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ScienceDirect Journal of Nutritional Biochemistry 26 (2015) 345 – 350
α-Tocopherol suppresses antiangiogenic effect of δ-tocotrienol in human umbilical vein endothelial cells☆ Akira Shibata a , Kiyotaka Nakagawa a,⁎, Tsuyoshi Tsuduki b , Teruo Miyazawa a a b
Food and Biodynamic Chemistry Laboratory, Graduate School of Agricultural Science, Tohoku University, Sendai 981-8555, Japan Laboratory of Food and Biomolecular Science, Graduate School of Agricultural Science, Tohoku University, Sendai 981-8555, Japan
Received 29 July 2014; received in revised form 8 November 2014; accepted 17 November 2014
Abstract Recently, tocotrienol (T3), a less well-known form of vitamin E, has gained attention as a potent hypocholesterolemic, anticancer and antiangiogenic agent. However, tocopherol (Toc), a commonly consumed form of vitamin E, has been reported to inhibit T3’s effects (hypocholesterolemic and anticancer activity). There has been no report on Toc’s effect on the antiangiogenic action of T3 during cotreatment. The aim of this study is to determine if and to what extent Toc affects the antiangiogenic effects of δ-T3 (the most potent isomer). This was achieved through cotreatment of human umbilical vein endothelial cells (HUVECs) with δ-T3 and Toc (α-, β-, γ- and δ-isomers). Toc, especially α-Toc, attenuated δ-T3-induced cytotoxicity and tube degradation in cotreated HUVECs, while α-Toc treatments did not exhibit any effects. A rat aortic ring assay also showed inhibition of δ-T3’s antiangiogenic effects by α-Toc. Further, in HUVEC study, cell cycle arrest and proapoptotic gene expression (p21, p27, caspase-3 and caspase-9) which were induced by δ-T3 were decreased by α-Toc treatment. α-Toc also suppressed δ-T3-induced dephosphorylation of vascular endothelial growth factor receptor 2 and Akt pathway proteins. Additionally, uptake of δ-T3 into HUVECs was decreased by α-Toc. Here we demonstrate that α-Toc not only has little antiangiogenic effect on endothelial cells but also reduces the antiangiogenic effects of δ-T3 through modulation of its cellular uptake and of relevant signal transduction pathways. Understanding T3’s antiangiogenic effects and interaction with Toc is important for developing medical applications. © 2015 Elsevier Inc. All rights reserved. Keywords: Angiogenesis; Tocotrienol; Tocopherol; VEGF; HUVEC
1. Introduction Angiogenesis takes place after activation of endothelial cells by angiogenic stimuli, and it can promote cancer growth by supplying nutrients and oxygen and removing waste products [1]. Therefore, antiangiogenesis is an effective defensive strategy against cancer progression [2]. Various anticancer agents that target angiogenesis have been developed, including natural and synthetic compounds [3]. Previous studies have shown that tocotrienol (T3) (especially the δ-isoform), an unsaturated vitamin E, has anticancer and antiangiogenic properties
Abbreviations: DLD-1, human colorectal adenocarcinoma; ERK, extracellular regulated kinase; HPLC, high-performance liquid chromatography; HUVEC, human umbilical vein endothelial cell; PTEN, phosphatase and tensin homolog deleted from chromosome 10; RT-PCR, reverse transcription polymerase chain reaction; T3, tocotrienol; Toc, tocopherol; VEGF, vascular endothelial growth factor; VEGFR-2, vascular endothelial growth factor receptor-2; WST-1, water-soluble tetrazolium salt ☆ Conflicts of interest: none. ⁎ Corresponding author at: Food and Biodynamic Chemistry Laboratory, Graduate School of Agricultural Science, Tohoku University, 1-1 TsutsumidoriAmamiyamachi, Aoba-ku, Sendai 981-8555, Japan. Tel.: +81 22 717 8906; fax: +81 22 717 8905. E-mail address: [email protected] (K. Nakagawa). http://dx.doi.org/10.1016/j.jnutbio.2014.11.010 0955-2863/© 2015 Elsevier Inc. All rights reserved.
in vitro and in animal models [4–7]. Many studies have demonstrated that T3 has potent proapoptotic and antiproliferative activities in cancer cells (e.g., breast, colon and liver cancer cells) [8–10]. Importantly, at the same T3 dose, almost no adverse effects on growth or function are seen in normal counterpart cells [11]. Previous studies by our group have shown that δ-T3 has the most potent antiangiogenic activities in vitro and in vivo. Conversely, tocopherol (Toc), a major form of vitamin E found in many foods, does not [12]. Therefore, δ-T3 would be a promising anticancer agent or an adjuvant for minimizing tumor angiogenesis. Recently, we showed that δ-T3-induced cytotoxicity was attenuated by α-Toc through inhibition of cellular uptake of δ-T3 into colon cancer cells [13]. Results from the study suggest that α-Toc inhibits a physiological function of δ-T3. Thus, there is a possibility that α-Toc may attenuate the antiangiogenic activity of δ-T3. However, the antiangiogenic effect of δ-T3 during cotreatment of these compounds is unknown. If α-Toc inhibits the effects of δ-T3, careful consideration is required to predict the antiangiogenic effects of δ-T3. In the present study, we cotreated human umbilical vein endothelial cells (HUVECs) with δ-T3 and Toc (α-, β-, γ- and δ-isomers) and investigated whether Toc inhibits the antiangiogenic effects of T3. Cytotoxicity, tube formation, apoptosis and cell-cycle-related gene expression, phosphorylation of signal proteins and cellular T3 accumulation were evaluated to explore the relationship between these compounds. To further examine whether Toc inhibits the antiangiogenic
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effect of T3 in a more physiological condition, a rat aortic ring assay was also performed.
Table 1 Primer sequences for quantitative real-time PCR amplification of selected human genes. Gene
GenBank (accession no.)
Sequence (5’–3’)
p21(CDKN1A)-F p21(CDKN1A)-R p27(CDKN1B)-F p27(CDKN1B)-R Caspase-3(CASP3)-F Caspase-3(CASP3)-R Caspase-9(CASP9)-F Caspase-9(CASP9)-R β-Actin(ACTB)-F β-Actin(ACTB)-R
NM_000389 NM_000389 NM_004064 NM_004064 NM_032991 NM_032991 NM_032996 NM_032996 NM_001101 NM_001101
AGCAGAGGAAGACCATGTGGAC TTTCGACCCTGAGAGTCTCCAG TGCAACCGACGATTCTTCTACTCAA CAAGCAGTGATGTATCTGATAAACAAGGA ATGGAAGCGAATCAATGGAC GGCTCAGAAGCACACAAACA TGCTCAGACCAGAGATTCGC CTTTCTGCTCGACATCACCAA TGGCACCCAGCACAATGAA CTAAGTCATAGTCCGCCTAGAAGCA
2. Materials and methods 2.1. Chemicals All Toc isomers (α-, β-, γ- and δ-Toc) were purchased from Eisai (Tokyo, Japan). δ-T3 was kindly gifted by Tama Biochemical (Tokyo, Japan). δ-T3 was selected for this study because it has been reported to show the most potent antiangiogenic activities among the four T3 isomers (α-, β-, γ- and δ-T3) [14]. Water-soluble tetrazolium salt-1 (WST-1) reagent was obtained from Dojindo Laboratories (Kumamoto, Japan). All other reagents used in this study were of analytical grade. 2.2. Cells and culture HUVECs were obtained from Kurabo (Osaka, Japan). The cells were cultured in a base medium (HuMedia-EB2) supplemented with 2% fetal bovine serum, 10 μg/L human epidermal growth factor, and 5 μg/L human basic fibroblast growth factor (Kurabo). Confluent HUVECs (passages 5 to 8) were used in all experiments. The cells were maintained at 37°C in a humidified atmosphere of 95% air and 5% CO2.
lengths were counted in four randomly selected microscopic fields on day 12 by using Lumina Vision software (Olympus, Tokyo, Japan). 2.8. Total RNA isolation and mRNA analysis
2.3. Preparation of test medium Each Toc isomer or δ-T3 was dissolved in ethanol at a concentration of 50 mM. The stock solution was then diluted with 1%–2% fetal bovine serum/HuMedia-EB2 with or without 10 μg/L vascular endothelial growth factor (VEGF) (R&D system, Minneapolis, MN, USA) to achieve the desired final concentration of vitamin E (0–50 μM). The final concentration of ethanol in the test medium was less than 0.1% (v/v), which did not affect cell viability. Medium with ethanol alone was similarly prepared and used as control medium.
After treating HUVECs with the test medium for 24 h, total RNA was isolated with the RNeasy Plus Mini kit (Qiagen, Valencia, CA, USA) for reverse transcription polymerase chain reaction (RT-PCR). cDNA was synthesized using a Ready-To-Go T-Primed FirstStrand kit (GE Healthcare, Piscataway, NJ, USA). Quantitative RT-PCR amplification was performed with the DNA Engine Opticon 2 system (MJ Research, Waltham, MA, USA) using SYBR Premix Ex Taq II (Takara Bio, Otsu, Japan) and gene specific primers for p21 (CDKN1A), p27 (CDKN1B), caspase-3 (CASP3), caspase-9 (CASP9) and β-actin (ACTB) (Table 1). PCR conditions for these primers were 95°C for 1 min, 95°C for 5 s and 60°C for 20 s over 40 cycles.
2.4. Cell viability assay HUVECs were plated at 3×103 cells/well in 96-well culture plates to determine cell viability. After 24 h, the medium was replaced with the test medium containing the compounds of interest (δ-T3 and Toc). Cells were incubated in test medium for 24 h, and cell viability was determined using the reagent WST-1 [15] according to the manufacturer’s instruction. 2.5. Tube formation assay Twenty-four-well plates were coated with 350 μl of Matrigel (Becton Dickinson, Bedford, MA, USA), and incubated at 37°C for 1 h for solidification. HUVECs were suspended in test medium containing the appropriate concentration of δ-T3 and α-Toc at 6×104 cells/well (500 μl/well). The cell suspension was plated onto the surface of the Matrigel and incubated for 18 h. Following incubation, the cells were fixed with 4% paraformaldehyde solution and photographed. Lengths of tube-structured cells were quantified using a Kurabo Angiogenesis Imaging Analyzer (imaging software; Kurabo).
2.9. Western blot analysis Confluent HUVECs were preincubated in a VEGF-free test medium for 6 h, then stimulated by the addition of 10 ng/ml VEGF and incubated for 5 min. Cellular proteins (60 μg/well) were separated by sodium dodecyl sulfate polyacrylamide gel electrophoresis (4%–20% e-PAGEL; Atto, Tokyo, Japan), and protein bands were transferred to polyvinylidene fluoride membranes (GE Healthcare). Following blocking, the membranes were incubated with primary antibodies for phospho-Akt (Ser473), phospho-Akt (Thr308), Akt, phospho-extracellular signal-regulated kinase (ERK), ERK, phospho-phosphatase and tensin homologue deleted on chromosome 10 (PTEN), PTEN, phospho-vascular endothelial growth factor receptor-2 (VEGFR-2) and β-actin, followed by a horseradishperoxidase-conjugated secondary antibody. All antibodies were purchased from Cell Signaling Technology (Beverly, MA, USA). ECL Plus reagents (GE Healthcare) were used for detection. 2.10. Apoptosis assay
2.6. Ethics statement The animal study was performed according to the protocols approved by the Institutional Committee for Use and Care of Laboratory Animals of Tohoku University, which was granted by Tohoku University Ethics Review Board (2008-Noudou-30), and the Guide for Care and Use of Laboratory Animals published by the US National Institutes of Health (NIH publication 85-23, revised 1996). All animals were anesthetized to minimize their suffering. 2.7. Rat aortic ring assay Ex vivo angiogenesis was studied by culturing rings of rat aorta in collagen gel [16]. Male Wistar rats (body weight ~200 g) were obtained from CLEA (Tokyo, Japan) and maintained in a clean, pathogen-free environment. The rats were housed individually in a temperature- (23°C) and humidity-controlled room with a 12-h light–dark cycle. Prior to the assay, each rat was acclimatized for 1 week. Once acclimatized, rats were sacrificed by bleeding from the right femoral artery under diethyl ether anesthesia. A section of thoracic aorta was removed and thoroughly washed with RPMI-1640 medium (Sigma, St. Louis, MO, USA) to remove any traces of blood. The aortic section was then turned inside out and cut into short segments of about 1.0–1.5 mm. A collagen gel matrix solution was made using 8 vol of porcine tendon collagen solution (3 mg/ml) (Nitta Gelatin, Osaka, Japan), 1 vol of 10× Eagle's MEM (GIBCO Laboratories, BRL, Rockville, MD, USA) and 1 vol of reconstitution buffer (0.08 M NaOH and 200 mM HEPES) and mixed gently at 4°C. Each aortic segment was placed in the center of a well on a 12-well culture plate and covered with 0.3 ml of prepared gel matrix solution. The solution was allowed to gel at 37°C for 20 min and was then overlaid with 1 ml of test medium (EGM2 medium; Clonetics, San Diego, CA, USA) containing 20 μg/L VEGF, δ-T3 and α-Toc. The plates were incubated for 12 days at 37°C in a fully humidified system of 5% CO2. The medium was changed on days 5 and 10 of the culture. The tube-structure
HUVECs were seeded 1×104 cells/well in gelatin-treated 96-well culture plates with culture medium and incubated until the cells reached confluence. After incubation, the medium was replaced with the test medium containing the compounds (δ-T3 and α-Toc). Cells were incubated in test medium for 6 h, and apoptosis ratio was evaluated. APOPercentage Dye kit (Biocolor Ltd., Belfast, UK) was used to quantify apoptosis, according to the manufacturer's instructions. This assay uses a dye that stains cells as they undergo the membrane ‘flip-flop’ event when phosphatidylserine is translocated to the outer membrane. This event is considered to be diagnostic of apoptosis but not necrosis. Digital images of APOPercentage-Dye-labeled cells, which appear bright pink against a white background, were used to quantify apoptotic cell ratio. Using Adobe Photoshop, the level of apoptosis was measured and expressed as a pixel number. 2.11. Cellular uptake of δ-T3 Cellular vitamin E content was determined by high-performance liquid chromatography (HPLC) with fluorescence detection. After confluent HUVECs were incubated with test medium for 6 h, cellular T3 and Toc were extracted as described previously [17]. HPLC was performed at 35°C using a silica column (ZORBAX Rx-SIL, 4.6×250 mm; Agilent, Palo Alto, CA, USA) with a mobile phase of hexane/1,4-dioxane/2-propanol (988:10:2) at a flow rate of 1.0 ml/min. T3 and Toc were detected with an RF-10AXL fluorescence detector (excitation 294 nm, emission 326 nm; Shimadzu, Kyoto, Japan). 2.12. Statistical analysis The data are expressed as the mean±S.D. Statistical analyses were performed using analysis of variance followed by Bonferroni/Dunn tests for multiple comparisons. Differences were considered significant at Pb.05.
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Fig. 1. Toc decreases δ-T3-induced reduction of HUVEC viability (A) and tube degradation (B). HUVECs were treated simultaneously with the indicated concentrations of δ-T3 and one of four Toc isomers, and incubated for 24 h. Cell viabilities were determined by WST-1 assay postincubation (A). Following incubation of the cells on a Matrigel-coated plate for 18 h, cultures were photographed, and the lengths of tube-structured cells were quantified using angiogenesis imaging software (B). Cell viability and tube length are expressed as the percentage to control. Values are mean±S.D. (n=6). Means without a common letter differ, Pb.05.
3. Results 3.1. Toc suppresses δ-T3-induced cytotoxicity and tube degradation δ-T3 showed an approximately 40% reduction of cell viability at 5 μM compared with vehicle in HUVECs following a 24 h incubation. When Toc isomers (α-, β-, γ- and δ-Toc) over a range of concentrations (5–50 μM) were cotreated with δ-T3, all exhibited a dose-dependent suppression of δ-T3-induced cytotoxicity in HUVECs. Of the isomers, α-, β- and γ-Toc completely suppressed the effect of δ-T3 at 50 μM (Fig. 1A). Because α-Toc showed the most potent inhibitory effect among the four Toc isomers, we used α-Toc for the following experiments. Effects of α-Toc on δ-T3-induced tube degradation were assessed by using a Matrigel assay. When HUVECs were cultured on Matrigel in the presence of test medium containing VEGF, elongated and robust tube-like structures formed when compared to control-treated cells (Fig. 1B). Treatment with δ-T3 potently inhibited the tube formation in the VEGF-treated group. Although α-Toc alone did not exhibit any effects, it dose-dependently attenuated δ-T3-induced tube degradation. To further examine whether α-Toc inhibits the antiangiogenic effect of δ-T3 in a more representative physiological environment, a rat aortic ring assay was conducted. This assay closely reflects several stages in angiogenesis, including endothelial cell proliferation, migration and tube formation. Here, VEGF significantly increased vessel sprouting
Fig. 2. α-Toc attenuates inhibition of tube formation by δ-T3 in rat aorta. Aortic segments were embedded in collagen gel layers and incubated with EBM-2 medium in the absence or presence of VEGF, δ-T3 and α-Toc. Medium was changed on day 5 and day 10. After 12 days, cultures were photographed (A) and vessel lengths were quantified (B). Values are mean±S.D. (n=4). Means without a common letter differ, Pb.05.
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compared with the control, and δ-T3 potently reduced microvessel growth in the VEGF-treated group (Fig. 2A and B). While α-Toc did not exhibit any effects even at 50 μM, it dose-dependently suppressed the antiangiogenic effect of δ-T3 in an ex vivo model. 3.2. α-Toc attenuates δ-T3-induced proapoptotic/cell cycle arrest gene expression and antiangiogenic signaling To elucidate the mechanism of α-Toc in suppressing the antiangiogenic effect of δ-T3, we measured the expression of genes involved in cell cycle and apoptosis. Quantitative RT-PCR showed that α-Toc decreases, in a dose-dependent manner, the expression of δ-T3-induced cell cycle arrest genes p21 (CDKN1A) and p27 (CDKN1B), and proapoptotic genes caspase-3 (CASP3) and caspase-9 (CASP9) (Fig. 3A). Gene expression changes induced by δ-T3 were completely suppressed by 50 μM of α-Toc.
α-Toc alone had no significant effect on gene expression when compared to control. Next, we investigated the VEGFR-2/Akt pathway, which includes PTEN and ERK, and which is critical for VEGF-induced angiogenic steps (e.g., cell proliferation, tube formation and antiapoptosis) [18]. Western blot analysis showed that VEGF-treated cells showed increased phosphorylation of Akt (Thr308 and Ser473), PTEN, ERK and VEGFR-2 when compared with control, whereas δ-T3 reduced VEGF-induced phosphorylation of these proteins (Fig. 3B). This effect was attenuated in a dose-dependent manner by α-Toc, causing an overall decrease of dephosphorylation of the proteins affected by δ-T3, and 50 μM of α-Toc almost completely inhibited the effects of δ-T3. α-Toc alone did not exhibit any effects on protein phosphorylation, and neither δ-T3 nor α-Toc changed total protein levels. Next, the cells were examined using the APOPercentage Dye kit to evaluate whether the apoptotic event was
Fig. 3. α-Toc rescues δ-T3-induced changes on mRNA expression (A) and protein (B) phosphorylation in HUVECs. HUVECs were treated for 24 h with the indicated concentrations of δ-T3 and α-Toc. mRNA levels of p21 (CDKN1A), p27 (CDKN1B), caspase-3 (CASP3) and caspase-9 (CASP9) were measured by real-time RT-PCR (A). Values are mean±S.D. (n=6). Means without a common letter differ, Pb.05. Protein phosphorylation levels of VEGFR-2/Akt pathway were analyzed by Western blotting (B). β-Actin acted as a loading control. Each blot is a representative of three replicate experiments.
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Fig. 4. α-Toc suppresses δ-T3-induced proapoptotic event in HUVECs. APOPercentage assay was used to assess the degree of apoptosis as described in Materials and Methods. The level of apoptosis was measured by a pixel number using Adobe Photoshop. Fold increase in apoptosis levels was expressed as the pixel numbers relative to control. Values are mean±S.D. (n=4–6). Means without a common letter differ, Pb.05.
caused by δ-T3. As shown in Fig. 4, HUVECs underwent flip-flop by treating δ-T3 for 6 h. δ-T3-induced proapoptotic event was suppressed by cotreatment with α-Toc in a dose-dependent manner, whereas α-Toc alone did not exhibit apoptotic effect. 3.3. α-Toc suppresses cellular uptake of δ-T3 on HUVECs To further elucidate the mechanism of the inhibitory effect of α-Toc on the antiangiogenic effects of δ-T3 in HUVECs, we measured cellular δT3 and α-Toc levels. Intracellular concentration of δ-T3 was 29.8± 1.5 nmol/mg protein, when HUVECs were incubated with 5 μM of δ-T3 for 6 h. This was significantly higher than in cotreated HUVECs, where δT3 (5 μM treatment) and α-Toc (50 μM treatment) had intracellular concentrations of 21.1±1.1 nmol/mg protein and 49.1±7.6 nmol/mg protein, respectively, following a 6 h incubation. Further, α-Toc decreased the uptake of δ-T3 into HUVECs in a dose-dependent manner (Fig. 5). This decrease in uptake is in concordance with the suppression of the antiangiogenic action of δ-T3 by adding α-Toc. When treated with 50 μM α-Toc alone, the intracellular concentration was 64.9± 3.3 nmol/mg protein. 4. Discussion The goal of this study was to clarify influence of Toc on T3-induced antiangiogenesis in vitro and in other relevant systems. This study demonstrated that α-Toc attenuates δ-T3's antiangiogenic response
Fig. 5. α-Toc prevents cellular uptake of δ-T3 in HUVECs. HUVECs were treated simultaneously for 6 h with the indicated concentrations of δ-T3 and α-Toc. Cellular contents of δ-T3 and α-Toc were measured by HPLC with fluorescence detection. Values are means±S.D. (n=6).
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(proliferation and tube formation) via suppression of δ-T3-induced protein dephosphorylation and reduction of cellular δ-T3 uptake. As shown in Fig. 1A, all Toc isomers attenuated δ-T3-induced cytotoxicity and did not show any inhibitory effects (as seen in δ-T3) on HUVEC viability. In this study, the inhibitory potency of the isomers against δ-T3 effects varied in the order α-Nγ-Nβ-Nδ-Toc. It has previously been shown that γ-Toc and δ-Toc induce apoptosis in some cancer cells, but α-Toc does not show this effect [19]. Cellular uptake of γ-Toc and δ-Toc was greater compared with that of α-Toc in intestinal epithelial cell lines, which in part explains the superior biopotency for both γ-Toc and δ-Toc compared with α-Toc in these cells [20]. Further, structurally, the δ-isomer lacks the 5- and 7-methyl groups attached onto its chroman ring; so it easily passes through cell membranes more than the α-isomer. This information seems to reflect the aforementioned order (α-Nγ-Nβ-Nδ-Toc) of isomer potency observed in our study. Similar results were obtained with our previous studies using human colon cancer cells [13]. We also aimed to determine whether α-Toc affects δ-T3's antiangiogenic effect on the key steps of angiogenesis, namely, tube formation. Here, α-Toc dose-dependently attenuated δ-T3-induced tube degradation. Importantly, α-Toc did not have any inhibitory effects on the tube formation in vitro or on vessel sprouting ex vivo (Figs. 1B, 2A and B). Concordant to these phenotypic results (Figs. 1–2), α-Toc dosedependently attenuated δ-T3-induced cell cycle arrest and proapoptotic signals at the mRNA level (Fig. 3A). Furthermore, we performed APOPercentage apoptosis assay and confirmed that α-Toc suppressed δ-T3-induced proapoptotic event in HUVECs (Fig. 4). These observations suggest that α-Toc suppresses the upstream events of δ-T3-induced cell cycle arrest or apoptosis. These genes are regulated by intracellular signal transduction from growth factor receptors such as the VEGFR-2/Akt pathway, which is involved in tumor angiogenesis and is correlated with tumor progression [21]. Activated Akt promotes cell survival and protects against apoptosis through multiple mechanisms. This includes regulating p21 and p27, and protein degradation or suppression of activation for caspase-3 and caspase-9 [22]. Some apoptotic inducers such as 7-ketocholesterol have been shown to induce cell death through inactivation of the phosphatidylinositol 3-kinase/Akt signaling pathway, which is known to be highly specific to lipid raft domains, and this event was canceled by α-Toc [23]. Therefore, we speculated that α-Toc may suppress δ-T3's antiangiogenic effect via regulating VEGFR-2/Akt activity in HUVECs. δ-T3 inhibited VEGF-induced phosphorylation of Akt (Thr308 and Ser473), PTEN and VEGFR-2 without changing total protein levels (Fig. 3B). α-Toc dose-dependently attenuated δ-T3-induced dephosphorylation of VEGFR-2/Akt signal proteins, but α-Toc alone did not have any effects. Further, α-Toc recovered downstream signals of the VEGFR2/Akt pathway, such as ERK1/2, which is involved in cell proliferation and survival [24]. These findings suggest that the inhibitory effect of α-Toc on the antiangiogenic effect of δ-T3 is at least in part mediated by regulation of VEGFR-2/Akt activity in endothelial cells. The present results regarding gene expression and protein phosphorylation suggested that α-Toc regulates an upstream target of signaling pathways or limits the cellular availability of δ-T3. To address the latter issue, we evaluated cellular vitamin E contents, finding that α-Toc decreased cellular δ-T3 uptake (Fig. 5). Similar results were observed in our previous study using human colorectal adenocarcinoma (DLD-1 cell line) cells [13]. Together with these results, we suggest a possibility that α-Toc abrogates the δ-T3-induced antiangiogenic signal by suppressing, in part, cellular δ-T3 uptake. The mechanism of α-Toc for reduction of cellular δ-T3 uptake (e.g., membrane stability alteration by α-Toc) is a subject of ongoing investigation. It is important to clarify the inhibitory effect of α-Toc on the antiangiogenic activity of T3 because α-Toc is the most abundant form of vitamin E in the human body and is common in natural food. T3 is also found in some food sources such as rice bran and palm, where the ratio of Toc to T3 is 50:50 and 25:75, respectively [25].
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Therefore, Toc, especially when it coexists with T3, may significantly alter the antiangiogenic effect of T3, which has potential to be chemopreventive ingredient if consumed regularly. α-Toc has also been shown to inhibit the anticancer and cholesterol-lowering effects of T3 [26]. It is difficult to use Toc-free T3 in an in vivo study, and a high-purity T3 will be necessary to clarify the real therapeutic value of T3. Our result suggests that Toc coexisting with T3 may interfere with the chemopreventive activity of T3 and that Toc-free T3 effectively suppresses angiogenic diseases to overcome the interference effect of α-Toc. Further research into the interaction mechanisms between T3 and Toc will be of great interest, and those clarification is essential to future chemotherapeutic applications. References [1] Tonini T, Rossi F, Claudio PP. Molecular basis of angiogenesis and cancer. Oncogene 2003;22:6549–56. [2] Shojaei F. Anti-angiogenesis therapy in cancer: current challenges and future perspectives. Cancer Lett 2012;320:130–7. [3] Wahl O, Oswald M, Tretzel L, Herres E, Arend J, Efferth T. Inhibition of tumor angiogenesis by antibodies, synthetic small molecules and natural products. Curr Med Chem 2011;18:3136–55. [4] Hodul PJ, Dong Y, Husain K, Pimiento JM, Chen J, Zhang A, et al. Vitamin E δ-tocotrienol induces p27(Kip1)-dependent cell-cycle arrest in pancreatic cancer cells via an E2F-1dependent mechanism. PLoS One 2013;8:e52526. http://dx.doi.org/10.1371/journal. pone.0052526. [5] Manu KA, Shanmugam MK, Ramachandran L, Li F, Fong CW, Kumar AP, et al. First evidence that γ-tocotrienol inhibits the growth of human gastric cancer and chemosensitizes it to capecitabine in a xenograft mouse model through the modulation of NF-κB pathway. Clin Cancer Res 2012;18:2220–9. [6] Shibata A, Nakagawa K, Sookwong P, Tsuzuki T, Oikawa S, Miyazawa T, et al. Tumor anti-angiogenic effect and mechanism of action of δ-tocotrienol. Biochem Pharmacol 2008;76:330–9. [7] Shibata A, Nakagawa K, Sookwong P, Tsuduki T, Tomita S, Shirakawa H, et al. Tocotrienol inhibits secretion of angiogenic factors from human colorectal adenocarcinoma cells by suppressing hypoxia-inducible factor-1α. J Nutr 2008;138:2136–42. [8] Guthrie N, Gapor A, Chambers AF, Carroll KK. Inhibition of proliferation of estrogen receptor-negative MDA-MB-435 and -positive MCF-7 human breast cancer cells by palm oil tocotrienols and tamoxifen, alone and in combination. J Nutr 1997;127:544S–8S. [9] Xu WL, Liu JR, Liu HK, Qi GY, Sun XR, Sun WG, et al. Inhibition of proliferation and induction of apoptosis by γ-tocotrienol in human colon carcinoma HT-29 cells. Nutrition 2009;25:555–66.
[10] Wada S, Satomi Y, Murakoshi M, Noguchi N, Yoshikawa T, Nishino H. Tumor suppressive effects of tocotrienol in vivo and in vitro. Cancer Lett 2005;229: 181–91. [11] Yap WN, Zaiden N, Tan YL, Ngoh CP, Zhang XW, Wong YC, et al. Id1, inhibitor of differentiation, is a key protein mediating anti-tumor responses of γ-tocotrienol in breast cancer cells. Cancer Lett 2010;291:187–99. [12] Shibata A, Nakagawa K, Sookwong P, Tsuduki T, Oikawa S, Miyazawa T. δ-Tocotrienol suppresses VEGF induced angiogenesis whereas α-tocopherol does not. J Agric Food Chem 2009;57:8696–704. [13] Shibata A, Nakagawa K, Sookwong P, Tsuduki T, Asai A, Miyazawa T. α-Tocopherol attenuates the cytotoxic effect of δ-tocotrienol in human colorectal adenocarcinoma cells. Biochem Biophys Res Commun 2010;397:214–9. [14] Nakagawa K, Shibata A, Yamashita S, Tsuzuki T, Kariya J, Oikawa S, et al. In vivo angiogenesis is suppressed by unsaturated vitamin E, tocotrienol. J Nutr 2007; 137:1938–43. [15] Ishiyama M, Tominaga H, Shiga M, Sasamoto K, Ohkura Y, Ueno K. A combined assay of cell viability and in vitro cytotoxicity with a highly water-soluble tetrazolium salt, neutral red and crystal violet. Biol Pharm Bull 1996;19:1518–20. [16] Blacher S, Devy L, Burbridge MF, Roland G, Tucker G, Noël A, et al. Improved quantification of angiogenesis in the rat aortic ring assay. Angiogenesis 2001;4:133–42. [17] Sookwong P, Nakagawa K, Murata K, Kojima Y, Miyazawa T. Quantitation of tocotrienol and tocopherol in various rice brans. J Agric Food Chem 2007;55:461–6. [18] Ferrara N, Gerber HP, LeCouter J. The biology of VEGF and its receptors. Nat Med 2003;9:669–76. [19] Constantinou C, Papas A, Constantinou AI. Vitamin E and cancer: an insight into the anticancer activities of vitamin E isomers and analogs. Int J Cancer 2008;123: 739–52. [20] Elisia I, Kitts DD. Different tocopherol isoforms vary in capacity to scavenge free radicals, prevent inflammatory response, and induce apoptosis in both adult- and fetal-derived intestinal epithelial cells. Biofactors 2013;39:663–71. [21] Schlaeppi JM, Wood JM. Targeting vascular endothelial growth factor (VEGF) for anticancer therapy, by anti-VEGF neutralizing monoclonal antibodies and VEGF receptor tyrosine-kinase inhibitors. Cancer Metastasis Rev 1999;18:473–81. [22] Li L, Ittmann MM, Ayala G, Tsai MJ, Amato RJ, Wheeler TM, et al. The emerging role of the PI3-K-Akt pathway in prostate cancer progression. Prostate Cancer Prostatic Dis 2005;8:108–18. [23] Royer MC, Lemaire-Ewing S, Desrumaux C, Monier S, Pais de Barros JP, Athias A, et al. 7-Ketocholesterol incorporation into sphingolipid/cholesterol-enriched (lipid raft) domains is impaired by vitamin E: a specific role for α-tocopherol with consequences on cell death. J Biol Chem 2009;284:15826–34. [24] Wang K, Zheng J. Signaling regulation of fetoplacental angiogenesis. J Endocrinol 2012;212:243–55. [25] Aggarwal BB, Sundaram C, Prasad S, Kannappan R. Tocotrienols, the vitamin E of the 21st century: its potential against cancer and other chronic diseases. Biochem Pharmacol 2010;80:1613–31. [26] Qureshi AA, Pearce BC, Nor RM, Gapor A, Peterson DM, Elson CE. Dietary α-tocopherol attenuates the impact of γ-tocotrienol on hepatic 3-hydroxy-3-methylglutaryl coenzyme A reductase activity in chickens. J Nutr 1996;126:389–94.
ScienceDirect Journal of Nutritional Biochemistry 26 (2015) 345 – 350
α-Tocopherol suppresses antiangiogenic effect of δ-tocotrienol in human umbilical vein endothelial cells☆ Akira Shibata a , Kiyotaka Nakagawa a,⁎, Tsuyoshi Tsuduki b , Teruo Miyazawa a a b
Food and Biodynamic Chemistry Laboratory, Graduate School of Agricultural Science, Tohoku University, Sendai 981-8555, Japan Laboratory of Food and Biomolecular Science, Graduate School of Agricultural Science, Tohoku University, Sendai 981-8555, Japan
Received 29 July 2014; received in revised form 8 November 2014; accepted 17 November 2014
Abstract Recently, tocotrienol (T3), a less well-known form of vitamin E, has gained attention as a potent hypocholesterolemic, anticancer and antiangiogenic agent. However, tocopherol (Toc), a commonly consumed form of vitamin E, has been reported to inhibit T3’s effects (hypocholesterolemic and anticancer activity). There has been no report on Toc’s effect on the antiangiogenic action of T3 during cotreatment. The aim of this study is to determine if and to what extent Toc affects the antiangiogenic effects of δ-T3 (the most potent isomer). This was achieved through cotreatment of human umbilical vein endothelial cells (HUVECs) with δ-T3 and Toc (α-, β-, γ- and δ-isomers). Toc, especially α-Toc, attenuated δ-T3-induced cytotoxicity and tube degradation in cotreated HUVECs, while α-Toc treatments did not exhibit any effects. A rat aortic ring assay also showed inhibition of δ-T3’s antiangiogenic effects by α-Toc. Further, in HUVEC study, cell cycle arrest and proapoptotic gene expression (p21, p27, caspase-3 and caspase-9) which were induced by δ-T3 were decreased by α-Toc treatment. α-Toc also suppressed δ-T3-induced dephosphorylation of vascular endothelial growth factor receptor 2 and Akt pathway proteins. Additionally, uptake of δ-T3 into HUVECs was decreased by α-Toc. Here we demonstrate that α-Toc not only has little antiangiogenic effect on endothelial cells but also reduces the antiangiogenic effects of δ-T3 through modulation of its cellular uptake and of relevant signal transduction pathways. Understanding T3’s antiangiogenic effects and interaction with Toc is important for developing medical applications. © 2015 Elsevier Inc. All rights reserved. Keywords: Angiogenesis; Tocotrienol; Tocopherol; VEGF; HUVEC
1. Introduction Angiogenesis takes place after activation of endothelial cells by angiogenic stimuli, and it can promote cancer growth by supplying nutrients and oxygen and removing waste products [1]. Therefore, antiangiogenesis is an effective defensive strategy against cancer progression [2]. Various anticancer agents that target angiogenesis have been developed, including natural and synthetic compounds [3]. Previous studies have shown that tocotrienol (T3) (especially the δ-isoform), an unsaturated vitamin E, has anticancer and antiangiogenic properties
Abbreviations: DLD-1, human colorectal adenocarcinoma; ERK, extracellular regulated kinase; HPLC, high-performance liquid chromatography; HUVEC, human umbilical vein endothelial cell; PTEN, phosphatase and tensin homolog deleted from chromosome 10; RT-PCR, reverse transcription polymerase chain reaction; T3, tocotrienol; Toc, tocopherol; VEGF, vascular endothelial growth factor; VEGFR-2, vascular endothelial growth factor receptor-2; WST-1, water-soluble tetrazolium salt ☆ Conflicts of interest: none. ⁎ Corresponding author at: Food and Biodynamic Chemistry Laboratory, Graduate School of Agricultural Science, Tohoku University, 1-1 TsutsumidoriAmamiyamachi, Aoba-ku, Sendai 981-8555, Japan. Tel.: +81 22 717 8906; fax: +81 22 717 8905. E-mail address: [email protected] (K. Nakagawa). http://dx.doi.org/10.1016/j.jnutbio.2014.11.010 0955-2863/© 2015 Elsevier Inc. All rights reserved.
in vitro and in animal models [4–7]. Many studies have demonstrated that T3 has potent proapoptotic and antiproliferative activities in cancer cells (e.g., breast, colon and liver cancer cells) [8–10]. Importantly, at the same T3 dose, almost no adverse effects on growth or function are seen in normal counterpart cells [11]. Previous studies by our group have shown that δ-T3 has the most potent antiangiogenic activities in vitro and in vivo. Conversely, tocopherol (Toc), a major form of vitamin E found in many foods, does not [12]. Therefore, δ-T3 would be a promising anticancer agent or an adjuvant for minimizing tumor angiogenesis. Recently, we showed that δ-T3-induced cytotoxicity was attenuated by α-Toc through inhibition of cellular uptake of δ-T3 into colon cancer cells [13]. Results from the study suggest that α-Toc inhibits a physiological function of δ-T3. Thus, there is a possibility that α-Toc may attenuate the antiangiogenic activity of δ-T3. However, the antiangiogenic effect of δ-T3 during cotreatment of these compounds is unknown. If α-Toc inhibits the effects of δ-T3, careful consideration is required to predict the antiangiogenic effects of δ-T3. In the present study, we cotreated human umbilical vein endothelial cells (HUVECs) with δ-T3 and Toc (α-, β-, γ- and δ-isomers) and investigated whether Toc inhibits the antiangiogenic effects of T3. Cytotoxicity, tube formation, apoptosis and cell-cycle-related gene expression, phosphorylation of signal proteins and cellular T3 accumulation were evaluated to explore the relationship between these compounds. To further examine whether Toc inhibits the antiangiogenic
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effect of T3 in a more physiological condition, a rat aortic ring assay was also performed.
Table 1 Primer sequences for quantitative real-time PCR amplification of selected human genes. Gene
GenBank (accession no.)
Sequence (5’–3’)
p21(CDKN1A)-F p21(CDKN1A)-R p27(CDKN1B)-F p27(CDKN1B)-R Caspase-3(CASP3)-F Caspase-3(CASP3)-R Caspase-9(CASP9)-F Caspase-9(CASP9)-R β-Actin(ACTB)-F β-Actin(ACTB)-R
NM_000389 NM_000389 NM_004064 NM_004064 NM_032991 NM_032991 NM_032996 NM_032996 NM_001101 NM_001101
AGCAGAGGAAGACCATGTGGAC TTTCGACCCTGAGAGTCTCCAG TGCAACCGACGATTCTTCTACTCAA CAAGCAGTGATGTATCTGATAAACAAGGA ATGGAAGCGAATCAATGGAC GGCTCAGAAGCACACAAACA TGCTCAGACCAGAGATTCGC CTTTCTGCTCGACATCACCAA TGGCACCCAGCACAATGAA CTAAGTCATAGTCCGCCTAGAAGCA
2. Materials and methods 2.1. Chemicals All Toc isomers (α-, β-, γ- and δ-Toc) were purchased from Eisai (Tokyo, Japan). δ-T3 was kindly gifted by Tama Biochemical (Tokyo, Japan). δ-T3 was selected for this study because it has been reported to show the most potent antiangiogenic activities among the four T3 isomers (α-, β-, γ- and δ-T3) [14]. Water-soluble tetrazolium salt-1 (WST-1) reagent was obtained from Dojindo Laboratories (Kumamoto, Japan). All other reagents used in this study were of analytical grade. 2.2. Cells and culture HUVECs were obtained from Kurabo (Osaka, Japan). The cells were cultured in a base medium (HuMedia-EB2) supplemented with 2% fetal bovine serum, 10 μg/L human epidermal growth factor, and 5 μg/L human basic fibroblast growth factor (Kurabo). Confluent HUVECs (passages 5 to 8) were used in all experiments. The cells were maintained at 37°C in a humidified atmosphere of 95% air and 5% CO2.
lengths were counted in four randomly selected microscopic fields on day 12 by using Lumina Vision software (Olympus, Tokyo, Japan). 2.8. Total RNA isolation and mRNA analysis
2.3. Preparation of test medium Each Toc isomer or δ-T3 was dissolved in ethanol at a concentration of 50 mM. The stock solution was then diluted with 1%–2% fetal bovine serum/HuMedia-EB2 with or without 10 μg/L vascular endothelial growth factor (VEGF) (R&D system, Minneapolis, MN, USA) to achieve the desired final concentration of vitamin E (0–50 μM). The final concentration of ethanol in the test medium was less than 0.1% (v/v), which did not affect cell viability. Medium with ethanol alone was similarly prepared and used as control medium.
After treating HUVECs with the test medium for 24 h, total RNA was isolated with the RNeasy Plus Mini kit (Qiagen, Valencia, CA, USA) for reverse transcription polymerase chain reaction (RT-PCR). cDNA was synthesized using a Ready-To-Go T-Primed FirstStrand kit (GE Healthcare, Piscataway, NJ, USA). Quantitative RT-PCR amplification was performed with the DNA Engine Opticon 2 system (MJ Research, Waltham, MA, USA) using SYBR Premix Ex Taq II (Takara Bio, Otsu, Japan) and gene specific primers for p21 (CDKN1A), p27 (CDKN1B), caspase-3 (CASP3), caspase-9 (CASP9) and β-actin (ACTB) (Table 1). PCR conditions for these primers were 95°C for 1 min, 95°C for 5 s and 60°C for 20 s over 40 cycles.
2.4. Cell viability assay HUVECs were plated at 3×103 cells/well in 96-well culture plates to determine cell viability. After 24 h, the medium was replaced with the test medium containing the compounds of interest (δ-T3 and Toc). Cells were incubated in test medium for 24 h, and cell viability was determined using the reagent WST-1 [15] according to the manufacturer’s instruction. 2.5. Tube formation assay Twenty-four-well plates were coated with 350 μl of Matrigel (Becton Dickinson, Bedford, MA, USA), and incubated at 37°C for 1 h for solidification. HUVECs were suspended in test medium containing the appropriate concentration of δ-T3 and α-Toc at 6×104 cells/well (500 μl/well). The cell suspension was plated onto the surface of the Matrigel and incubated for 18 h. Following incubation, the cells were fixed with 4% paraformaldehyde solution and photographed. Lengths of tube-structured cells were quantified using a Kurabo Angiogenesis Imaging Analyzer (imaging software; Kurabo).
2.9. Western blot analysis Confluent HUVECs were preincubated in a VEGF-free test medium for 6 h, then stimulated by the addition of 10 ng/ml VEGF and incubated for 5 min. Cellular proteins (60 μg/well) were separated by sodium dodecyl sulfate polyacrylamide gel electrophoresis (4%–20% e-PAGEL; Atto, Tokyo, Japan), and protein bands were transferred to polyvinylidene fluoride membranes (GE Healthcare). Following blocking, the membranes were incubated with primary antibodies for phospho-Akt (Ser473), phospho-Akt (Thr308), Akt, phospho-extracellular signal-regulated kinase (ERK), ERK, phospho-phosphatase and tensin homologue deleted on chromosome 10 (PTEN), PTEN, phospho-vascular endothelial growth factor receptor-2 (VEGFR-2) and β-actin, followed by a horseradishperoxidase-conjugated secondary antibody. All antibodies were purchased from Cell Signaling Technology (Beverly, MA, USA). ECL Plus reagents (GE Healthcare) were used for detection. 2.10. Apoptosis assay
2.6. Ethics statement The animal study was performed according to the protocols approved by the Institutional Committee for Use and Care of Laboratory Animals of Tohoku University, which was granted by Tohoku University Ethics Review Board (2008-Noudou-30), and the Guide for Care and Use of Laboratory Animals published by the US National Institutes of Health (NIH publication 85-23, revised 1996). All animals were anesthetized to minimize their suffering. 2.7. Rat aortic ring assay Ex vivo angiogenesis was studied by culturing rings of rat aorta in collagen gel [16]. Male Wistar rats (body weight ~200 g) were obtained from CLEA (Tokyo, Japan) and maintained in a clean, pathogen-free environment. The rats were housed individually in a temperature- (23°C) and humidity-controlled room with a 12-h light–dark cycle. Prior to the assay, each rat was acclimatized for 1 week. Once acclimatized, rats were sacrificed by bleeding from the right femoral artery under diethyl ether anesthesia. A section of thoracic aorta was removed and thoroughly washed with RPMI-1640 medium (Sigma, St. Louis, MO, USA) to remove any traces of blood. The aortic section was then turned inside out and cut into short segments of about 1.0–1.5 mm. A collagen gel matrix solution was made using 8 vol of porcine tendon collagen solution (3 mg/ml) (Nitta Gelatin, Osaka, Japan), 1 vol of 10× Eagle's MEM (GIBCO Laboratories, BRL, Rockville, MD, USA) and 1 vol of reconstitution buffer (0.08 M NaOH and 200 mM HEPES) and mixed gently at 4°C. Each aortic segment was placed in the center of a well on a 12-well culture plate and covered with 0.3 ml of prepared gel matrix solution. The solution was allowed to gel at 37°C for 20 min and was then overlaid with 1 ml of test medium (EGM2 medium; Clonetics, San Diego, CA, USA) containing 20 μg/L VEGF, δ-T3 and α-Toc. The plates were incubated for 12 days at 37°C in a fully humidified system of 5% CO2. The medium was changed on days 5 and 10 of the culture. The tube-structure
HUVECs were seeded 1×104 cells/well in gelatin-treated 96-well culture plates with culture medium and incubated until the cells reached confluence. After incubation, the medium was replaced with the test medium containing the compounds (δ-T3 and α-Toc). Cells were incubated in test medium for 6 h, and apoptosis ratio was evaluated. APOPercentage Dye kit (Biocolor Ltd., Belfast, UK) was used to quantify apoptosis, according to the manufacturer's instructions. This assay uses a dye that stains cells as they undergo the membrane ‘flip-flop’ event when phosphatidylserine is translocated to the outer membrane. This event is considered to be diagnostic of apoptosis but not necrosis. Digital images of APOPercentage-Dye-labeled cells, which appear bright pink against a white background, were used to quantify apoptotic cell ratio. Using Adobe Photoshop, the level of apoptosis was measured and expressed as a pixel number. 2.11. Cellular uptake of δ-T3 Cellular vitamin E content was determined by high-performance liquid chromatography (HPLC) with fluorescence detection. After confluent HUVECs were incubated with test medium for 6 h, cellular T3 and Toc were extracted as described previously [17]. HPLC was performed at 35°C using a silica column (ZORBAX Rx-SIL, 4.6×250 mm; Agilent, Palo Alto, CA, USA) with a mobile phase of hexane/1,4-dioxane/2-propanol (988:10:2) at a flow rate of 1.0 ml/min. T3 and Toc were detected with an RF-10AXL fluorescence detector (excitation 294 nm, emission 326 nm; Shimadzu, Kyoto, Japan). 2.12. Statistical analysis The data are expressed as the mean±S.D. Statistical analyses were performed using analysis of variance followed by Bonferroni/Dunn tests for multiple comparisons. Differences were considered significant at Pb.05.
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Fig. 1. Toc decreases δ-T3-induced reduction of HUVEC viability (A) and tube degradation (B). HUVECs were treated simultaneously with the indicated concentrations of δ-T3 and one of four Toc isomers, and incubated for 24 h. Cell viabilities were determined by WST-1 assay postincubation (A). Following incubation of the cells on a Matrigel-coated plate for 18 h, cultures were photographed, and the lengths of tube-structured cells were quantified using angiogenesis imaging software (B). Cell viability and tube length are expressed as the percentage to control. Values are mean±S.D. (n=6). Means without a common letter differ, Pb.05.
3. Results 3.1. Toc suppresses δ-T3-induced cytotoxicity and tube degradation δ-T3 showed an approximately 40% reduction of cell viability at 5 μM compared with vehicle in HUVECs following a 24 h incubation. When Toc isomers (α-, β-, γ- and δ-Toc) over a range of concentrations (5–50 μM) were cotreated with δ-T3, all exhibited a dose-dependent suppression of δ-T3-induced cytotoxicity in HUVECs. Of the isomers, α-, β- and γ-Toc completely suppressed the effect of δ-T3 at 50 μM (Fig. 1A). Because α-Toc showed the most potent inhibitory effect among the four Toc isomers, we used α-Toc for the following experiments. Effects of α-Toc on δ-T3-induced tube degradation were assessed by using a Matrigel assay. When HUVECs were cultured on Matrigel in the presence of test medium containing VEGF, elongated and robust tube-like structures formed when compared to control-treated cells (Fig. 1B). Treatment with δ-T3 potently inhibited the tube formation in the VEGF-treated group. Although α-Toc alone did not exhibit any effects, it dose-dependently attenuated δ-T3-induced tube degradation. To further examine whether α-Toc inhibits the antiangiogenic effect of δ-T3 in a more representative physiological environment, a rat aortic ring assay was conducted. This assay closely reflects several stages in angiogenesis, including endothelial cell proliferation, migration and tube formation. Here, VEGF significantly increased vessel sprouting
Fig. 2. α-Toc attenuates inhibition of tube formation by δ-T3 in rat aorta. Aortic segments were embedded in collagen gel layers and incubated with EBM-2 medium in the absence or presence of VEGF, δ-T3 and α-Toc. Medium was changed on day 5 and day 10. After 12 days, cultures were photographed (A) and vessel lengths were quantified (B). Values are mean±S.D. (n=4). Means without a common letter differ, Pb.05.
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compared with the control, and δ-T3 potently reduced microvessel growth in the VEGF-treated group (Fig. 2A and B). While α-Toc did not exhibit any effects even at 50 μM, it dose-dependently suppressed the antiangiogenic effect of δ-T3 in an ex vivo model. 3.2. α-Toc attenuates δ-T3-induced proapoptotic/cell cycle arrest gene expression and antiangiogenic signaling To elucidate the mechanism of α-Toc in suppressing the antiangiogenic effect of δ-T3, we measured the expression of genes involved in cell cycle and apoptosis. Quantitative RT-PCR showed that α-Toc decreases, in a dose-dependent manner, the expression of δ-T3-induced cell cycle arrest genes p21 (CDKN1A) and p27 (CDKN1B), and proapoptotic genes caspase-3 (CASP3) and caspase-9 (CASP9) (Fig. 3A). Gene expression changes induced by δ-T3 were completely suppressed by 50 μM of α-Toc.
α-Toc alone had no significant effect on gene expression when compared to control. Next, we investigated the VEGFR-2/Akt pathway, which includes PTEN and ERK, and which is critical for VEGF-induced angiogenic steps (e.g., cell proliferation, tube formation and antiapoptosis) [18]. Western blot analysis showed that VEGF-treated cells showed increased phosphorylation of Akt (Thr308 and Ser473), PTEN, ERK and VEGFR-2 when compared with control, whereas δ-T3 reduced VEGF-induced phosphorylation of these proteins (Fig. 3B). This effect was attenuated in a dose-dependent manner by α-Toc, causing an overall decrease of dephosphorylation of the proteins affected by δ-T3, and 50 μM of α-Toc almost completely inhibited the effects of δ-T3. α-Toc alone did not exhibit any effects on protein phosphorylation, and neither δ-T3 nor α-Toc changed total protein levels. Next, the cells were examined using the APOPercentage Dye kit to evaluate whether the apoptotic event was
Fig. 3. α-Toc rescues δ-T3-induced changes on mRNA expression (A) and protein (B) phosphorylation in HUVECs. HUVECs were treated for 24 h with the indicated concentrations of δ-T3 and α-Toc. mRNA levels of p21 (CDKN1A), p27 (CDKN1B), caspase-3 (CASP3) and caspase-9 (CASP9) were measured by real-time RT-PCR (A). Values are mean±S.D. (n=6). Means without a common letter differ, Pb.05. Protein phosphorylation levels of VEGFR-2/Akt pathway were analyzed by Western blotting (B). β-Actin acted as a loading control. Each blot is a representative of three replicate experiments.
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Fig. 4. α-Toc suppresses δ-T3-induced proapoptotic event in HUVECs. APOPercentage assay was used to assess the degree of apoptosis as described in Materials and Methods. The level of apoptosis was measured by a pixel number using Adobe Photoshop. Fold increase in apoptosis levels was expressed as the pixel numbers relative to control. Values are mean±S.D. (n=4–6). Means without a common letter differ, Pb.05.
caused by δ-T3. As shown in Fig. 4, HUVECs underwent flip-flop by treating δ-T3 for 6 h. δ-T3-induced proapoptotic event was suppressed by cotreatment with α-Toc in a dose-dependent manner, whereas α-Toc alone did not exhibit apoptotic effect. 3.3. α-Toc suppresses cellular uptake of δ-T3 on HUVECs To further elucidate the mechanism of the inhibitory effect of α-Toc on the antiangiogenic effects of δ-T3 in HUVECs, we measured cellular δT3 and α-Toc levels. Intracellular concentration of δ-T3 was 29.8± 1.5 nmol/mg protein, when HUVECs were incubated with 5 μM of δ-T3 for 6 h. This was significantly higher than in cotreated HUVECs, where δT3 (5 μM treatment) and α-Toc (50 μM treatment) had intracellular concentrations of 21.1±1.1 nmol/mg protein and 49.1±7.6 nmol/mg protein, respectively, following a 6 h incubation. Further, α-Toc decreased the uptake of δ-T3 into HUVECs in a dose-dependent manner (Fig. 5). This decrease in uptake is in concordance with the suppression of the antiangiogenic action of δ-T3 by adding α-Toc. When treated with 50 μM α-Toc alone, the intracellular concentration was 64.9± 3.3 nmol/mg protein. 4. Discussion The goal of this study was to clarify influence of Toc on T3-induced antiangiogenesis in vitro and in other relevant systems. This study demonstrated that α-Toc attenuates δ-T3's antiangiogenic response
Fig. 5. α-Toc prevents cellular uptake of δ-T3 in HUVECs. HUVECs were treated simultaneously for 6 h with the indicated concentrations of δ-T3 and α-Toc. Cellular contents of δ-T3 and α-Toc were measured by HPLC with fluorescence detection. Values are means±S.D. (n=6).
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(proliferation and tube formation) via suppression of δ-T3-induced protein dephosphorylation and reduction of cellular δ-T3 uptake. As shown in Fig. 1A, all Toc isomers attenuated δ-T3-induced cytotoxicity and did not show any inhibitory effects (as seen in δ-T3) on HUVEC viability. In this study, the inhibitory potency of the isomers against δ-T3 effects varied in the order α-Nγ-Nβ-Nδ-Toc. It has previously been shown that γ-Toc and δ-Toc induce apoptosis in some cancer cells, but α-Toc does not show this effect [19]. Cellular uptake of γ-Toc and δ-Toc was greater compared with that of α-Toc in intestinal epithelial cell lines, which in part explains the superior biopotency for both γ-Toc and δ-Toc compared with α-Toc in these cells [20]. Further, structurally, the δ-isomer lacks the 5- and 7-methyl groups attached onto its chroman ring; so it easily passes through cell membranes more than the α-isomer. This information seems to reflect the aforementioned order (α-Nγ-Nβ-Nδ-Toc) of isomer potency observed in our study. Similar results were obtained with our previous studies using human colon cancer cells [13]. We also aimed to determine whether α-Toc affects δ-T3's antiangiogenic effect on the key steps of angiogenesis, namely, tube formation. Here, α-Toc dose-dependently attenuated δ-T3-induced tube degradation. Importantly, α-Toc did not have any inhibitory effects on the tube formation in vitro or on vessel sprouting ex vivo (Figs. 1B, 2A and B). Concordant to these phenotypic results (Figs. 1–2), α-Toc dosedependently attenuated δ-T3-induced cell cycle arrest and proapoptotic signals at the mRNA level (Fig. 3A). Furthermore, we performed APOPercentage apoptosis assay and confirmed that α-Toc suppressed δ-T3-induced proapoptotic event in HUVECs (Fig. 4). These observations suggest that α-Toc suppresses the upstream events of δ-T3-induced cell cycle arrest or apoptosis. These genes are regulated by intracellular signal transduction from growth factor receptors such as the VEGFR-2/Akt pathway, which is involved in tumor angiogenesis and is correlated with tumor progression [21]. Activated Akt promotes cell survival and protects against apoptosis through multiple mechanisms. This includes regulating p21 and p27, and protein degradation or suppression of activation for caspase-3 and caspase-9 [22]. Some apoptotic inducers such as 7-ketocholesterol have been shown to induce cell death through inactivation of the phosphatidylinositol 3-kinase/Akt signaling pathway, which is known to be highly specific to lipid raft domains, and this event was canceled by α-Toc [23]. Therefore, we speculated that α-Toc may suppress δ-T3's antiangiogenic effect via regulating VEGFR-2/Akt activity in HUVECs. δ-T3 inhibited VEGF-induced phosphorylation of Akt (Thr308 and Ser473), PTEN and VEGFR-2 without changing total protein levels (Fig. 3B). α-Toc dose-dependently attenuated δ-T3-induced dephosphorylation of VEGFR-2/Akt signal proteins, but α-Toc alone did not have any effects. Further, α-Toc recovered downstream signals of the VEGFR2/Akt pathway, such as ERK1/2, which is involved in cell proliferation and survival [24]. These findings suggest that the inhibitory effect of α-Toc on the antiangiogenic effect of δ-T3 is at least in part mediated by regulation of VEGFR-2/Akt activity in endothelial cells. The present results regarding gene expression and protein phosphorylation suggested that α-Toc regulates an upstream target of signaling pathways or limits the cellular availability of δ-T3. To address the latter issue, we evaluated cellular vitamin E contents, finding that α-Toc decreased cellular δ-T3 uptake (Fig. 5). Similar results were observed in our previous study using human colorectal adenocarcinoma (DLD-1 cell line) cells [13]. Together with these results, we suggest a possibility that α-Toc abrogates the δ-T3-induced antiangiogenic signal by suppressing, in part, cellular δ-T3 uptake. The mechanism of α-Toc for reduction of cellular δ-T3 uptake (e.g., membrane stability alteration by α-Toc) is a subject of ongoing investigation. It is important to clarify the inhibitory effect of α-Toc on the antiangiogenic activity of T3 because α-Toc is the most abundant form of vitamin E in the human body and is common in natural food. T3 is also found in some food sources such as rice bran and palm, where the ratio of Toc to T3 is 50:50 and 25:75, respectively [25].
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