; Sigma-Aldrich) and H2O2 (10–500 μM) for 12 h, or pre-treated with KCHO-1 (10–200 μg/ml; Sigma-Aldrich) for 12 h. SnPP IX (50 μM; Sigma-Aldrich) was used as an inhibitor of HO, and trolox (50 μM) was used as a positive control, incubated with 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT; Sigma-Aldrich) at a final concentration of 0.5 mg/ml for 4 h. Subsequently, the formazan that had formed was dissolved in acidic 2-propanol. Optical density (OD) was measured at 590 nm using a microplate reader (model no. 680; Bio-Rad Laboratories, Inc., Hercules, CA, USA). The OD of formazan in the control (untreated) cells was considered to represent 100% viability.

ROS measurement

To measure ROS, HT22 cells (2.5×104 cells/well in 24-well plates) were treated with 5 mM glutamate (5 mM) in the presence or absence of KCHO-1 (10–200 μg/ml) or SnPP IX (50 μM) for 8 h. After washing with phosphate-buffered saline (PBS), the cells were stained with 10 μM 2′,7′-dichlorofluorescein diacetate in Hank's balanced salt solution for 30 min in the dark. The cells were then washed twice with PBS and extracted with 1% Triton X-100 in PBS for 10 min at 37°C. Fluorescence was recorded at an excitation wavelength of 490 nm and an emission wavelength of 525 nm (Spectramax Gemini XS; Molecular Devices, Sunnyvale, CA, USA). Cells were immediately observed under a laser-scanning confocal microscope (TCS SP2; Leica Microsystems, Wetzlar, Germany). Dichlorofluorescein fluorescence was excited at 488 nm with an argon laser, and the resulting emission was filtered with a 515-nm long pass filter.

Western blot analysis

HT22 cells were treated with KCHO-1 (10–200 μg), harvested and pelleted by centrifugation at 200 × g for 3 min. Subsequently, the cells were washed with PBS and lysed using radioimmunoprecipitation assay lysis buffer [25 mmol/l Tris-HCl buffer, pH 7.6; 150 mmol/l NaCl; 1% NP-40; 1% sodium deoxycholate; 0.1% sodium dodecyl sulfate (SDS)]. Protein concentration was determined using Bradford Assay Reagent (Bio-Rad Laboratories, Inc.). An equal amount of protein (30 μg) from each sample was separated by 12% SDS-polyacrylamide gel electrophoresis and was then electrophoretically transferred onto a Hybond-enhanced chemiluminescence (ECL) nitrocellulose membrane (Bio-Rad Laboratories, Inc.). The membrane was blocked with 5% skim milk and incubated with primary antibodies at 4°C overnight, then incubated with secondary antibodies at room temperature for 1 h. The bands were then visualized using ECL (RPN2232; GE Healthcare Life Sciences, Chalfont, UK) and quantified by densitometry using Image J software (version 1.47; National Institutes of Health, Bethesda, MA, USA). In the figures, representative blots from three independent experiments are presented, and the data are presented as the mean ± standard deviation of three independent experiments. Nuclear and cytoplasmic cell extracts were prepared using NE-PER Nuclear and Cytoplasmic Extraction Reagents (Pierce Biotechnology, Inc., Rockford, IL USA).

Reverse transcription-quantitative polymerase chain reaction (PCR) analysis

Total RNA was isolated from the cells using TRIzol® (Invitrogen; Thermo Fisher Scientific, Inc.), according to the manufacturer's protocol, and was quantified spectrophotometrically at 260 nm (ND-1000; Thermo Fisher Scientific, Inc.). Total RNA (1 μg) was reverse transcribed using the High Capacity RNA-to-cDNA kit (Applied Biosystems; Thermo Fisher Scientific, Inc.) according to the manufacturer's protocol. cDNA was amplified using the SYBR Premix Ex Taq kit (Takara Bio. Inc., Shiga, Japan) on a StepOnePlus Real-Time PCR system (Applied Biosystems; Thermo Fisher Scientific, Inc.). Briefly, each reaction volume contained 10 μl SYBR Green PCR Master Mix, 0.8 μM each primer and diethyl pyrocarbonate-treated water, with a final reaction volume of 20 μl. The primer sequences were designed using PrimerQuest (Integrated DNA Technologies, Coralville, IA, USA). The primer sequences were as follows: HO-1, forward 5′-CTC TTG GCT GGC TTC CTT-3′, reverse 5′-GGC TCC TTC CTC CTT TCC-3′; and glyceraldehyde 3-phosphate dehydrogenase (GAPDH), forward 5′-ACT TTG GTA TCG TGG AAG GACT-3′ and reverse 5′-GTA GAG GCA GGG ATG ATG TTCT-3. The optimal conditions for PCR amplification were established according to the manufacturer's protocol. The thermal cycling conditions used were as follows: Pre-denaturation at 95°C for 10 min; denaturation at 95°C for 15 sec; and annealing at 60°C for 1 min. A total of 40 cycles were performed. The data were analyzed using StepOne software (version 2.3; Applied Biosystems; Thermo Fisher Scientific, Inc.), and the cycle number at the linear amplification threshold (quantification cycle; Cq) was recorded for the endogenous control gene and the target gene. Relative gene expression (target gene expression normalized to the expression of the endogenous control gene) was calculated using the comparative Cq method (2−ΔΔCq) (27).

Statistical analysis

The data are presented as the mean ± standard deviation of at least three independent experiments. To compare three or more groups, one-way analysis of variance followed by the Newman-Keuls post-hoc test was conducted. Statistical analysis was performed using GraphPad Prism software, version 3.03 (GraphPad Software, Inc., San Diego, CA, USA). P<0.05 was considered to indicate a statistically significant difference.

Results

Effects of KCHO-1, glutamate and H2O2 on HT22 cell viability

The present study examined KCHO-1 cytotoxicity on HT22 cells using the MTT method. As shown in Fig. 1A, HT22 cells were incubated with 10–400 μg/ml KCHO-1, and cell viability was unchanged following treatment with all doses of KCHO-1. Therefore, in the present study, KCHO-1 was used at a concentration of 10–200 μg/ml. Subsequently, the effects of glutamate (0.5–20 mM) and H2O2 (10–500 μM) were determined on the viability of HT22 cells. Glutamate significantly reduced cell viability when used at a concentration of >2 mM (Fig. 1B). H2O2 induced cell death when used at a concentration of >50 μM (Fig. 1C). Therefore, glutamate and H2O2 were subsequently used at concentrations of 5 mM and 100 μM, respectively.

Figure 1 Effects of (A) KCHO-1, (B) glutamate and (C) hydrogen peroxide (H2O2) on the viability of HT22 cells. HT22 cells were incubated for 48 h with various concentrations of KCHO-1 (10–400 μg/ml), glutamate (0.5–20 mM) or H2O2 (10–500 μM). Data are presented as the mean ± standard deviation of three independent experiments. *P<0.05 vs. the control untreated cells.

Effects of KCHO-1 on glutamate-induced oxidative neurotoxicity and ROS generation in HT22 cells

The present study investigated whether KCHO-1 affected glutamate-induced oxidative cell toxicity and ROS generation in HT22 cells. Cell viability was lower in the glutamate-treated cells compared with in the control group, whereas pretreatment with KCHO-1 (100–200 μg/ml) increased viability in a dose-dependent manner (Fig. 2A). In addition, glutamate treatment doubled ROS production, whereas KCHO-1 markedly attenuated this increase (Fig. 2B). The known antioxidant trolox was used as a positive control.

Figure 2 Effects of KCHO-1 on glutamate-induced (A) oxidative neurotoxicity and (B) reactive oxygen species (ROS) generation in HT22 cells. HT22 cells were pretreated with KCHO-1 for 12 h and were then incubated for 12 h with glutamate (5 mM). Exposure of HT22 cells to glutamate increased ROS production. Data are presented as the mean ± standard deviation of three independent experiments. Trolox (50 μM) was used as a positive control. #P<0.05 vs. the control group. *P<0.05 vs. glutamate-treated cells.

Effects of KCHO-1 on H2O2-induced oxidative neurotoxicity and ROS generation in HT22 cells

The present study also determined the protective action of KCHO-1 against H2O2-induced neurotoxicity in HT22 cells. Compared with the untreated cells, treatment with H2O2 caused cell death and induced ROS production; however, pretreatment with KCHO-1 (100–200 μg/ml) increased viability in a concentration-dependent manner (Fig. 3A). Furthermore, KCHO-1 significantly suppressed H2O2-induced ROS generation (Fig. 3B).

Figure 3 Effects of KCHO-1 on hydrogen peroxide (H2O2)-induced (A) oxidative neurotoxicity and (B) reactive oxygen species (ROS) generation in HT22 cells. HT22 cells were pretreated with KCHO-1 for 12 h and were then incubated for 12 h with H2O2 (100 μM). Exposure of HT22 cells to H2O2 increased ROS production. Data are presented as the mean ± standard deviation of three independent experiments. Trolox (50 μM) was used as a positive control. #P<0.05 vs. the control group. *P<0.05 vs. H2O2-treated cells.

Effects of KCHO-1 on the mRNA and protein expression levels of HO-1 in HT22 cells

The present study detected HO-1 expression in KCHO-1-treated HT22 cells. The HT22 cells were treated with non-cytotoxic concentrations of KCHO-1 (10–200 μg/ml) for 12 h, and HO-1 mRNA (Fig. 4A) and protein expression levels (Fig. 4B) were increased in a dose-dependent manner. As a HO-1 inducer, CoPP was used as a positive control and dose-dependently increased HO-1 mRNA and protein expression levels (Fig. 4A and B).

Figure 4 Effects of KCHO-1 on heme oxygenase (HO)-1 (A) mRNA and (B) protein expression in HT22 cells. HT22 cells were incubated with the indicated concentrations of KCHO-1 for 12 h (10–200 μg/ml). Data are presented as the mean ± standard deviation of three independent experiments. Cobalt protopor-phyrin IX (CoPP; 20 μM) was used as a positive control. *P<0.05 vs. the control untreated cells.

Effects of KCHO-1 on Nrf2 nuclear translocation in HT22 cells

Nrf2 nuclear translocation is a key inducer of HO-1 expression; therefore, the present study investigated whether pretreatment of HT22 cells with KCHO-1 also upregulated Nrf2 nuclear translocation (Fig. 5A and B). Cells were treated with KCHO-1 for 0.5, 1.0 or 1.5 h at a concentration of 200 μg/ml. Nrf2 levels gradually decreased in the cytoplasm of HT22 cells (Fig. 5A), whereas nuclear Nrf2 levels markedly increased in a time-dependent manner (Fig. 5B).

Figure 5 Effects of KCHO-1 on the nuclear translocation of nuclear factor erythroid-derived 2-related factor-2 (Nrf2) in HT22 cells. HT22 cells were treated with KCHO-1 (200 μg/ml) for 0.5, 1 or 1.5 h. The nuclei were fractionated from the cytosol using PER-Mammalian Protein Extraction buffer. Nrf2 (A) cytoplasmic and (B) nuclear protein expression was detected using western blot analysis, and representative blots of three independent experiments are shown. Data are presented as the mean ± standard deviation of three independent experiments. *P<0.05 vs. the control untreated cells.

Effects of KCHO-1-induced HO-1 expression via Nrf2 nuclear translocation on glutamate- and H2O2-induced oxidative neurotoxicity

The present study subsequently assessed whether KCHO-1-induced HO-1 upregulation was responsible for the observed cytoprotective effects. HT22 cells were co-treated with 200 μg/ml KCHO-1 for 12 h in the absence or presence of the HO inhibitor SnPP IX. SnPP partially inhibited the ability of KCHO-1 to suppress glutamate-induced cytotoxicity and ROS generation (Fig. 6A and B). Furthermore, SnPP partially inhibited the ability of KCHO-1 to suppress H2O2-induced cytotoxicity and ROS generation (Fig. 6C and D). These results suggest that HO-1 expression may be required for the inhibition of H2O2-induced ROS generation.

Figure 6 Effects of KCHO-1-induced nuclear factor erythroid-derived 2-related factor-2-mediated heme oxygenase-1 expression on glutamate- and hydrogen peroxide (H2O2)-induced oxidative neurotoxicity and reactive oxygen species (ROS) generation in HT22 cells. HT22 cells were treated with 200 μg/ml KCHO-1 in the presence or absence of 50 μM tin protoporphyrin IX (SnPP). (A) Cell viability and (B) ROS generation were measured following treatment with glutamate (5 mM) for 12 h. (C) Cell viability and (D) ROS generation were also measured following treatment with H2O2 (100 μM) for 12 h. Data are presented as the mean ± standard deviation of three independent experiments. *P<0.05.

Effects of KCHO-1-induced ERK activation on HO-1 expression, and glutamate- and H2O2-induced neurotoxicity

To investigate the role of MAPKs in KCHO-1-induced HO-1 expression, the present study examined the effects of specific inhibitors, including PD98059 (ERK inhibitor), SP600125 (JNK inhibitor) and SB203580 (p38 inhibitor). As shown in Fig. 7A, ERK inhibition suppressed KCHO-1-induced HO-1 expression, whereas JNK and p38 inhibition did not. In addition, ERK phosphorylation was detected following KCHO-1 treatment between 15 and 60 min (Fig. 7B). PD98059 also partially reversed the ability of KCHO-1 to inhibit glutamate- and H2O2-induced cell toxicity (Fig. 7C and D). Data from the HPLC analysis of KCHO-1 was obtained in the form of chromatograms by monitoring responses at 254 nm. As presented in Fig. 8, the retention time of the main peak was 38.858 min.

Figure 7 Effects of KCHO-1 induced extracellular signal-regulated kinase activation on heme oxygenase (HO)-1 expression, and glutamate- and hydrogen peroxide (H2O2)-induced neurotoxicity. (A) Cells were incubated with the indicated concentration of KCHO-1 (200 μg/ml) for 12 h in the presence or absence of PD98059 (20 μM), SP600125 (25 μM) or SB203580 (20 μM), and western blotting was performed with a HO-1 antibody. (B) HT22 cells were treated with KCHO-1 (200 μg/ml) for the indicated times. HT22 cells untreated or treated with KCHO-1 (100 or 200 μg/ml) in the presence or absence of PD98059 (20 μM) for 12 h were exposed to (C) 5 mM glutamate or (D) 100 μM H2O2 for 12 h. Data are presented as the mean ± standard deviation of three independent experiments. *P<0.05 vs. glutamate (5 mM)- or H2O2 (100 μM)-treated cells; ##P<0.05 vs. KCHO-1-treated cells; #P<0.05 vs. the control group.

Figure 8 High-performance liquid chromatography (HPLC) chromatograms of KCHO-1. Chromatograms were obtained via HPLC using KCHO-1 dissolved in acetonitrile at a concentration of 10 mg/ml. The method utilized a hydro column at a flow rate of 1 ml/min, with an acetonitrile/water/formic acid gradient for 110 min, and the wavelength was detected at 254 nm. The ultraviolet spectra peak at 38.858 min was identified as the major component of KCHO-1.

Discussion

Oxidative stress in brain tissue may occur physiologically, as a result of neurodegenerative disorders (28). Therefore, the authors of the present study have focused on the mechanism of action of natural products against neurodegenerative diseases via HO-1 regulation (29–32). In our previous study, the extract KCHO-1 was developed (26). The present study investigated the association of HO-1 with the neuroprotective action of KCHO-1, via Nrf2 nuclear translocation. To determine the therapeutic potential of KCHO-1, its direct neuroprotective effects on glutamate- and H2O2-induced oxidative damage were investigated in HT22 mouse hippocampal cells.

The HT22 immortalized neuronal cell line has been used as an in vitro model for mechanistic identification of glutamate-induced oxidative damage. In the central nervous system, glutamate is the main excitatory neurotransmitter that is released by nerve cells in the brain; however, glutamate toxicity induces neuronal cell death, which is associated with acute insults and chronic neurodegenerative disorders (33,34). Glutamate-mediated oxidative stress is caused by inhibiting cellular cystine uptake, leading to glutathione depletion or ROS generation and elevated Ca2+ levels (35). H2O2 is the product of a non-radical two-electron reduction of oxygen, and has been reported to have a key role in oxidative cell death (6). Therefore, it may be therapeutically beneficial to reduce the damaging effects of oxidative glutamate or H2O2 toxicity. As shown in Fig. 1, the present study initially evaluated the action of glutamate (5 mM) and H2O2 (100 μM) on the viability of HT22 cells. Subsequently, it was investigated whether KCHO-1 was able to affect glutamate- or H2O2-induced oxidative neurotoxicity and ROS generation in HT22 cells. KCHO-1 significantly suppressed glutamate- and H2O2-induced cell damage and ROS generation (Figs. 2 and 3).

In our previous studies, it was demonstrated that HO-1 expression may have an important role in the protection of HT22 cells (36,37). It has been suggested that the role of HO-1 in heme degradation may offer cells protection against oxidative insults and maintain cellular homeostasis. The antioxidant activities of HO-1 have been observed in Alzheimer's disease, sepsis, endotoxemia, surgical stress, ischemia reperfusion injury and psychological stress (26,38). In the present study, cells were treated with non-cytotoxic concentrations of KCHO-1. The results indicated that the mRNA and protein expression levels of HO-1 were increased in HT22 cells (Fig. 4). Furthermore, the present study assessed whether KCHO-1-mediated HO-1 upregulation was responsible for its protective effects on HT22 cells. Treatment with the HO-1 inhibitor SnPP partially reversed the ability of KCHO-1 to inhibit H2O2-induced cell death and ROS generation (Fig. 6). These results suggested that HO-1 expression may be required to inhibit H2O2-induced ROS generation. Nrf2 is a basic leucine zipper transcription factor, which resides in the cytoplasm bound to Keap-1. Following stimulation with inducers, Nrf2 translocates into the nucleus (39–41). Nrf2 has been reported to induce the expression of antioxidant proteins, including HO-1 (42). The present study revealed that KCHO-1 significantly upregulated Nrf2 and efficiently promoted its translocation into the nucleus, thus suggesting that KCHO-1-induced HO-1 expression may be associated with Nrf2 nuclear translocation (Fig. 5).

The present study also demonstrated that the ERK pathway is involved in KCHO-1-induced HO-1 expression (Fig. 7). MAPK is one of the most common cellular response signaling pathways, which responds to various extracellular stimuli. There are three subfamilies of MAPK: p38 kinase, ERK1/2 and JNK (43). MAPKs are initiated in response to various extracellular stimuli, particularly oxidative stress. Previous studies have reported that activation of MAPK pathways may contribute to HO-1 gene expression (44,45). In the present study, KCHO-1-induced HO-1 gene expression was shown to be associated with the ERK pathway, since treatment with the ERK inhibitor, PD98059, suppressed KCHO-1-induced HO-1 expression; however, JNK and p38 inhibition did not affect HO-1 expression. As expected, treatment with the ERK pathway inhibitor also abolished KCHO-1-induced cytoprotection (Fig. 7). These results indicated that KCHO-1-induced HO-1 expression in HT22 cells may be mediated by the Nrf2 or ERK pathways.

In conclusion, the results of the present study suggested that KCHO-1 may effectively prevent glutamate- or H2O2-induced oxidative cell damage in a murine hippocampal cell line. KCHO-1-induced HO-1 upregulation via ERK and Nrf2 pathways appears to have a central role in the protection of HT22 cells. These results may provide an insight into the mechanisms underlying KCHO-1-induced neuronal cell protection and HO-1 enzyme induction. Therefore, KCHO-1 may be considered a potential agent for the treatment of neurodegenerative diseases.

Acknowledgments

The present study was supported by the Traditional Korean Medicine R&D Program funded by the Ministry of Health & Welfare through the Korea Health Industry Development Institute (KHIDI) (grant no. HI11C2142).

References