Caffeic Acid Phenethyl Ester from the Twigs of Cinnamomum cassia

(2) The failure of these cellular regulators in malignant cells can arise from genetic ... pharmacological properties that can treat diarrhea, edema, ...
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Caffeic Acid Phenethyl Ester from the Twigs of Cinnamomum cassia Inhibits Malignant Cell Transformation by Inducing c‑Fos Degradation Seung Ho Shin,†,∥ Seoung Rak Lee,‡,∥ Eunjung Lee,§ Ki Hyun Kim,*,‡ and Sanguine Byun*,⊥ †

Program in Bioinformatics and Computational Biology, University of Minnesota, Minneapolis, Minnesota 55455, United States School of Pharmacy, Sungkyunkwan University, Suwon 440-746, Republic of Korea § Traditional Alcoholic Beverage Research Team, Korea Food Research Institute, Seongnam 13539, Republic of Korea ⊥ Division of Bioengineering, Incheon National University, Incheon 22012, Republic of Korea ‡

S Supporting Information *

ABSTRACT: The twigs of Cinnamomum cassia, commonly referred to as Cinnamomi Ramulus, are widely used as one of the primary ingredients in Chinese/Korean traditional medicines that have anticancer effects. However, the active constituents responsible for its anticancer effects and their molecular mechanisms still remain to be elucidated. Caffeic acid phenethyl ester (CAPE) and caffeic acid (CA) were isolated for the first time from C. cassia using LC-MS-guided phytochemical isolation methods. CAPE significantly suppressed EGF- and TPA-induced cell transformation of JB6 P+ cells at sub-micromolar concentrations, whereas CA, a structurally similar compound to CAPE, had no such effect. The antiproliferative and chemopreventive activity of CAPE was found to arise through the inhibition of AP-1 transcriptional activity via the promotion of c-Fos degradation. These findings demonstrate that CAPE may contribute to the chemopreventive/chemotherapeutic effects of C. cassia through downregulating c-Fos.

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on the active constituents of Cinnamomi Ramulus, which refers to tiny twigs of C. cassia (Lauraceae). The bark and twigs of C. cassia have been reported to possess pharmacological properties that can treat diarrhea, edema, blood circulatory dysfunction, amenorrhea, and gastrointestinal disorders.15 Previous research has shown that an extract from the twigs of C. cassia exhibits anti-inflammatory, antibacterial, and antioxidant properties16,17 and contains biologically active constituents including cinnamaldehyde, protocatechuic acid, coumarin, eugenol, cinnamic acid, and daucosterol.18−21 Cinnamaldehyde, a primary constituent of the extract, harbors numerous biological properties including antiviral, antiinflammatory, analgesic, antipyretic, and antibacterial effects.18,21−24 Recent reports have suggested that an extract from the twigs of C. cassia can induce antiproliferative effects with cinnamaldehyde being a primary active constituent.23,25,26 However, the moderate potency of cinnamaldehyde at the micromolar range led us to investigate other active constituents in C. cassia twigs and their molecular mechanisms. In the present study, our LC/MS analysis of the MeOH extract of C. cassia twigs revealed for the first time that the MeOH extract contains caffeic acid phenethyl ester (CAPE), an anticancer component of propolis extract.27−30 An LC-MSguided isolation technique was applied for the separation of the target constituents. As a result, CAPE (1) and caffeic acid (CA)

ancer is one of the leading causes of death worldwide, and the incidence is expected to grow.1 Cancer develops through the successive accumulation of loss in control mechanisms for cell proliferation, angiogenesis, invasion, and cell death.2 The failure of these cellular regulators in malignant cells can arise from genetic mutations influenced by socioeconomic, cultural, and dietary factors.3,4 Despite ongoing efforts to identify effective cancer treatments, significant unmet needs exist. As a potential source of novel treatments, numerous natural products have been investigated for anticancer activity. Historically, anticancer herb formulations have been developed and used in the form of traditional medicines. Cinnamomi Ramulus is a major ingredient of traditional Chinese and Korean anticancer herbal medicine. One prominent example is Geiji-Bokryung-Hwan, which includes Cinnamomi Ramulus [Cinnamomum cassia Blume (Lauraceae)] as an ingredient.5 Previous studies of Geiji-Bokryung-Hwan have shown anticancer effects in a two-stage mouse skin cancer model and human hepatocarcinoma cells.6 In addition, Cinnamomi Ramulus has similarly been used as an ingredient in Yang-Dan-Tang, a Chinese herbal formulation that was shown to inhibit lung cancer cell proliferation via G1/G2 phase arrest without affecting normal cells.7 Despite the widespread usage of Cinnamomi Ramulus in anticancer preparations, the chemical components responsible for the activity have not been comprehensively analyzed. As part of our ongoing search for anticancer compounds from natural resources,8−14 we focused © 2017 American Chemical Society and American Society of Pharmacognosy

Received: May 18, 2017 Published: July 6, 2017 2124

DOI: 10.1021/acs.jnatprod.7b00433 J. Nat. Prod. 2017, 80, 2124−2130

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(2) were isolated from subfractions E3 and E1 of an EtOAcsoluble fraction, respectively. Herein, we report the identification of anticancer compounds present in the twigs of C. cassia using an LC/MS-guided isolation method and elucidate c-Fos degradation as a potential mechanism of action of CAPE.

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RESULTS AND DISCUSSION

The dried twigs of C. cassia were extracted with 100% MeOH to obtain a crude MeOH extract, and LC/MS analysis revealed the presence of CAPE with a molecular ion peak at m/z 285 [M + H]+ in the positive ESI mode. The major fragment at m/z 163 in ESIMS2 indicated [C9H7O3]+ and another stable fragment at m/z 105 indicated [C8H9]+. On the basis of the molecular weight and fragmentation pattern, the compound was tentatively identified as CAPE (Figure S1).31 As CAPE was not previously reported from C. cassia, LC/MS-guided isolation was carried out for the target separation of CAPE. The high sensitivity and selectivity of the LC/MS-guided isolation method effectively reduced the time of analysis and consequently enabled fast isolation of target constituents. The target compound, CAPE (1), was isolated by preparative and semipreparative HPLC monitored by LC/MS analysis, which led to the identification of the E3 fraction containing the desired constituent. Similarly LC/MS screening of the E1 fraction depicted a peak of caffeic acid (2) with a molecular ion peak at m/z 181 [M + H]+ and its MS2 patterns, which was isolated by semipreparative HPLC. Using a combination of 1D NMR (1H and 13C NMR) and ESIMS data and comparing the spectroscopic data with previously reported values, the chemical structures of the isolates were determined as CAPE (1) and CA (2).32,33 This represents the first report of CAPE being identified in C. cassia. As epidermal growth factor (EGF) and 12-O-tetradecanoylphorbol 13-acetate (TPA) are well-known tumor promoters,34−36 we assessed the cancer chemopreventive potential of CAPE and CA on EGF- and TPA-induced malignant cell transformation in JB6 P+ cells. CAPE exerted strong inhibitory activity against EGF- and TPA-induced cell transformation in a dose-dependent manner, whereas CA exhibited little or no protective effect at identical concentrations (Figure 1A and B). It was also confirmed that CAPE and CA were not cytotoxic toward JB6 P+ cells (Figure 1C). These results demonstrate that CAPE (but not CA) can suppress malignant cell transformation, and this occurs in the absence of cytotoxicity. EGF treatment induced cell cycle progression, whereas CAPE treatment reversed this effect by causing G1 cell cycle arrest and reducing the G2/M phase population (Figure 2). However, CA had no such effect on cell cycle progression (Figure 2). AP-1 is a dimeric transcription factor that plays a major role in regulating malignant cell transformation and cancer progression.37,38 It was observed that CAPE, but not CA, significantly suppressed EGF- and TPA-induced AP-1 activity (Figure 3A and B). As there are several subcomponents that constitute the AP-1 complex,39 we further examined which component in AP-1 was responsible for CAPE-mediated reduction. Among the AP-1 components tested, c-Fos and phosphorylated-c-Jun were shown to be the most highly upregulated in the presence of EGF treatment (Figure S2). Next, we examined whether CAPE was involved in modulating EGF-induced c-Fos or phosphorylated-c-Jun activity. CAPE

Figure 1. Inhibitory effect of CAPE and CA on EGF- and TPAinduced cell transformation. (A and B) JB6 P+ cells were treated with EGF (10 ng/mL) for 12 days (A) or TPA (20 ng/mL) for 14 days (B). CAPE or CA was treated at the indicated concentrations. A total of 8000 cells/well were seeded, and the colony numbers were analyzed by Image-Pro Plus software. (C) At 24 and 48 h after treatments, CAPE and CA showed no toxicity up to 10 μM in the JB6 P+ cell line. Data are presented as means ± standard deviation of triplicate samples. The asterisks (*p < 0.05, **p < 0.01, and ***p < 0.001) indicate a significant difference versus the EGF only or TPA only treated group.

specifically targeted c-Fos while showing negligible effects toward phosphorylated-c-Jun (Figure 3C and D). To investigate the molecular mechanism responsible for CAPE-mediated suppression of c-Fos activity, we analyzed the levels of c-Fos protein and its upstream signaling pathway. Treatment with CAPE attenuated the phosphorylation of c-Fos and downregulated total protein levels of EGF-induced c-Fos (Figure 4). Conversely, treatment with CA did not affect EGFinduced c-Fos expression (Figure S3). Although the ERK-RSKCREB signaling pathway regulates c-Fos expression levels,40,41 CAPE treatment did not affect the phosphorylation of ERK, RSK, or CREB (Figure 4), suggesting that the decrease in c-Fos by CAPE occurs independently of these factors. Since c-Fos protein expression was reduced by CAPE treatment, we examined the mRNA level of c-Fos. CAPE did not decrease EGF-induced c-fos mRNA levels (Figure 5A), leading us to hypothesize that CAPE may suppress c-Fos levels 2125

DOI: 10.1021/acs.jnatprod.7b00433 J. Nat. Prod. 2017, 80, 2124−2130

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Figure 2. Effect of CAPE and CA on EGF-induced cell cycle progression. Cells were synchronized by serum deprivation for 24 h and were pretreated with various concentrations of CAPE or CA for 1 h before adding EGF. Cell cycle analysis was performed at 24 h after EGF (10 ng/mL) treatment (n = 3). Special symbols (singlet: p < 0.05, doublet: p < 0.01, and triplet: p < 0.001) indicate a significant difference versus the EGF only treated group (*G1, #S, and ϕG2/M).

Figure 4. CAPE downregulates c-Fos expression but not phosphorylations of the ERK-RSK-CREB pathway. JB6 P+ cells were serumstarved for 24 h and pretreated with CAPE (0.5 and 1 μM) for 1 h. Cells were collected 15 min after EGF (10 ng/mL) treatment, and immunoblot analysis was performed.

via protein degradation. While CAPE treatment alone decreased EGF-induced c-Fos, cotreatment with a proteasome inhibitor, MG132, prevented CAPE-mediated downregulation of c-Fos (Figure 5B). Collectively these results suggest that CAPE reduces c-Fos levels by inducing its degradation by the proteasome. In the current study, we have identified CAPE and CA from twigs of C. cassia using LC/MS-guided isolation techniques and demonstrated that CAPE can inhibit malignant cell transformation and AP-1 activity at sub-micromolar concentrations. We believe that the chemopreventive effect of CAPE primarily stems from its ability to suppress c-Fos by promoting its

degradation by the proteasome. Our results show that reduction in AP-1 activity and c-Fos levels mediated by CAPE occurs at concentrations of CAPE below 1 μM, emphasizing the physiological relevance and importance of the finding. AP-1 is a dimeric transcription factor that consists mainly of the members from the Fos and Jun subfamilies.39 AP-1 activity plays a critical role in various cellular events including

Figure 3. CAPE suppresses AP-1 activity. (A and B) CAPE inhibited AP-1 luciferase activity induced by EGF (10 ng/mL) (A) and TPA (20 ng/mL) (B) in a dose-dependent manner (n = 3). JB6 P+ cells were stably transfected with an AP-1-luciferase reporter plasmid and serum-starved in 0.1% FBS-containing MEM. The cells were pretreated with various concentrations of CAPE or CA for 1 h prior to EGF or TPA treatment. AP-1 activity was measured 4 h after EGF or TPA treatment. (C and D) TransAM AP-1 family transcription assay kit was used to detect c-Fos (C) and p-c-Jun (D) transcriptional activities, respectively (n = 3). JB6 P+ cells were stimulated with EGF for 1 h. The asterisks (*p < 0.05, **p < 0.01, and ***p < 0.001) indicate a significant difference versus the EGF-treated group. 2126

DOI: 10.1021/acs.jnatprod.7b00433 J. Nat. Prod. 2017, 80, 2124−2130

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C. cassia can attenuate AP-1 activity and may be the cause of the anticancer effects observed. We also confirmed the presence of CA in the twigs of C. cassia. However, although the two compounds share similar structures, only CAPE showed strong effects against malignant transformation, cell cycle progression, and AP-1 activity at the concentrations tested. Although CA has also been shown to possess cancer chemopreventive and chemotherapeutic effects in previous studies,52−54 when compared head-to-head in identical conditions, it appears that the presence of the phenethyl group in CA (i.e., CAPE) significantly enhances its bioactivity based on our findings and others.55 In addition, through comparing CAPE and its derivatives, a previous study has shown that the phenethyl group of CAPE is required for heme oxygenase 1 inducing activity in human umbilical vein endothelial cells.56 Identification of the direct binding target of CAPE and further research on the structure−activity relationships of these compounds could be beneficial for the design of structures with improved bioactivity. In conclusion, we have identified CAPE and CA present in the twigs of C. cassia (Cinnamomi Ramulus) using LC-MSguided isolation techniques and demonstrated that CAPE possesses strong cancer chemopreventive activity through the downregulation of c-Fos (Figure 6). These findings support the

Figure 5. CAPE induces proteosomal degradation of c-Fos. (A) CAPE does not reduce the mRNA expression of c-fos. CAPE was treated for 1 h prior to addition of EGF. After 15 min of EGF (10 ng/mL) treatment, total RNA was extracted. cDNA was synthesized and the gene expression was analyzed (n = 3). The asterisks (***p < 0.001) indicate a significant difference versus a control group without EGF. (B) JB6 P+ cells were serum-starved for 24 h and pretreated with MG132 (10 μM) and CAPE (0.5 and 1 μM), and cells were collected 15 min after EGF (10 ng/mL) treatment. The cell lysates were subjected to immunoblot analysis.

proliferation, differentiation, survival, apoptosis, and neoplastic transformation.37,38,42 Overexpression of AP-1 has been reported in various types of human cancers, and evidence has shown that AP-1 can play a crucial role in promoting tumorigenesis.37−39,42 Among the Fos family of transcription factors, only c-Fos and FosB are capable of malignant transformation.43 Overexpression of c-Fos can result in tumor development,43,44 while overexpression of other Fos and Jun members alone is generally insufficient for the generation of tumors in mice.43 Elevated expression of c-Fos is known to induce uncontrolled cell proliferation and has been detected in various human cancers.45 In our study, CAPE exerted potent chemopreventive effects and suppressed c-Fos levels. CAPE has been previously reported to possess anticancer effects,27,28,30,46−49 but the molecular mechanism responsible has not been fully understood. We observed that CAPE downregulated c-Fos through the induction of proteasomal degradation. While CAPE reduced c-Fos protein levels, it did not decrease its mRNA expression. Consistent with this finding, CAPE did not have any notable effect on the upstream ERK-RSK-CREB pathway, which is known to regulate c-Fos expression. Cinnamomi Ramulus refers to the twigs of C. cassia and has been used as an ingredient in traditional anticancer herbal medicine in Korea and China.5−7,50 On the basis of our findings, we believe that the anticancer effects observed in Cinnamomi Ramulus can be at least partially attributed to the effect of CAPE directly suppressing the proto-oncogene c-Fos. Interestingly, a study of 2-hydroxycinnamaldehyde, another compound present in C. cassia, also reported the suppression of cancer cell growth through the inactivation of AP-1.51 Although the concentration of 2-hydroxycinnamaldehyde needed to induce inhibitory effects against AP-1 was higher than that of CAPE, collectively these results show that components of

Figure 6. Proposed mechanism for the cancer chemopreventive effect of CAPE.

rationale of using this plant as an ingredient in traditional anticancer prescriptions and for the further development of CAPE as a lead for anticancer therapies.

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EXPERIMENTAL SECTION

General Experimental Procedures. IR spectra were recorded on a Bruker IFS-66/S FT-IR spectrometer. UV spectra were recorded with an Agilent 8453 UV−visible spectrophotometer (Agilent Technologies). NMR spectra were obtained from a Bruker AVANCE III 700 NMR spectrometer operating at 700 MHz (1H) and 175 MHz (13C) (Bruker), with chemical shifts given in ppm (δ). Preparative high-performance liquid chromatography (HPLC) used a Waters 1525 Binary HPLC pump with a Waters 996 photodiode array detector (Waters Corporation, Milford, CT, USA) using a 250 mm × 20 mm i.d., 10 μm, YMC-Pack ODS-AM C18(2) column (YMC America, Inc., Allentown, PA, USA). Semipreparative HPLC used a Shimadzu 2127

DOI: 10.1021/acs.jnatprod.7b00433 J. Nat. Prod. 2017, 80, 2124−2130

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116.0 (C-5), 115.7 (C-8), 114.8 (C-2); ESIMS (positive-ion mode) m/z 181 [M + H]+. Cell Culture. The JB6 P+ mouse epidermal (JB6 P+) cell line was cultured in monolayers at 37 °C in a 5% CO2 incubator in MEM containing 5% FBS and penicillin−streptomycin. Cells were cytogenetically tested and authenticated before the long-term storage in liquid nitrogen. Each vial of frozen cells was thawed and maintained in culture for a maximum of 8 weeks. Soft Agar Assay. Cells (8 × 103/mL) were suspended in 1 mL of 0.3% BME top agar containing 10% FBS with various concentrations of CAPE or CA with or without EGF (or TPA) and placed over a lower layer of solidified BME, 10% FBS, and 0.5% agar (3 mL) with the same concentrations of CAPE or CA with or without EGF (or TPA). The cultures were maintained at 37 °C in a 5% CO2 incubator for 1 to 2 weeks, and the number of colonies was counted under a microscope using Image-Pro Plus software (Media Cybernetics, Rockville, MD, USA).57 Cell Viability Assay. Cells were seeded in 96-well plates and incubated overnight before changing to 0.1% FBS-containing MEM. Indicated compounds were added, and at 24 and 48 h after the treatment, cell viability was measured using the CellTiter 96 AQueous MTS Reagent (Promega, Madison, WI, USA). Cell Cycle Assay. JB6 P+ cells were seeded (1.5 × 105 cells/well) in 60 mm dishes overnight followed by starvation with 0.1% FBScontaining MEM for 24 h. Chemicals with or without EGF were treated at various concentrations. The cells were trypsinized and then washed twice with cold PBS and fixed with ice-cold 70% ethanol at −20 °C overnight. Cells were washed twice with PBS, incubated with 20 mg/mL RNase A and 200 mg/mL propidium iodide in PBS at room temperature for 30 min avoiding light, and subjected to flow cytometry using the FACSCalibur flow cytometer.58 Luciferase Reporter Gene Assay. The JB6 P+ cell line was stably transfected with an AP-1 luciferase reporter plasmid and maintained in MEM supplemented with 5% FBS containing 200 μg/mL G418. Cells were seeded into a 96-well plate overnight. Cells were than starved in 0.1% FBS-containing MEM medium for another 24 h. The cells were treated with various concentrations of chemicals 1 h prior to EGF or TPA treatment. After 6 h, cells were harvested and disrupted with 100 mL of lysis buffer (0.1 mmol/L potassium phosphate pH 7.8, 1% Triton X-100, 1 mmol/L dithiothreitol, and 2 mmol/L EDTA). Then firefly luciferase activities were measured by Luminoskan Ascent (Thermo Fisher Scientific, Waltham, MA) using substrates provided in the reporter assay system (Promega). AP-1 Transcription Assay. c-Fos, c-jun, fosB, and Jun D activation was measured using the TransAM AP-1 family transcription assay kit (Active Motif, CA, USA) as per the manufacturer’s instructions. Briefly, cell extracts were added to a 96-well plate precoated with AP-1 binding oligonucleotides. The plate was incubated with primary antibodies specific for each AP-1 factor. Then, horseradish peroxidase-conjugated secondary antibody was added, followed by the addition of developing solution. Absorbance was measured at 450 nm. Immunoblotting. Cells were disrupted with cell lysis buffer (20 mmol/L Tris pH 7.5, 150 mmol/L NaCl, 1 mmol/L EDTA, 1 mmol/ L EGTA, 1% Triton X-100, 2.5 mmol/L sodium pyrophosphate, 1 mmol/L β-glycerophosphate, 1 mmol/L sodium vanadate, and 1 mmol/L phenylmethylsulfonyl fluoride), and the proteins were collected. The protein concentration was determined using a dyebinding protein assay kit (Bio-Rad) as described in the manufacturer’s manual. Total cellular protein extracts (10−60 μg) were separated by sodium dodecyl sulfate−polyacrylamide gel electrophoresis and electrophoretically transferred to a polyvinylidene difluoride membrane. After blocking, the membrane was incubated with a specific primary antibody at 4 °C overnight. Protein bands were visualized by a chemiluminescence detection kit after incubating with an AP-linked secondary antibody. Real-Time PCR. Total RNA was extracted from cultured cells using the RNeasy Plus Mini Kit (Qiagen, Valencia, CA, USA) following the manufacturer’s instructions. The reverse transcription reaction was done with amfiRivert cDNA Synthesis Platinum Master Mix

Prominence HPLC system with SPD-20A/20AV Series Prominence HPLC UV−vis detectors (Shimadzu, Tokyo, Japan). An Agilent 1200 Series HPLC system equipped with a diode array detector and a 6130 Series ESI mass spectrometer was used for LC-MS analysis using an analytical Kinetex C18 100 Å column (100 mm × 2.1 mm i.d., 5 μm) (Phenomenex, Torrance, CA, USA). Column chromatography used silica gel 60, 230−400 mesh, and RP-C18 silica gel, 230−400 mesh (Merck, Darmstadt, Germany). Silica gel 60 (Merck, 230−400 mesh) and reversed-phase (RP)-C18 silica gel (Merck, 230−400 mesh) were used for column chromatography. Merck precoated silica gel F254 plates and RP-18 F254s plates were used for thin-layer chromatography (TLC). Spots were detected on TLC under UV light or by heating after spraying with anisaldehyde−sulfuric acid. Chemicals and Reagents. Minimum essential medium Eagle (MEM), basal medium Eagle (BME), EGF, TPA, and the antibody against β-actin were purchased from Sigma-Aldrich (St. Louis, MO, USA). Fetal bovine serum (FBS) was purchased from Gemini Bioproducts (Sacramento, CA, USA). Antibodies to detect p-c-Fos (S32), c-Fos, p-CREB (S133), CREB, and p-RSK (T359/S363) were purchased from Cell Signaling Technology (Danvers, MA, USA). Antibodies for RSK, α-tubulin, p-ERK (T202/Y204), and ERK were purchased from Santa Cruz Biotechnology (Dallas, TX, USA). The protein assay kit was obtained from Bio-Rad Laboratories (Hercules, CA, USA). Plant Material. The twigs of C. cassia were acquired at the Kyoungdong Market, Seoul, Korea, in April 2015 and were verified by one of authors (K.H.K.). A voucher specimen (SKKU-GJ-2015-04) of materials was deposited in the herbarium of the School of Pharmacy, Sungkyunkwan University, Suwon, Korea. Extraction and Isolation. The dried and mashed twigs of C. cassia (10 g) were extracted with 100% MeOH (500 mL × 2) at room temperature for 3 days and then filtered. The crude extract was evaporated under a vacuum to afford a brown gum (1.1 g) of extract. The resultant MeOH extract was suspended in distilled H2O (500 mL) and successively partitioned with EtOAc (500 mL × 2) to provide an EtOAc-soluble extract (310 mg). The small aliquot of EtOAc-soluble extract was injected on LC/MS equipment eluted with a gradient solvent system of MeOH/H2O (1:9−1:0, flow rate of 0.3 mL/min, UV 254 nm) to identify the target constituent, caffeic acid phenethyl ester. On the basis of LC/MS data, the EtOAc-soluble extract was separated by using preparative RP-HPLC eluted with a gradient solvent system of MeOH/H2O (1:9−1:0, flow rate of 5.0 mL/min, UV 254 nm) to give five subfractions (E1−E5). All five subfractions were subjected to LC/MS prior to purification for the target isolation of CAPE (1) (3.5 mg, tR = 33.0 min), which was purified from subfraction E3 (35 mg) using semipreparative RP-HPLC with an isocratic solvent system of aqueous 39% MeOH (Phenomenex Luna C18, 250 mm × 10 mm i.d., 5 μm, flow rate of 2 mL/min, UV 254 nm). Additionally, LC/MS analysis indicated the presence of caffeic acid (2) (4.0 mg, tR = 21.0 min), which was isolated from subfraction E1 (20 mg) by utilizing semipreparative RP-HPLC with an isocratic solvent system of aqueous 19% MeOH (Phenomenex Luna C18, 250 mm × 10 mm i.d., 5 μm, flow rate of 2 mL/min, UV 254 nm). Caffeic acid phenethyl ester (1): amorphous powder; 1H NMR (CD3OD, 700 MHz) δ 7.46 (1H, d, J = 16.0 Hz, H-7), 7.21 (5H, m, H-1′, H-2′, H-3′, H-4′, and H-5′), 7.11 (1H, d, J = 2.0 Hz, H-2), 7.04 (1H, dd, J = 8.0, 2.0 Hz, H-6), 6.81 (1H, d, J = 8.0 Hz, H-5), 6.20 (1H, d, J = 16.0 Hz, H-8), 3.60 (2H, t, J = 7.5 Hz, H-8′), 2.59 (2H, t, J = 7.5 Hz, H-3′); 13C NMR (CD3OD, 175 MHz) δ 168.1 (C-9), 149.0 (C4), 148.8 (C-3), 148.4 (C-7), 138.6 (C-4′), 129.7 (C-3′ and C-5′), 128.6 (C-2′ and C-6′), 127.7 (C-1), 126.4 (C-1′), 122.2 (C-6), 115.6 (C-5), 115.0 (C-8), 114.1 (C-2), 60.6 (C-8′), 33.9 (C-7′); ESIMS (positive-ion mode) m/z 285 [M + H]+. Caffeic acid (2): amorphous powder; 1H NMR (CD3OD, 700 MHz) δ 7.36 (1H, d, J = 16.0 Hz, H-7), 7.01 (1H, d, J = 2.0 Hz, H-2), 6.95 (1H, dd, J = 8.0, 2.0 Hz, H-6), 6.75 (1H, d, J = 8.0 Hz, H-5), 6.11 (1H, d, J = 16.0 Hz, H-8); 13C NMR (CD3OD, 175 MHz) δ 171.5 (C9), 150.1 (C-4), 149.0 (C-3), 148.3 (C-7), 128.7 (C-1), 123.2 (C-6), 2128

DOI: 10.1021/acs.jnatprod.7b00433 J. Nat. Prod. 2017, 80, 2124−2130

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(GenDepot, Barker, TX, USA). Gene expression was assessed with a 7500 real-time PCR system (Applied Biosystems, Carlsbad, CA, USA) using the Power SYBR Green Master Mix (Life Technology, Grand Island, NY, USA). Reaction strips were incubated in a 96-well thermal cycling plate at 95 °C for 10 min and then underwent 40 cycles of 15 s at 95 °C and 1 min at 59 °C. All reactions were performed in triplicate. The following primers were used to detect expression: c-fos: 5′AGAGCATCAGCAACGTGGAG-3′ (sense), 5′-GGAAGGAGTCAGCTTCAGGG-3′ (antisense); gapdh: 5′-GGCATGGCCTTCCGTGT-3′ (sense), 5′-GGTTTCTCCAGGCGGCA-3′ (antisense). Statistical Analysis. One-way ANOVA was used to compare between groups with GraphPad Prism (v.6) software. As indicated in each figure legend, data are presented as mean values ± standard deviation. P values are indicated with an asterisk (*) and/or pound sign (#) (*P < 0.05; **P < 0.01; and ***P < 0.001).

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(9) Kim, K. H.; Moon, E.; Choi, S. U.; Pang, C.; Kim, S. Y.; Lee, K. R. J. Ethnopharmacol. 2015, 162, 231−237. (10) Kim, C. S.; Moon, E.; Choi, S. U.; Kim, S. Y.; Lee, K. R.; Kim, K. H. J. Antibiot. 2015, 68, 414−416. (11) Kang, H. R.; Eom, H. J.; Lee, S. R.; Choi, S. U.; Kang, K. S.; Lee, K. R.; Kim, K. H. Nat. Prod Commun. 2015, 10, 1929−1932. (12) Lee, S.; Moon, E.; Choi, S. U.; Kim, K. H. Chem. Biodiversity 2016, 13, 1391−1396. (13) Lee, S. R.; Park, J. Y.; Yu, J. S.; Lee, S. O.; Ryu, J. Y.; Choi, S. Z.; Kang, K. S.; Yamabe, N.; Kim, K. H. J. Agric. Food Chem. 2016, 64, 3804−3809. (14) Yu, J. S.; Baek, J.; Park, H. B.; Moon, E.; Kim, S. Y.; Choi, S. U.; Kim, K. H. Arch. Pharmacal Res. 2016, 39, 1628−1634. (15) He, S.; Zeng, K. W.; Jiang, Y.; Tu, P. F. Fitoterapia 2016, 112, 153−160. (16) Liang, M.-T.; Yang, C.-H.; Li, S.-T.; Yang, C.-S.; Chang, H.-W.; Liu, C.-S.; Cham, T.-M.; Chuang, L.-Y. Eur. Food Res. Technol. 2008, 227, 1387−1396. (17) Liao, J. C.; Deng, J. S.; Chiu, C. S.; Hou, W. C.; Huang, S. S.; Shie, P. H.; Huang, G. J. Evid Based Complement Alternat Med. 2012, 2012, 429320. (18) Hwang, S. H.; Choi, Y. G.; Jeong, M. Y.; Hong, Y. M.; Lee, J. H.; Lim, S. Gene 2009, 443, 83−90. (19) Jung, J.; Lee, J. H.; Bae, K. H.; Jeong, C. S. Yakugaku Zasshi 2011, 131, 1103−1110. (20) Liang, K.; Cui, S.; Zhang, Q.; Bi, K.; Qian, Z.; Jia, Y. Zhongguo Zhong Yao Za Zhi 2011, 36, 3298−3301. (21) Yang, L.; Zhao, Q.-c.; Tan, J.-j.; Shang, Z.-p.; DU, Z.-q. Practical Pharmacy Clin. Remedies 2010, 13, 183−184. (22) Chang, W. L.; Cheng, F. C.; Wang, S. P.; Chou, S. T.; Shih, Y. Environ. Toxicol. 2017, 32, 456−468. (23) Liu, R.; He, T.; Zeng, N.; Chen, T.; Gou, L.; Liu, J. Med. Plant 2015, 6, 4. (24) Sui, F.; Lin, N.; Guo, J. Y.; Zhang, C. B.; Du, X. L.; Zhao, B. S.; Liu, H. B.; Yang, N.; Li, L. F.; Guo, S. Y.; Huo, H. R.; Jiang, T. L. J. Asian Nat. Prod. Res. 2010, 12, 76−87. (25) Kim, E. C.; Kim, H. J.; Kim, T. J. Biosci., Biotechnol., Biochem. 2015, 79, 617−624. (26) Rad, S. K.; Kanthimathi, M. S.; Abd Malek, S. N.; Lee, G. S.; Looi, C. Y.; Wong, W. F. PLoS One 2015, 10, e0145216. (27) Akyol, S.; Ozturk, G.; Ginis, Z.; Armutcu, F.; Yigitoglu, M. R.; Akyol, O. Nutr. Cancer 2013, 65, 515−526. (28) Lin, H. P.; Lin, C. Y.; Huo, C.; Hsiao, P. H.; Su, L. C.; Jiang, S. S.; Chan, T. M.; Chang, C. H.; Chen, L. T.; Kung, H. J.; Wang, H. D.; Chuu, C. P. Oncotarget 2015, 6, 6684−6707. (29) Nomura, M.; Kaji, A.; Ma, W.; Miyamoto, K.; Dong, Z. Mol. Carcinog. 2001, 31, 83−89. (30) Tolba, M. F.; Esmat, A.; Al-Abd, A. M.; Azab, S. S.; Khalifa, A. E.; Mosli, H. A.; Abdel-Rahman, S. Z.; Abdel-Naim, A. B. IUBMB Life 2013, 65, 716−729. (31) Pellati, F.; Orlandini, G.; Pinetti, D.; Benvenuti, S. J. Pharm. Biomed. Anal. 2011, 55, 934−948. (32) Damtoft, S.; Jensen, S. R. Phytochemistry 1994, 37, 441−443. (33) Jaikang, C.; Chaiyasut, C.; Narongchai, P.; Niwatananun, K.; Narongchai, S.; Kusirisin, W. Med. Chem. 2011, 7, 99−105. (34) Lindsey, S.; Langhans, S. A. Int. Rev. Cell Mol. Biol. 2015, 314, 1−41. (35) Enomoto, T.; Yamasaki, H. Cancer Res. 1985, 45, 2681−2688. (36) Perera, F. P. Environ. Health Perspect 1991, 94, 231−235. (37) Matthews, C. P.; Colburn, N. H.; Young, M. R. Curr. Cancer Drug Targets 2007, 7, 317−324. (38) Angel, P.; Karin, M. Biochim. Biophys. Acta, Rev. Cancer 1991, 1072, 129−157. (39) Vaiopoulos, A. G.; Papachroni, K. K.; Papavassiliou, A. G. Int. J. Biochem. Cell Biol. 2010, 42, 1061−1065. (40) Wang, Y.; Prywes, R. Oncogene 2000, 19, 1379−1385. (41) Ahn, S.; Olive, M.; Aggarwal, S.; Krylov, D.; Ginty, D. D.; Vinson, C. Mol. Cell. Biol. 1998, 18, 967−977.

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jnatprod.7b00433. Additional information (PDF)

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AUTHOR INFORMATION

Corresponding Authors

*Tel (K. H. Kim): +82-31-290-7700. Fax: +82-31-290-7730. Email: [email protected]. *Tel (S. Byun): +82-32-835-8033. Fax: +82-32-835-0804. Email: [email protected]. ORCID

Ki Hyun Kim: 0000-0002-5285-9138 Sanguine Byun: 0000-0003-3903-5887 Author Contributions ∥

S. H. Shin and S. R. Lee contributed equally to this work.

Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This research was supported by the Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Science, ICT, & Future Planning (2015R1C1A1A02037383) to K.H.K. This work was supported by the NRF grant funded by the Korea government (MSIP) (NRF-2017R1C1B1006072) to S.B.

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REFERENCES

(1) Torre, L. A.; Bray, F.; Siegel, R. L.; Ferlay, J.; Lortet-Tieulent, J.; Jemal, A. Ca-Cancer J. Clin. 2015, 65, 87−108. (2) Vineis, P.; Schatzkin, A.; Potter, J. D. Carcinogenesis 2010, 31, 1703−1709. (3) Danaei, G.; Vander Hoorn, S.; Lopez, A. D.; Murray, C. J.; Ezzati, M. Lancet 2005, 366, 1784−1793. (4) Wu, S.; Powers, S.; Zhu, W.; Hannun, Y. A. Nature 2016, 529, 43−47. (5) Jang, S. B.; Baek, S. E.; Choi, K. H.; Yoo, J. E. J. Korean Obstet. Gynecol 2016, 29, 99−112. (6) Park, W. H.; Joo, S. T.; Park, K. K.; Chang, Y. C.; Kim, C. H. Immunopharmacol. Immunotoxicol. 2004, 26, 103−112. (7) Wang, T. C.; Fang, C. N.; Shen, C. C.; Wei, H. Y.; Weng, Y. P.; Lin, J. Y.; Hsieh-Li, H. M.; Lee, C. Y. Evid Based Complement Alternat Med. 2012, 2012, 276032. (8) Kim, K. H.; Kim, C. S.; Park, Y. J.; Moon, E.; Choi, S. U.; Lee, J. H.; Kim, S. Y.; Lee, K. R. Bioorg. Med. Chem. Lett. 2015, 25, 96−99. 2129

DOI: 10.1021/acs.jnatprod.7b00433 J. Nat. Prod. 2017, 80, 2124−2130

Journal of Natural Products

Article

(42) Papoudou-Bai, A.; Hatzimichael, E.; Barbouti, A.; Kanavaros, P. Clin. Exp. Med. 2016, DOI: 10.1007/s10238-016-0436-z. (43) Wisdon, R.; Verma, I. M. Mol. Cell. Biol. 1993, 13, 7429−7438. (44) Muhammad, N.; Bhattacharya, S.; Steele, R.; Phillips, N.; Ray, R. B. Clin. Cancer Res. 2017, 23, 3120. (45) Milde-Langosch, K. Eur. J. Cancer 2005, 41, 2449−2461. (46) Ha, J.; Choi, H. S.; Lee, Y.; Lee, Z. H.; Kim, H. H. Int. Immunopharmacol. 2009, 9, 774−780. (47) Huang, M. T.; Ma, W.; Yen, P.; Xie, J. G.; Han, J.; Frenkel, K.; Grunberger, D.; Conney, A. H. Carcinogenesis 1996, 17, 761−765. (48) Zheng, Z. S.; Xue, G. Z.; Grunberger, D.; Prystowsky, J. H. Oncol. Res. 1995, 7, 445−452. (49) Koltuksuz, U.; Mutus, H. M.; Kutlu, R.; Ozyurt, H.; Cetin, S.; Karaman, A.; Gurbuz, N.; Akyol, O.; Aydin, N. E. J. Pediatr Surg 2001, 36, 1504−1509. (50) Lu, C. C.; Lin, M. Y.; Chen, S. Y.; Shen, C. H.; Chen, L. G.; Hsieh, H. Y.; Chan, M. W.; Hsu, C. D. BMC Complementary Altern. Med. 2013, 13, 44. (51) Lee, C. W.; Lee, S. H.; Lee, J. W.; Ban, J. O.; Lee, S. Y.; Yoo, H. S.; Jung, J. K.; Moon, D. C.; Oh, K. W.; Hong, J. T. J. Pharmacol. Sci. 2007, 104, 19−28. (52) Rosendahl, A. H.; Perks, C. M.; Zeng, L.; Markkula, A.; Simonsson, M.; Rose, C.; Ingvar, C.; Holly, J. M.; Jernstrom, H. Clin. Cancer Res. 2015, 21, 1877−1887. (53) Yang, G.; Fu, Y.; Malakhova, M.; Kurinov, I.; Zhu, F.; Yao, K.; Li, H.; Chen, H.; Li, W.; Lim, D. Y.; Sheng, Y.; Bode, A. M.; Dong, Z.; Dong, Z. Cancer Prev. Res. 2014, 7, 1056−1066. (54) Kang, N. J.; Lee, K. W.; Shin, B. J.; Jung, S. K.; Hwang, M. K.; Bode, A. M.; Heo, Y. S.; Lee, H. J.; Dong, Z. Carcinogenesis 2009, 30, 321−330. (55) Dziedzic, A.; Kubina, R.; Kabala-Dzik, A.; Tanasiewicz, M. Evid Based Complement Alternat Med. 2017, 2017, 6793456. (56) Wang, X.; Stavchansky, S.; Kerwin, S. M.; Bowman, P. D. Eur. J. Pharmacol. 2010, 635, 16−22. (57) Byun, S.; Shin, S. H.; Park, J.; Lim, S.; Lee, E.; Lee, C.; Sung, D.; Farrand, L.; Lee, S. R.; Kim, K. H.; Dong, Z.; Lee, S. W.; Lee, K. W. Mol. Nutr. Food Res. 2016, 60, 1068−1078. (58) Shin, S. H.; Seo, S. G.; Min, S.; Yang, H.; Lee, E.; Son, J. E.; Kwon, J. Y.; Yue, S.; Chung, M. Y.; Kim, K. H.; Cheng, J. X.; Lee, H. J.; Lee, K. W. J. Agric. Food Chem. 2014, 62, 4306−4312.

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