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Cite This: J. Nat. Prod. 2018, 81, 1411−1416
Neuroprotective Compound from an Endophytic Fungus, Colletotrichum sp. JS-0367 Ji Hoon Song,†,# Changyeol Lee,‡,# Dahae Lee,§,# Soonok Kim,⊥ Sunghee Bang,‡ Myoung-Sook Shin,§ Jun Lee,∥,▽ Ki Sung Kang,*,§ and Sang Hee Shim*,‡ †
Department of Medicine, University of Ulsan College of Medicine, Seoul 05505, South Korea College of Pharmacy, Duksung Women’s University, Seoul 01369, South Korea § College of Korean Medicine, Gachon University, Seongnam 13120, South Korea ⊥ National Institute of Biological Resources, Incheon 22689, South Korea ∥ Herbal Medicine Research Division, Korea Institute of Oriental Medicine, Daejeon 34054, Republic of Korea ▽ Convergence Research Center for Diagnosis, Treatment and Care System of Dementia, Korea Institute of Science and Technology, Seoul 02792, South Korea
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‡
S Supporting Information *
ABSTRACT: Colletotrichum sp. JS-0367 was isolated from Morus alba (mulberry), identified, and cultured on a large scale for chemical investigation. One new anthraquinone (1) and three known anthraquinones (2−4) were isolated and identified using spectroscopic methods including 1D/2D-NMR and HRESIMS. Although the neuroprotective effects of some anthraquinones have been reported, the biological activities of the four anthraquinones isolated in this study have not been reported. Therefore, the neuroprotective effects of these compounds were determined against murine hippocampal HT22 cell death induced by glutamate. Compound 4, evariquinone, showed strong protective effects against HT22 cell death induced by glutamate by the inhibition of intracellular ROS accumulation and Ca2+ influx triggered by glutamate. Immunoblot analysis revealed that compound 4 reduced the phosphorylation of MAPKs (JNK, ERK1/2, and p38) induced by glutamate. Furthermore, compound 4 strongly attenuated glutamate-mediated apoptotic cell death.
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secondary metabolites have shown antimicrobial activities against bacteria and fungi.6 Oxidative stress is known to be a major cause of neuronal cytotoxicity in the central nervous system (CNS). The brain is highly vulnerable to oxidative stress because of its relatively high demand for oxygen and unsaturated fat content.7 Although glutamate is a well-known CNS excitatory neurotransmitter, an excess concentration can trigger neuronal cell death by excitotoxicity and oxidative stress and contributes to neurological diseases, including Alzheimer’s diseases, Parkinson’s disease, ischemic brain injuries, traumatic brain injuries, amyotrophic lateral sclerosis, and epilepsy.8−11 Glutamate-mediated oxidative stress is mainly
he extracts of root barks, leaves, and fruits of Morus alba L. (mulberry) have been used to improve health for a long time in oriental medicine.1 The leaves of mulberry trees have been valued as a primary food for silkworms for centuries. Previous studies have reported on the anti-inflammatory and hypoglycemic activities of the bark extract and the antidiabetic, antiatherosclerotic, antiobesity, and hepatoprotective activities of the leaf extract.2,3 Endophytic fungi, which live within the tissues of the plant, have notable mutualistic symbiotic relationships with their host plant.4 Endophytes have been considered as potential sources for various bioactive metabolites with intriguing structures, which could be useful drug candidates.4 Among the endophytes, Colletotrichum sp., a filamentous fungus, is distributed worldwide and is genetically diverse;5 its extracts and © 2018 American Chemical Society and American Society of Pharmacognosy
Received: January 10, 2018 Published: May 23, 2018 1411
DOI: 10.1021/acs.jnatprod.8b00033 J. Nat. Prod. 2018, 81, 1411−1416
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which was confirmed by HSQC and HMBC data (Figure 1). The HMBC data of the aromatic proton at δH 7.75 (H-5) with the
caused by the inhibition of cysteine uptake, which gives rise to a reduction in the levels of glutathione and the accumulation of reactive oxygen species (ROS) within the cell.12,13 Furthermore, intracellular calcium ion ([Ca2+]i) levels have a critical role in physiological and pathological brain functions. During glutamate-induced oxidative stress, the level of [Ca2+]i is dramatically increased, disrupting homeostasis and resulting in neuronal cell death. Oxidative stress and Ca2+ influx induced by glutamate contribute to the activation of various cellular signaling pathways. Mitogen-activated protein kinases (MAPKs) are well-known signaling molecules that can be activated by glutamate. Typically, MAPKs, a serine/threonine kinase family, include c-Jun N-terminal kinase (JNK), extracellular signal-regulated kinase (ERK), and p38 MAP kinase (p38).14 Previous reports have revealed that the persistent activation of ERK was related with neuronal cytotoxicity mediated by oxidative stress, whereas its specific inhibition prevented neuronal cell death by excessive glutamate exposure in primary cortical neurons.15 Moreover, it was recently reported that MAPKs, including ERK, JNK, and p38, were excessively phosphorylated during glutamate-induced cytotoxicity and were associated with glutamate-triggered apoptosis.16 Therefore, the inhibition of MAPKs may be beneficial to treat neurological diseases. In the present study, a bioactive secondary metabolite was isolated and identified from cultures of Colletotrichum sp., an endophytic fungus isolated from M. alba L.; the metabolite demonstrated potent neuroprotection against excessive glutamate-induced apoptosis in the immortalized murine HT22 hippocampal neuronal cell line, and the underlying molecular mechanism was investigated.
Figure 1. Observed key HMBC correlations of compound 1.
ketone carbon at δC 182.3 (C-10), the methyl carbon at δC 22.4 (C-11), and the aromatic methine carbon at δC 118.53 (C-7), in addition to the HMBC data of the methoxy proton at δH 4.05 (8-OCH3) with the carbon at δC 160.9 (C-8), indicated that the methyl group and one of the methoxy groups were attached to the C-6 and C-8 of the anthraquinone ring, respectively (Figure 1). The attachment of the remaining methoxy group to the carbon at δC 139.1 (C-2) was confirmed by HMBC. The carbon resonance for C-2 appeared at a relatively high field due to two electrondonating groups in the ortho positions, which could be observed in the cases of trioxygenated anthraquinones.17 In addition, the exchangeable hydroxy proton showed HMBC correlation with carbons at δC 155.9 (C-1), 139.1 (C-2), and 112.5 (C-9a), which suggested that one hydroxyl group and one methoxy group were attached to C-1 and C-2, respectively (Figure 1). Through the HMBC correlation of the remaining aromatic methine proton at δH 7.38 (H-4) with the carbonyl carbon at δC 182.3 (C-10), together with the carbon chemical shift at δC 154.1 (C-3), one more hydroxy group was assumed to be attached to C-3. The known compounds were identified as 1-hydroxy-2,3,8trimethoxy-6-methylanthraquinone (2),18 1,2-dihydroxy-3,8dimethoxy-6-methylanthraquinone (3),19 and evariquinone (4),17 respectively, through comparison of the NMR data with literature values. This study presents the first report of compound 2 from nature, although it was once reported as an intermediate of a chemical synthesis.17 Previously, studies reported that 6-methyl-1,3,8,-trihydroxyanthraquinone, a derivative of anthraquinone, prevented tauand beta-amyloid-induced neuronal cell death.20,21 Therefore, to examine the neuroprotective effect, HT22 cells with or without compounds were treated with 5 mM glutamate for 24 h, and the cell viability was then measured using the EZ-Cytox cell viability assay. As a result, the cell viability markedly decreased by treatment with glutamate. When the compounds were added, the viability significantly increased only in the presence of compound 4 (evariquinone) in a concentration-dependent manner compared with compounds 1−3 (Figure 2A−D). Regarding the structures and activities of the anthraquinone derivatives, an increase in the substitution of hydroxyl groups in anthraquinones seems to improve its neuroprotective effect. These hydroxyl groups trap excess electrons balancing ROS, and therefore methylation of the phenolic hydroxyl group decreased the neuroprotection. Consistently, microscopic images also showed that compound 4 markedly prevented HT22 cell death induced by excessive glutamate stimulation (Figure 2E). These data indicate that compound 4 is a strong neuroprotectant against HT22 cell death induced by glutamate. High concentrations of glutamate are known to produce excessive amounts of intracellular ROS, which result in neuronal cell death.22 Antioxidants, such as N-acetylcysteine, can prevent oxidative-stress-mediated neuronal cytotoxicity both in vitro and in vivo. As shown in Figure 3A, the DPPH scavenging activity of
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RESULTS AND DISCUSSION Colletotrichum sp. JS0367 was isolated from the leaves of M. alba L., cultured, and extracted with ethyl acetate, and one new anthraquinone (1) and three known anthraquinones (2−4) were isolated from the extracts.
The molecular formula of compound 1 was established to be C17H14O6 by 1H and 13C NMR spectroscopic data and (+) HRESIMS. The 1H NMR data of 1 exhibited signals for three aromatic protons at δH 7.75 (s, H-5), 7.38 (s, H-4), and 7.13 (s, H-7); two methoxy groups at δH 4.11 (s, 2-OCH3) and 4.05 (s, 8-OCH3); one methyl group at δH 2.50 (s, H3-11); and one hydroxy proton at δH 13.55 (1H, s). The 13C NMR data of 1 showed 17 carbon signals, corresponding to two carbonyl carbons, 12 aromatic carbons (nine nonprotonated sp2 carbons), two methoxy carbons, and one methyl carbon. The presence of two carbonyl carbons together with two benzenoid moieties suggested that this compound has an anthraquinone skeleton, 1412
DOI: 10.1021/acs.jnatprod.8b00033 J. Nat. Prod. 2018, 81, 1411−1416
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Figure 2. Neuroprotective effects of compounds 1−4. HT22 cells were exposed to 5 mM glutamate (Glu) for 24 h with or without the indicated concentrations of compounds (Com) 1−4, and cell viability was determined: (A) compound 1; (B) compound 2; (C) compound 3; (D) compound 4. The bars denote the percentage of cell viabilities (mean ± SEM, *p < 0.05, **p < 0.001 compared with the glutamate-treated group); (E) microscopic images represent the protective effect of Com 4. Scale bar, 50 μm.
Figure 3. Compound 4 blocked the accumulation of glutamate-induced intracellular ROS. (A) The antioxidant activity of compound 4 (Com 4) was determined with an in vitro DPPH scavenging assay. (B) To determine the levels of intracellular ROS, HT22 cells were exposed to 5 mM glutamate in the presence or absence of compound 4, and the fluorescent intensity was then measured after staining the cells with DCF-DA. Bars denote the fold change in fluorescence intensity compared with the control cells (mean ± SEM, **p < 0.001 compared with the glutamate-treated group).
acute brain insults.23 Thus, the present study investigated whether compound 4 blocked the elevation of Ca2+ within the HT22 cells. Microscopic images revealed that treatment with glutamate markedly increased the level of intracellular Ca2+, whereas compound 4 prevented the glutamate-induced increase in intracellular Ca2+ (Figure 4A). The quantitative results show that the fluorescence intensity of the glutamate-treated cells increased by 3.0-fold compared with that of the control cells, whereas the presence of compound 4 significantly reduced the fluorescence intensity of Fluo-4 to 1.4- and 0.98-fold of the untreated cells (Figure 4B). In addition to the inhibitory effect of compound 4 against the glutamate-mediated accumulation of intracellular ROS, this compound also blocked the increase of Ca2+ induced by glutamate within the HT22 cells. The MAPK signaling pathways are known to be associated with neuronal cell death induced by glutamate. Increases in intracellular ROS and Ca2+ have been suggested to be able to
compound 4 dramatically increased in a concentration-dependent manner, and its IC50 was 42.2 μM with a maximum activity at 100 μM (70 ± 1.78%) (Figure 3A). On the basis of these data, we hypothesized that compound 4 may prevent the glutamateinduced accumulation of intracellular ROS. To confirm this, HT22 cells were stained with 2′,7′-dichlorofluorescin diacetate (DCF-DA), a membrane-permeable indicator for ROS, and then, the fluorescent intensity of DCF was measured. The results showed that the fluorescent intensity of DCF was increased by the exposure to glutamate, while it was significantly reduced by treatment with compound 4 (Figure 3B). Hence, these data indicate that compound 4 contributed to the prevention of oxidative stress triggered by the excessive glutamate stimulation in HT22 cells through its antioxidant property. In addition to ROS, an abnormal increase in intracellular Ca2+ is a contributory factor to neuronal cell death in various neuropathological conditions, such as neurodegenerative diseases and 1413
DOI: 10.1021/acs.jnatprod.8b00033 J. Nat. Prod. 2018, 81, 1411−1416
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Figure 4. Compound 4 blocked the accumulation of intracellular ROS induced by glutamate. The cells were treated with 5 mM glutamate and 50 or 100 μM compound 4 for 8 h and then stained with Fluo-4 AM. The images were obtained with a fluorescent microscope. Scale bar, 50 μM. The fluorescence intensity was quantitatively analyzed and presented as a fold change compared with the control cells (mean ± SEM, **p < 0.001 compared with the glutamate-treated group).
Figure 5. Compound 4 inhibits glutamate-induced activation of MAPKs: (A) The cells were incubated with 5 mM glutamate (Glu) and 50 or 100 μM compound 4 (Com 4) for 8 h. The immunoreactive bands were obtained with specific antibodies to p-JNK, JNK, p-ERK1/2, ERK1/2, p-p38, p38, and GAPDH. (B) Quantitative analysis of the immunoreactive bands. The bar graphs represent the fold change in the phosphorylation of the MAPKs (mean ± SEM, **p < 0.001 compared with the glutamate-treated group).
presence of compound 4 significantly reduced the percentage of annexin V-positive apoptotic cells compared with the glutamateexposed cells (Figure 6). In addition, few (or no) PI-positive cells were found in the glutamate-treated cells (Figure 6). These data suggest that compound 4 attenuates glutamate-mediated apoptotic cytotoxicity in HT22 cells.
activate MAPK signaling pathways, including JNK, ERK1/2, and p38.15,16 Recent studies have suggested that the inhibition of phosphorylation of MAPKs prevents apoptotic cell death in HT22 neuronal cells treated with excessive glutamate.16 Therefore, Western blot analysis was performed to elucidate the effect of compound 4 on the MAPK phosphorylation induced by exposure to glutamate for 8 h. The data show that treatment with glutamate markedly increased the phosphorylation of JNK, ERK1/2, and p38, whereas the presence of compound 4 completely blocked these effects of glutamate (Figure 5A). Quantitative analysis showed that compound 4 significantly prevented the phosphorylation of ERK1/2, JNK, and p38 (Figure 5B). Therefore, these data indicated that the prevention of the accumulation of ROS and Ca2+ reduced the glutamate-mediated activation of MAPKs. High concentrations of glutamate have been reported to induce apoptotic cell death.15 Therefore, to confirm the antiapoptotic effect of compound 4, the HT22 cells were first exposed to 5 mM glutamate for 10 h in the absence or presence of 50 or 100 μM compound 4 and then stained with propidium iodide (PI) and Alexa Fluor-488 annexin V. Our results show that the number of annexin V-positive cells increased by the exposure to glutamate, which indicats an increase in apoptotic cell death, whereas the
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EXPERIMENTAL SECTION
General Experimental Procedures. A Bruker UHR ESI Q-TOF mass spectrometer (Bruker, Germany) was used for the HRESIMS spectra. The NMR spectra were obtained with a Varian NMR Systems 500 MHz (1H: 500 MHz/13C: 125 MHz; Varian, USA) using the solvent signals (CDCl3: δH 7.24/δC 77.0; Cambridge Isotope Laboratories, Inc., USA) as internal standards; chemical shifts are indicated as δ values. High-performance liquid chromatography (HPLC) was performed on an Agilent 1260 Infinity HPLC system (Agilent Technologies, USA) equipped with a G1311C quaternary pump, a G1329B autosampler, a G1315D PDA detector, and a G1316A column oven using a Zorbax SB-C18 column (4.6 × 150 mm, Agilent Technologies, USA). Semipreparative HPLC was performed on a Waters 600 controller (Waters, USA) with a 996 PDA detector using a Zorbax SB-C18 column (21.2 mm × 25 cm, Agilent Technologies, USA). Column chromatography was performed over silica gel 60 (70−230 mesh, Merck, Germany). Thinlayer chromatography (TLC) was carried out on precoated silica gel 60 1414
DOI: 10.1021/acs.jnatprod.8b00033 J. Nat. Prod. 2018, 81, 1411−1416
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Figure 6. Compound 4 prevents apoptotic HT22 cell death induced by glutamate. To assess the antiapoptotic effect of compound 4 (Com 4), the cells were first exposed to 5 mM glutamate (Glu) in the absence or presence of 50 or 100 μM Com 4 for 10 h and then stained with annexin V and propidium iodide (PI). The number of apoptotic cells was analyzed with Tali-Image-based cytometric analysis. The bar graph (right) represents the percentage of cells stained positive for annexin V, an indicator of apoptotic cells (mean ± SEM, **p < 0.001 compared with the glutamate-treated group). 1,3-Dihydroxy-2,8-dimethoxy-6-methylanthraquinone (1): yellow solid; UV (CHCl3) λmax 216, 278, 422 nm; IR (KBR) νmax 3361, 2922, 2851, 1738, 1626, 1580, 1458, 1065, 1020 cm−1; 1H NMR data (500 MHz, CDCl3) δ 13.55 (1H, s, 1-OH), 7.75 (1H, s, H-5), 7.38 (1H, s, H-4), 7.13 (1H, s, H-7), 6.36 (1H, s, 3-OH), 4.11 (3H, s, 2-OCH3), 4.05 (3H, s, 8-OCH3), 2.50 (3H, s, H3-11); 13C NMR data (125 MHz, CDCl3) δ 188.1 (C-9), 182.3 (C-10), 160.9 (C-8), 155.9 (C-1), 154.1 (C-3), 147.1 (C-6), 139.1 (C-2), 135.3 (C-10a), 128.9 (C-4a), 121.1 (C-5), 118.53 (C-7), 118.51 (C-8a), 112.5 (C-9a), 106.7 (C-4), 61.1 (2-OCH3), 56.6 (8-OCH3), 22.4 (C-11); (+) HRESIMS m/z, 337.0682 [M + Na]+, calcd for C17H14O6Na, 337.0683. Cell Culture. HT22 cells, an immortalized mouse hippocampal cell line, were cultured in Dulbecco’s modified Eagle’s medium (DMEM; Corning, Manassas, VA, USA) with 10% fetal bovine serum (Atlas, Fort Collins, CO, USA) and penicillin/streptomycin (Gibco, Grand Island, NY, USA) added. The cells were grown and treated at 37 °C in a humidified atmosphere supplemented with 5% CO2. Measurement of Cell Viability. HT22 cells were placed into a 96-well plate at a density of 1 × 104 cells/well and kept there for 24 h for the adherence of the cells. Glutamate (Sigma, St. Louis, MO, USA) was then added to the cells to a final concentration of 5 mM with the compounds at the indicated concentrations for 24 h. Cell viability was quantitatively measured with an EZ-Cytox cell viability assay kit (Daeil Lab Service, Seoul, Korea) in accordance with the manufacturer’s instruction. Briefly, the cells were treated with 10 μL of Ez-CyTox reagent for 30 min. The absorbance value at 450 nm was measured with an E-Max microplate reader (Molecular Devices, Sunnyvale, CA, USA). The cell viability was indicated as a percentage of the live control cells. Measurement of Reactive Oxygen Species. The intracellular levels of ROS were measured with DCF-DA (Sigma). The cells were placed into black 96-well plates at a density of 1 × 104 cells/well. After exposure to 5 mM glutamate for 8 h, the cells were further treated with 10 μM DCF-DA for 30 min and washed three times with phosphatebuffered saline (PBS) to remove any residual DCF-DA. The fluorescence intensity of DCF at 495 nm (excitation) and 517 nm (emission) was measured with a SPARK 10 M fluorescent microplate reader (Tecan, Männedorf, Switzerland), and the fluorescent intensity was indicated as a percentage of the control. Fluorescent images were acquired with an IX50 fluorescent microscope equipped with a CCD camera (Olympus, Tokyo, Japan). Live Cell Staining for Intracellular Calcium Ion Measurement. To determine the intracellular Ca2+ levels, 5× 104 cells/well were plated onto 24-well plates. The HT22 cells were treated with glutamate and compound 4 for 8 h and then stained with 2 μM Fluo-4 AM (Invitrogen, Eugene, OR, USA), a fluorescent indicator for Ca2+. The cells were briefly washed with phenol-red-free medium and serum. Fluorescent images were obtained with a fluorescence microscope equipped with a CCD camera (Olympus) and quantitatively analyzed with the ImageJ software (National Institutes of Health, Bethesda, MD, USA).
F254 and RP-18 F254S plates (Merck, Germany) using a UV detector and 10% H2SO4 reagent to visualize the compounds. All solvents used for all experiments were of analytical quality. Isolation, Identification, and Culture of Fungus JS-0367. The fungal strain (JS-0367) was isolated from the leaves of the mulberry tree, Morus alba L., collected in Inje-Gun, Gangwon Province, South Korea (3797′99″ N, 12845′23″E) on July 21, 2011. The tissues were cut into small pieces (0.5 × 0.5 cm), and the surfaces were sterilized with a series of 2% sodium hypochlorite for 1 min, 70% ethanol for 1 min, and finally washed with sterilized distilled water. The sterilized plant tissues were transferred to malt extract agar (Difco; BD Diagnostic Systems, Sparks, MD, USA) medium with 50 ppm kanamycin, 50 ppm chloramphenicol, and 50 ppm Rose and incubated for approximately 7 days at 22 °C for the fungal strains to be grown out of the plant tissues. An actively growing fungus was transferred and cultured on potato dextrose agar (PDA) (Difco). This strain was identified as Colletotrichum sp. by the culture morphology on PDA, the microscopic observation of conidia, and the internal transcribed spacer (ITS) sequences.24 This fungus produced long cylindrical conidia similar to that of C. gigasporum.25 Phylogenetic analysis with the ITS sequences suggested this fungus is a close relative of C. gigasporum. The fungus was deposited in the Wildlife Genetic Resources Bank of the National Institute of Biological Resources under the accession number NIBR0000115974. A stock solution of the fungus in 20% glycerol was stored in a liquid nitrogen tank at the Wildlife Genetic Resources Bank at the National Institute of Biological Resources (Incheon, Korea) until used (Code No. JS-0367). Extraction and Isolation of Metabolites. The fungus was grown in Erlenmeyer flasks (5 × 1000 mL) in PDB medium containing potato dextrose broth (24 g) and distilled water (1000 mL). After inoculation, each flask was incubated for 21 days with shaking once daily. The cultures were extracted with EtOAc five times. The EtOAc extracts (3.16 g) were subjected to column chromatography over silica gel with an elution of CHCl3−acetone−MeOH gradient (v/v/v, 100:0:0 → 25:1:0 → 4:1:0 → 1:1:0.1) solvents to give six fractions (1−6). Compound 2 (2.0 mg) was purely isolated from fraction 2 through silica gel column chromatography with a gradient elution of n-hexane−EtOAc (v/v, 60:1 → 15:1). Compound 1 (4.5 mg) was obtained from fraction 3 by HPLC with a C18 column using a gradient solvent system of H2O−MeOH (v/v, 80:20 → 0:100). Fraction 4 was separated by HPLC with a C18 column using a gradient solvent system of H2O−MeOH (v/v, 65:35 → 0:100) to yield seven fractions (4A−4G). Fraction 4E was purified by HPLC with a C18 column using an isocratic H2O−MeOH (v/v, 42:58) solvent to yield compound 3 (1.3 mg). Fraction 5 was separated by C18 column chromatography using a gradient system of acetone−MeOH−water (v/v/v, 1:1:1 → 10:10:1) to yield five fractions (5A−5E). Fraction 5B was purified by HPLC with a C18 column using a gradient solvent system of H2O−MeOH (v/v, 45:55 → 30:70) to yield compound 4 (4.7 mg). 1415
DOI: 10.1021/acs.jnatprod.8b00033 J. Nat. Prod. 2018, 81, 1411−1416
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Measurement and Quantification of Apoptotic Cell Death Using Tali-Image-Based Cytometry. To evaluate the effect of compound 4 on apoptotic cell death induced by glutamate, we performed image-based cytometric analysis. Briefly, cells were cultured on six-well plates at a density of 2 × 105 cells/well, treated with 5 mM glutamate and compound 4 for 10 h, harvested, and washed with PBS. An equal number of cells was suspended with annexin binding buffer and labeled with annexin V Alexa Fluor 488 (Invitrogen) for an additional 20 min. The cells were further stained with propidium iodide in annexin binding buffer. The apoptotic cells were identified with a Tali-Imagebased cytometer (Invitrogen) and analyzed with the TaliPCApp (version 1.0). The data were expressed as representative images and as a percentage of the proportion of annexin V-positive cells. Western Blot Analysis. The cells were incubated with 5 mM glutamate with or without 50 μM casuarinin for 4 h, collected, and lysed with RIPA buffer containing protease inhibitor cocktail (Roche, Indianapolis, IN, USA). SDS-polyacrylamide gel electrophoresis was performed for separation of equal amounts of proteins, which were transferred to polyvinylidene difluoride membranes (Merck Millipore, Darmstadt, Germany). The membranes were blocked with 5% skim milk, probed with primary antibodies for ERK1/2, p-ERK1/2, JNK, pJNK, p38, p-p38, and GAPDH (Cell Signaling, Danvers, MA, USA) for 1 h, and treated with appropriate secondary antibodies as required (Cell Signaling). For visualization of the immunoreactive bands, SuperSignal West Femto Maximum Sensitivity Substrate (Thermo Scientific, Rockford, IL, USA) and a Fusion Solo Chemiluminescence System (PEQLAB Biotechnologie GmbH, Erlangen, Germany) were used, and they were quantitatively analyzed with the ImageJ software and presented a foldincrease compared with the control cells. Statistical Analysis. The data are presented as the mean ± SEM. Statistical significance was assessed with Student’s t test with p values of less than 0.05 considered to be statistically significant.
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(2) Eo, H. J.; Park, J. H.; Park, G. H.; Lee, M. H.; Lee, J. R.; Koo, J. S.; Jeong, J. B. BMC Complementary Altern. Med. 2014, 14, 200−9293. (3) Oh, K. S.; Ryu, S. Y.; Lee, S.; Seo, H. W.; Oh, B. K.; Kim, Y. S.; Lee, B. H. J. Ethnopharmacol. 2009, 122, 216−220. (4) Tan, R. X.; Zou, W. X. Nat. Prod. Rep. 2001, 18, 448−459. (5) Lima, S. J.; Figueiredo, J. G.; Gomes, R. G.; Stringari, D.; Goulin, E. H.; Adamoski, D.; Kava-Cordeiro, V.; Galli-Terasawa, L. V.; Glienke, C. ISRN Microbiol. 2012, 2012, 215716. (6) Wedge, D. E.; Nagle, D. G. J. Nat. Prod. 2000, 63, 1050−1054. (7) Coyle, J. T.; Puttfarcken, P. Science 1993, 262, 689−695. (8) Hynd, M. R.; Scott, H. L.; Dodd, P. R. Neurochem. Int. 2004, 45, 583−595. (9) Blandini, F.; Porter, R. H. P.; Greenamyre, J. T. Mol. Neurobiol. 1996, 12, 73−94. (10) Arundine, M.; Tymianski, M. Cell. Mol. Life Sci. 2004, 61, 657− 668. (11) Meldrum, B. S. Neurology 1994, 44, S14−S23. (12) Shih, A. Y.; Erb, H.; Sun, X.; Toda, S.; Kalivas, P. W.; Murphy, T. H. J. Neurosci. 2006, 26, 10514−10523. (13) Tan, S.; Wood, M.; Maher, P. J. Neurochem. 1998, 71, 95−105. (14) Davis, R. J. J. Biol. Chem. 1993, 268, 14553−14556. (15) Stanciu, M.; Wang, Y.; Kentor, R.; Burke, N.; Watkins, S.; Kress, G.; Reynolds, I.; Klann, E.; Angiolieri, M. R.; Johnson, J. W.; DeFranco, D. B. J. Biol. Chem. 2000, 275, 12200−12206. (16) Fukui, M.; Song, J. H.; Choi, J.; Choi, H. J.; Zhu, B. T. Eur. J. Pharmacol. 2009, 617, 1−11. (17) Bringmann, G.; Lang, G.; Steffens, S.; Gunther, E.; Schaumann, K. Phytochemistry 2003, 63, 437−443. (18) Roberge, G.; Brassard, P. J. Org. Chem. 1981, 46, 4161−4166. (19) Gonzalez, A. G.; Barrera, J. B.; Davila, B. B.; Valencia, E.; Dominguez, X. A. Phytochemistry 1992, 31, 255−258. (20) Liu, T.; Jin, H.; Sun, Q. R.; Xu, J. H.; Hu, H. T. Brain Res. 2010, 1347, 149−160. (21) Pickhardt, M.; Gazova, Z.; von Bergen, M.; Khlistunova, I.; Wang, Y.; Hascher, A.; Mandelkow, E. M.; Biernat, J.; Mandelkow, E. J. Biol. Chem. 2005, 280, 3628−3635. (22) Nakao, N.; Brundin, P. Prog. Brain Res. 1998, 116, 245−263. (23) Wojda, U.; Salinska, E.; Kuznicki, J. IUBMB Life 2008, 60, 575− 590. (24) Cannon, P. F.; Damm, U.; Johnston, P. R.; Weir, B. S. Stud. Mycol. 2012, 73, 181−213. (25) Rakotoniriana, E. F.; Scauflaire, J.; Rabemanantsoa, C.; UrvegRatsimamanga, S.; Corbisier, A. M.; Quetin-Leelereq, J.; Declerck, S.; Munaut, F. Mycol. Prog. 2013, 12, 403−412.
ASSOCIATED CONTENT
S Supporting Information *
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jnatprod.8b00033. 1 H and 13C NMR, HSQC, HMBC, COSY, ESIMS, and HRESIMS spectra for compound 1; morphology and phylogenetic tree of JS-0367 (PDF)
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AUTHOR INFORMATION
Corresponding Authors
*E-mail:
[email protected]. Tel: +82-31-750-5402. *E-mail:
[email protected]. Tel: +82-2-901-8774. ORCID
Sang Hee Shim: 0000-0002-0134-0598 Author Contributions #
J. H. Song, C. Lee, and D. Lee contributed to this work equally.
Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS This research was supported by the National Research Foundation of Korea (NRF-2018R1A2B6001733), by the National Institute of Biological Resources (NIBR201830101), and also by Korea Institute of Planning and Evaluation for Technology in Food, Agriculture, Forestry and Fisheries (IPET) through the High Value-added Food Technology Development Program, funded by the Ministry of Agriculture, Food and Rural Affairs (MAFRA) (grant number 116001-3).
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REFERENCES
(1) Wang, Y.; Xiang, L.; Wang, C.; Tang, C.; He, X. PLoS One 2013, 8, e71144. 1416
DOI: 10.1021/acs.jnatprod.8b00033 J. Nat. Prod. 2018, 81, 1411−1416
