Dioscin Restores the Activity of the Anticancer Agent Adriamycin in

The purpose of this study was to investigate the ameliorating effect of dioscin (1) on multidrug resistance (MDR) in adriamycin (ADR)-resistant erythr...
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Dioscin Restores the Activity of the Anticancer Agent Adriamycin in Multidrug-Resistant Human Leukemia K562/Adriamycin Cells by Down-Regulating MDR1 via a Mechanism Involving NF-κB Signaling Inhibition Lijuan Wang,† Qiang Meng,†,‡ Changyuan Wang,†,‡ Qi Liu,†,‡ Jinyong Peng,†,‡ Xiaokui Huo,† Huijun Sun,†,‡ Xiaochi Ma,†,‡ and Kexin Liu*,†,‡,§ †

Department of Clinical Pharmacology, College of Pharmacy, Dalian Medical University, Dalian 116044, People’s Republic of China Provincial Key Laboratory for Pharmacokinetics and Transport, Liaoning, Dalian Medical University, Dalian 116044, People’s Republic of China § Institute of Cancer Stem Cell, Dalian Medical University, Dalian 116044, People’s Republic of China ‡

ABSTRACT: The purpose of this study was to investigate the ameliorating effect of dioscin (1) on multidrug resistance (MDR) in adriamycin (ADR)-resistant erythroleukemic cells (K562/adriamycin, K562/ADR) and to clarify the molecular mechanisms involved. High levels of multidrug resistance 1 (MDR1) mRNA and protein and reduced ADR retention were found in K562/ADR cells compared with parental cells (K562). Dioscin (1), a constituent of plants in the genus Discorea, significantly inhibited MDR1 mRNA and protein expression and MDR1 promoter and nuclear factor κ-B (NFκB) activity in K562/ADR cells. MDR1 mRNA and protein suppression resulted in the subsequent recovery of intracellular drug accumulation. Additionally, inhibitor κB-α (IκB-α) degradation was inhibited by 1. Dioscin (1) reversed ADR-induced MDR by down-regulating MDR1 expression by a mechanism that involves the inhibition of the NF-κB signaling pathway. These findings provide evidence to support the further investigation of the clinical application of dioscin (1) as a chemotherapy adjuvant.

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Dioscin (1) is a constituent of plants in the genus Discorea inclusive of medicinal plants such as Dioscorea nipponica Makino and Dioscorea zingiberensis C.H. Wright (Dioscoreaceae). Pharmacological research has demonstrated that dioscin (1) has anti-inflammatory, lipid-lowering, anticancer, and hepatoprotective effects.12−14 Furthermore, it has been widely used as an important raw material for the synthesis of steroid hormone drugs.15 Whether dioscin (1) can restore the activity of adriamycin (ADR) in cancer cells is presently unclear. In the current study a human leukemic cell line, adriamycin-resistant erythroleukemic cells (K562/adriamycin, K562/ADR) derived from parental cells (K562), was used to study the effect of 1 on ADR resistance. To understand the molecular mechanism involved, the effect of dioscin on MDR1 expression and the relationship between MDR1 expression and NF-κB activity were also investigated.

rug resistance is a formidable obstacle in cancer chemotherapy. Tumor cells can develop resistance to structurally diverse and mechanistically unrelated anticancer drugs, a phenomenon termed multidrug resistance (MDR).1 MDR is associated with the overexpression of the multidrug resistance 1 (MDR1) protein. MDR1, a 170 kD plasma membrane glycoprotein encoded by the MDR1 gene, is a multidrug transporter that functions as an adenosine triphosphate-dependent drug efflux pump.2 In tumor cells, MDR1 pumps out anticancer drugs, leading to drug resistance at the cellular level.3,4 The mechanism of MDR1-mediated drug resistance is related to nuclear factor κ-B (NF-κB), a protein complex acting as a transcription factor. Phosphorylation of inhibitor κB-α (IκB-α) is required for NF-κB activation. A significant increase in p-IκB-α leads to the activation of NF-κB.5 Previous studies have demonstrated that the up-regulation of MDR1 is due to the up-regulation of NF-κB.5−8 MDR1-mediated drug resistance can be effectively overcome by either blocking MDR1 drug pump function or inhibiting its expression.9 Inhibition of MDR1-mediated drug efflux leads to the resensitization of MDR cancer cells to chemotherapeutic agents, allowing for successful chemotherapy in patients with multidrug-resistant tumors.10 Currently, many clinical anticancer drugs such as certain alkaloids, anthracycline antibiotics, and epipodophyllotoxin derivatives can induce MDR.11 The discovery and development of safe and effective MDR reversal agents is urgently required. © 2013 American Chemical Society and American Society of Pharmacognosy

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RESULTS AND DISCUSSION MDR is often the result of the overexpression of drug efflux proteins such as MDR1.16 In MDR cancer cells, the intracellular concentration of drugs is reduced by drug efflux transport carried out by efflux transporters that decreases the therapeutic effect of anticancer drugs. One strategy for Received: January 24, 2013 Published: April 26, 2013 909

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reversing MDR in cells expressing ABC transporters is the combined use of anticancer drugs with efflux transporter modulators.17 Since the first MDR1 inhibitor, verapamil, was found in 1981, several agents have been reported to overcome MDR.6,8,18 Unfortunately, these drugs have unfavorable side effects or toxicity profiles at clinical doses and thus have limited therapeutic applications. In previous studies it was reported that certain flavonoids and natural phytochemicals can inhibit MDR1 expression.19−21 Dioscin (1), a steroidal saponin, is a traditional oriental herbal medicine component that exhibits potent antiviral, immunomodulatory, and cytotoxic activities.22−25 It is not known what effect dioscin (1) has on human leukemic cell multidrug resistance. The aim of this study was to investigate the potential reversal effect of dioscin (1) and to clarify the molecular mechanisms involved. It is reported that certain breast cancer cells are resistant to adriamycin.26 ADR is an antileukemia drug used widely in the clinic. K562/ADR cells overexpress the MDR1 protein and resist ADR-induced apoptosis.27 To characterize MDR in the K562/ADR cell line, various concentrations of ADR (from 0.1 to 100 μM) were used and cell viability was analyzed using the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) method. ADR exerted cytotoxicity against K562 and K562/ADR cells with IC50 values of 3.8 ± 0.8 μM and 42.4 ± 2.6 μM, respectively. The results indicated that K562/ADR cells are 11.0-fold more resistant to ADR when compared to parental K562 cells (Figure 1A). To confirm the overexpression of MDR1 in K562/ADR cells,26 MDR1 expression was measured in both K562/ADR and K562 cells using quantitative real-time PCR (qRT-PCR) and Western blot analysis. MDR1 mRNA and protein expression in K562/ADR cells was significantly higher than that for parental K562 cells (6.7-fold for MDR1 mRNA and 3.4-fold for MDR1 protein, respectively) (Figure 1B and C), confirming that MDR1 is overexpressed in K562/ADR cells. Recently, combination treatment approaches incorporating an MDR-reversal agent with ADR therapy have gained wide attention. This approach aims to enhance ADR antitumor activity in MDR phenotypes.28 Dioscin (1) is a potential adjuvant in such applications and the focus of the present study. The cytotoxicity of dioscin (1) at various concentrations (from 0.1 to 100 μM) in K562 and K562/ADR cells was compared. Both K562 and K562/ADR cells were nearly equally sensitive to dioscin (1). This compound exerted cytotoxicity in K562 and K562/ADR cells with IC50 values of 4.7 ± 1.3 μM and 3.3 ± 0.8 μM, respectively (p > 0.05) (Figure 2). These results indicate that 1 is effective against both drug-sensitive parental and MDR cancer cells. Nontoxic concentrations of dioscin (1) were determined in K562 and K562/ADR cells. The IC10 values were 0.4 ± 0.1 μM and 0.3 ± 0.1 μM, respectively. A

Figure 1. Characterization of K562 and K562/ADR cells. (A) Effects of ADR on the viability of K562 and K562/ADR cells. (B) Expression of MDR-1 mRNA by qRT-PCR analysis in K562 and K562/ADR cells. (C) MDR1 protein expression was detected in K562 and K562/ADR cells by Western blot. Means ± SD of three experiments are presented. Results marked with an asterisk are statistically significant (*p ≤ 0.05, **p ≤ 0.01).

Figure 2. Effects of dioscin (1) on the viability of K562 and K562/ ADR cells. Means ± SD of three experiments are presented.

concentration of 0.3 μM dioscin (1) was chosen for further use in this study.29 To determine whether dioscin (1) can affect the activity of ADR in K562/ADR and K562 cells, cell viabilities were analyzed, again using a MTT assay. Co-incubation with dioscin (1) (IC10) and various concentrations of ADR did not affect the cytotoxicity of ADR in K562 cells (Figure 3A). The IC50 values of ADR and dioscin (1) + ADR were 3.8 ± 0.8 μM and 3.4 ± 910

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dioscin + ADR treatments were 42.4 ± 2.6 μM and 2.4 ± 0.1 μM, respectively. The co-incubation of 1 and ADR decreased significantly the IC50 value of ADR in K562/ADR cells compared to ADR alone (p ≤ 0.01). These results indicate that dioscin (1) potentiates the cytotoxic effect of ADR in K562/ADR cells. Both MDR1 mRNA and protein levels increased as K562/ ADR cells developed resistance. A reduction of MDR1 expression at both the transcriptional and translational levels is possibly one of the mechanisms by which certain modulators or agents can reverse an MDR phenotype.30,31 Thus, the effects of dioscin (1) on MDR1 gene and protein expression were examined by qRT-PCR and Western blotting. After K562/ADR cells were treated with dioscin (1) (0.3 μM), the MDR1 mRNA concentration decreased by 93.1% (Figure 4A) and further declined by 97.8% after 24 h (Figure 4B). The observed change in MDR1 protein expression in K562/ADR cells was confirmed by Western blot analysis following incubation of the cells for 24 h with various concentrations of dioscin (1) or 0.3 μM 1 for the time indicated (0−48 h). Following treatment with 1 (0.3 μM), MDR1 protein content in K562/ADR cells decreased by 91.4% (Figure 4C) and further declined by 84.7% after 24 h (Figure 4D). On treating with 0.3 μM 1 for 24 h, MDR1 gene and protein expression decreased to nearly zero. The results indicated that dioscin (1) suppresses MDR1 in K562/ADR cells and can reverse ADR resistance. A beneficial property of ADR for the purposes of this study is its autofluorescence capacity.32 Fluorescence intensity is an indicator of ADR intracellular accumulation and may be shown

Figure 3. Effects of dioscin (1) on the cytotoxicity of ADR in K562 (A) and K562/ADR (B) cells. Means ± SD of three experiments are presented. Results marked with an asterisk are statistically significant (**p ≤ 0.01).

0.8 μM in K562 cells, respectively. However, dioscin markedly increased the cytotoxicity of ADR against K562/ADR cells (Figure 3B). In K562/ADR cells, the IC50 values for ADR and

Figure 4. Effects of dioscin (1) on MDR1 gene and protein expression in K562/ADR cells. (A, B) Cells were treated with different concentrations of 1 for 24 h and 0.3 μM 1 for 0−48 h. MDR1 gene expression was analyzed by qRT-PCR. (C, D) Cells were treated with various concentrations of 1 for 24 h or 0.3 μM 1 for 0−48 h, and its effects on MDR1 protein expression were analyzed by Western blotting. Means ± SD of three experiments are presented. Results marked with an asterisk are statistically significant (*p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001). 911

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were analyzed. The activity of NF-κB and the MDR1 promoter was markedly enhanced by TNF-α. The increase in activity caused by TNF-α was sharply inhibited by 1 (Figure 6A and B), suggesting that dioscin inhibits the activity of MDR1, at least in part, by down-regulating NF-κB activity. To further determine the effect of dioscin (1) on the NF-κB signaling pathway, K562/ADR cells were treated with various concentrations of 1 and TNF-α. Western blot analyses was performed to detect changes in MDR1 and IκB-α protein expression. MDR1 expression was increased by TNF-α. This effect was significantly inhibited by 1 (Figure 6C). Furthermore, TNF-α significantly increased IκB-α phosphorylation, which was subsequently decreased following treatment with 1 at 12−48 h (Figure 6D). These data suggest that dioscin suppresses NF-κB activation by inhibiting the phosphorylation of IκB-α, thus inhibiting the expression of MDR1. In summary, these data strongly imply that dioscin (1) (1) can increase the intracellular accumulation of ADR in K562/ ADR cells at nontoxic concentrations, (2) works by downregulating MDR1 by a mechanism involving the inhibition of the NF-κB signaling pathway, and (3) is potentially a new, potent, and clinically relevant MDR reversal agent. These findings provide evidence in support of the further investigation into the clinical application of dioscin (1) as an adjuvant in cancer chemotherapy.

by representative histograms of K562, K562/ADR, and K562/ ADR cells pretreated with 0.3 μM dioscin (1) for 24 h. The mean fluorescence intensity of ADR in K562/ADR cells was 46.3% lower than that of K562 cells. Dioscin (1) treatment enhanced the intracellular accumulation of ADR in K562/ADR cells. The fluorescence intensity of ADR in dioscin (1)-treated K562/ADR cells increased by 78.4% compared to untreated K562/ADR cells. In turn, the fluorescence intensity of ADR in dioscin-treated K562/ADR cells increased by 59.8% compared with K562 cells (Figure 5). These results demonstrated that intracellular ADR increases in K562/ADR cells following treatment with dioscin (1).

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EXPERIMENTAL SECTION

General Experimental Procedures. Dioscin (1) was kindly provided by Professor Jinyong Peng (College of Pharmacy, Dalian Medical University, Dalian, People’s Republic of China). The purity of 1 was 96.55% as determined by HPLC.39 Adriamycin was purchased from Shenzhen Main Luck Pharmaceuticals, Inc. (Shenzhen, People’s Republic of China). RPMI 1640 medium was from Gibco BRL (Grand Island, NY, USA). Fetal bovine serum (FBS) was purchased from Invitrogen Life Technologies Corporation (Invitrogen, Carlsband, CA, USA). TNF-α was from Peprotech (Rocky Hill, NJ, USA). Antibodies against phospho-IκB-α and horseradish peroxidase-conjugated antimouse IgG antibodies were purchased from Cell Signaling Technology (Beverly, MA, USA). Antibodies against β-actin and MDR1 were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA). MTT was purchased from USB Corporation (Cleveland, OH, USA). All other reagents and solvents were of analytical grade. Cell Culture. K562 and K562/ADR cells were purchased from Nanjing KeyGen Biotech Co., Ltd. (Nanjing, People’s Republic of China) and were grown in RPMI 1640 medium supplemented with 10% FBS, 100 units/mL penicillin, and 0.1 mg/mL streptomycin and kept in an incubator at 37 °C with 95% humidity and 5% CO2. Cultures initiated at a density of 105 cells/mL grew exponentially to about 106 cells/mL in 3 days. K562/ADR cells were cultured in RPMI 1640 medium in the presence of 1.8 μM ADR. Cells were grown in RPMI 1640 medium without ADR for 2 weeks prior to experiments. For the assays, and in order to have cells in the exponential growth phase, cultures were initiated at 5 × 105 cells/mL and used 24 h later, reaching a density of about (8−10) × 105 cells/mL. Cytotoxicity Assays. Cell viability was determined using an MTT assay. K562 and K562/ADR cells were disaggregated through a pipet and counted with a hemacytometer. Cells were seeded in 96-well plates (4 × 103 cells/well) and then treated with various concentrations of ADR with or without dioscin (1). After incubation for 48 h, 10 μL of MTT reagent (5 mg/mL in phosphate-buffered saline, PBS) was added to each well and left to incubate for an additional 4 h. A 100 mL aliquot of SDS−isobutanol−HCl solution (10% SDS, 5% isobutanol, and 12 μM HCl) was added and left to incubate overnight. Relative cell viability was obtained on a microplate reader (Bio-Rad, San Diego, CA, USA) with a 570 nm filter. The inhibition rate of various concentrations of 1 on the two cell lines was evaluated at IC10. IC10 values were calculated from three

Figure 5. Effects of dioscin (1) on ADR accumulation in K562 and K562/ADR cells: (A) untreated K562 cells; (B) untreated K562/ADR cells; (C) K562/ADR cells treated with 0.3 μM 1. Means ± SD of three experiments are presented. Results marked with an asterisk are statistically significant (*p ≤ 0.05, ***p ≤ 0.001).

MDR1 expression has been studied in vitro in K562/ADM,20 A549,21 MCF-7/ADR,19 HepG2/ADM,33 and Caco-234 cells and in vivo.35 Up-regulation of MDR1 gene expression has been correlated with the NF-κB signaling pathway,36 cyclooxygenase2,37 and protein kinase C.38 Among them, the NF-κB signaling pathway is reported most frequently as the molecular mechanism underpinning the up-regulation of MDR1 gene expression.5,7,8,36 In recent studies, a 5′-serial truncation analysis of the MDR1 promoter revealed a mutation of a consensus NF-κB binding site.7 Additionally, IκB degradation was shown to result in NFκB activation.8 Furthermore, the up-regulation of MDR1 gene expression is mediated by a NF-κB signaling pathway that requires a NF-κB binding site located distal to the MDR1 promoter.36 Since up-regulation of MDR1 has been found to be NF-κB-dependent, it is believed that NF-κB inhibitors could decrease MDR1 protein expression and restore chemosensitivity.5 As activation of NF-κB requires the phosphorylation of IκB-α, these experiments were designed to determine a possible role of NF-κB activation in dioscin (1)-mediated MDR reversal and to examine whether NF-κB activation is inhibited through inhibiting the phosphorylation of IκB-α. Tumor necrosis factor α (TNF-α), a NF-κB signaling pathway stimulator, was used to investigate NF-κB activity. TNF-α can increase NF-κB activity, the level of IκB-α phosphorylation, and MDR1 protein expression.5,8 To better understand the molecular mechanism of MDR reversal by dioscin (1), the effects of 1 on the MDR1 and NF-κB promoter 912

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Figure 6. Effects of dioscin (1) on the promoter activities of MDR1 and NF-κB, and IκB-α degradation in K562/ADR cells. Luciferase activity of MDR1 (A) and NF-κB (B) reporter genes was measured using a dual-luciferase reporter assay. (C) K562/ADR cells were incubated for 24 h with 0.3 μM 1 and 5 nM TNF-α. Cells were then lysed and subjected to Western blot analysis using MDR1 and β-actin antibodies. (D) K562/ADR cells were incubated with 0.3 μM 1 for 0−48 h followed by TNF-α for 30 min. The cells were lysed and subjected to Western blot analysis using phospho-IκB-α and β-actin antibodies. Means ± SD of three experiments are presented. Results marked with an asterisk are statistically significant (*p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001). independent experiments using SPSS. A dioscin concentration of 0.3 μM (IC10) was chosen for use in this study. Quantitative RT-PCR. Total RNA was extracted using TRIzol Reagent (Invitrogen, Shanghai, People’s Republic of China), according to the manufacturer’s protocol. RNA pellets were resuspended in diethyl pyrocarbonate-treated deionized water. RNA samples were analyzed by agarose gel electrophoresis, and integrity was examined by visualization of intact 18S and 28S rRNA under ultraviolet light. One microgram of total RNA was used to prepare cDNA by reverse transcription using a PrimeScript RT reagent kit (Takara, Dalian, People’s Republic of China). The primer sequences were as follows: MDR1 (forward): 5′- GGAGCCTACTTGGTGGCACATAA-3′, (reverse): 5′-TGGCATAGTCAGGAGCAAATGAAC-3′; β-actin (forward): 5′-ATTGAACACGGCATTGTCAC-3′, (reverse): 5′CATCGGAACCGCTCATTG-3′. cDNA was amplified using a SYBR Premix Ex Taq kit (Takara, Dalian, People’s Republic of China) and an Mx3000p instrument (Agilent). The PCR protocol was as follows: one cycle of denaturation at 95 °C for 30 s; 40 cycles of denaturation at 95 °C for 5 s; and annealing at 60 °C for 20 s. PCR products were analyzed using the ΔΔCT method40 with β-actin as the standard gene. Western Blotting. After incubation with dioscin (1) and TNF-α at 37 °C for 0−48 h, cells were collected and washed with PBS. Protein was extracted using the total protein extraction kit (KeyGen Biotech, Nanjing, People’s Republic of China), according to the manufacturer’s protocol. Protein concentrations were determined using the BCA protein assay kit (KeyGen Biotech, Nanjing, People’s Republic of China) following the manufacturer’s protocol. For Western blot analysis, protein was first denatured by mixing with an equal volume of 2× sample loading buffer (100 μL of loading buffer and 4 μL of mercaptoethanol) and then boiled at 100 °C for 5 min. An aliquot (60 μg of protein) of the supernatant was run on a 10% SDS-

polyacrylamide gel by electrophoresis. Proteins were transferred onto a PVDF membrane (Millipore, Billerica, MA, USA) and blocked with 5% nonfat milk in Tris-buffered saline with 0.1% Tween-20 (TTBS) for 2 h at 37 °C. β-Actin served as the loading control. Membranes were incubated overnight at 4 °C with 1:100 and 1:1000 dilutions of monoclonal antibodies for MDR1 and Phospho-IκB-α (Ser32/36), respectively, and with a 1:500 dilution of monoclonal antibody for β-actin. After incubation with a primary antibody, membranes were rinsed three times with TTBS and incubated with a 1:5000 dilution of anti-mouse horseradish peroxidase-conjugated secondary antibody for 2 h at 37 °C. After extensive washing with TBST, membranes were exposed to enhanced chemiluminescenceplus reagents (ECL) from Beyotime Institute of Biotechnology (Haimen, People’s Republic of China), according to the manufacturer’s protocol. Emitted light was recorded with a BioSpectrum-410 multispectral imaging system using a Chemi 410 HR camera. Protein bands were visualized and photographed under transmitted ultraviolet light. Images were used for semiquantitative measurements based on band densitometry. Plasmid Construction. Human MDR1 (hMDR1)-Luc reporter plasmid (Takara, Dalian, People’s Republic of China) and NF-κB-Luc reporter plasmid (Beyotime Institute of Biotechnology) were used. hMDR1-Luc, a 1269 bp region (residues −969 to +300) of the hMDR1 promoter, was amplified with PCR using a 5′-prime (5′TTTCTCTATCGATAGGTACCCTGGACCCAGCTTTCACTAT) and 3′-prime (5′-CCGGAATGCCAAGCTTTTCTCCCCCTATTTACACTAAT) primer. The gel-purified PCR product was digested using KpnI and HindIII and then cloned into pGL3-basic firefly luciferase reporter vectors (Promega, Madison, WI, USA) using an InFusion HD cloning kit (Takara, Dalian, People’s Republic of China). The cloning of the hMDR1 promoter fragment into the pGL3 promoter vector was confirmed using restriction enzyme digestion and 913

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direct sequencing, which showed a similarity of 100%. NF-κB-luc contained a firefly luciferase gene driven by four copies of the NF-κB response element. All the resulting plasmids were amplified in Escherichia coli. Transient Transfection and Luciferase Assay. To determine promoter activity, a dual-luciferase reporter assay system (Promega) was applied. Briefly, cells were plated in 24-well plates overnight and co-transfected transiently with hMDR1-Luc or NF-κB-Luc construct and pRL-SV plasmid (Renilla luciferase expression for normalization) (Promega), using the LipofectAMINETM 2000 reagent (Invitrogen, Carlsbad, CA, USA). Cells were then exposed to dioscin (1) and TNFα for 24 h. Luciferase activity in cell lysates was measured using a TD20 luminometer (Turner Designs, Sunnyvale, CA, USA). Relative luciferase activity was calculated by normalizing MDR1 or NF-κB promoter-driven firefly luciferase activity to Renilla luciferase activity (Luminoskan Ascent, Thermo Electron). Accumulation of Adriamycin. K562 and K562/ADR cells were plated in six-well plates at a concentration of 4 × 106 cells in 4 mL of growth medium. After incubation with dioscin (1) at 37 °C for 24 h, 0.3 μM 1 was added to designated K562/ADR cells for another 24 h. K562 and K562/ADR cells were incubated with 25 μM ADR (with or without 1) for 1 h at 37 °C. After three washes with ice-cold PBS, the cells were subjected to flow cytometry (Becton Dickinson, San Diego, CA, USA) with excitation measured at 488 nm and emission measured at 575 nm. Statistical Analysis. All experiments were done in triplicate. For Western blot quantitation analysis, the sum of the density of protein bands was calculated and the amount of β-actin was used for normalization. The SPSS 19.0 statistical package was employed to perform correlation analysis. One-way analysis of variance (ANOVA) was used to determine the significance of differences between treatment groups. The Fisher’s least significant difference (LSD) test was used for multigroup comparisons. Statistical significance was considered to be p ≤ 0.05.

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AUTHOR INFORMATION

Corresponding Author

*Tel/fax: +86 411 86110407. E-mail: [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported by a grant from the National Natural Science Foundation of the People’s Republic of China (Nos. 81273580, 81072694) and the Dalian Government (No. 2010E12SF060).

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REFERENCES

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