Dihydroartemisin inhibits glioma invasiveness via a ROS to P53 to β-catenin signaling

Accepted Manuscript Title: Dihydroartemisin inhibits glioma invasiveness via a ROS to P53 to ␤-catenin signalling Authors: Zhongyou Que, Ping Wang, Yi Hu, Yixue Xue, Xiaobai Liu, Chengbin Qu, Jun Ma, Yunhui Liu PII: DOI: Reference:

S1043-6618(16)30509-6 http://dx.doi.org/doi:10.1016/j.phrs.2017.01.014 YPHRS 3475

To appear in:

Pharmacological Research

Received date: Revised date: Accepted date:

27-5-2016 10-1-2017 12-1-2017

Please cite this article as: Que Zhongyou, Wang Ping, Hu Yi, Xue Yixue, Liu Xiaobai, Qu Chengbin, Ma Jun, Liu Yunhui.Dihydroartemisin inhibits glioma invasiveness via a ROS to P53 to ␤-catenin signalling.Pharmacological Research http://dx.doi.org/10.1016/j.phrs.2017.01.014 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Title Dihydroartemisin inhibits glioma invasiveness via a ROS to P53 to β-catenin signalling

Zhongyou Que1,2, Ping Wang3, Yi Hu1,2, Yixue Xue3, Xiaobai Liu1,2, Chengbin Qu1,2, Jun Ma3, Yunhui Liu1,2*

1

Department of Neurosurgery, Shengjing Hospital of China Medical University,

Shenyang 110004, People’s Republic of China 2

Liaoning Research Center for Translational Medicine in Nervous System Disease,

Shenyang 110004, People’s Republic of China 3Department of Neurobiology, College of Basic Medicine, China Medical University, Shenyang 110122, People’s Republic of China

Correspondence to: Yunhui Liu, Department of Neurosurgery, Shengjing Hospital of China Medical University, Shenyang 110004, People’s Republic of China Telephone number: +86 024 96615-36111 Fax number: +86 024 2295 8989 Email: [email protected]

1

Graphical abstract

Abstract Dihydroartemisinin(DHA) is the active metabolic derivative of artemisinin. DHA has potential therapeutic effects on glioma but the detailed mechanism is unclear. In this study, we investigated the role and the underlying mechanisms of DHA in its inhibition of glioma cells. U87 cells are wild-type p53 glioblastoma cells and U251 cells containmutant p53.DHA inhibited the proliferation, migration and invasion of glioma cells in a dose-dependent manner. DHA promoted reactive oxygen species production and activated p53 in two glioma cell lines, U87 and U251. In U87 cells, DHA significantly up-regulated the expression of p–β-catenin(S45) and inhibited EGFR, β-catenin, p–β-catenin(Y333) and matrix metalloprotease7/9 activity. In U251 cells, DHA significantly up-regulated p–β-catenin(S45), p–β-catenin(Y333) and EGFR, but the expression of β-cateninwas unchanged. We also found that DHA and sh–β-catenin prevented the proliferation of U87 and U251 cells in vivo. In 2

conclusion, DHA inhibited the migration and invasion of human glioma cells with different types of p53 via different pathways.

Abbreviations: ChIP, chromatin immunoprecipitation; CKIα, casein kinase Iα; CNS, central nervous system; DHA, dihydroartemisinin; ECM, extracellular matrix; EGFR, epidermal growth factor receptor; ERK1/2, extracellular signal-regulated kinase1/2; GBM, glioblastoma multiforme; GSK3β, glycogen synthase kinase 3β; MMPs, matrix metalloproteinases; PKM2, pyruvate kinase M2; ROS, reactive oxygen species

Keywords dihydroartemisinin, glioma, migration, invasion, P53, β-catenin

Chemical compounds studied in this article: dihydroartemisinin (PubChem CID:456410)

3

1. Introduction Gliomas are the most common malignant primary central nervous system (CNS) tumors [1]. Glioblastoma multiforme (GBM), accounting for more than 50% of gliomas, is the most malignant type of glioma[2]. Enormous achievements have been made in the treatment of GBM usingmicrosurgery, radiotherapy and chemotherapy with temozolomide in the last few decades [3,4]. However, the prognosis for GBM patients still remains very poor, and the 5-year survival rate is less than 5%, even after aggressive surgery with adjuvant radio-chemotherapy [5,6]. Rapid cellular migration and aggressive invasiveness are the major biological malignant properties of GBM that contribute most to the poor prognosis of this disease. The extracellular matrix (ECM) is the major component of a cancer’s microenvironment and its remodeling, triggered by matrix metalloproteinases (MMPs), is an essential step in the development and metastasis of malignant tumors [7]. Within the family of MMPs, MMP7 and MMP9 play important roles in the migration and invasion of malignant gliomas [8]. Thus, the attenuation of tumor cell migration and invasion will be the key element in the successful treatment of GBM. Artemisinin, a highly effective antimalarial agent, was first extracted from the tropical plant,Artemisia annua, by Chinese scientists in the 1970s [9]. In recent years, artemisinin and its derivatives have drawn the attention of researchers to their anti-cancer therapeutic effects, such as inhibiting cell proliferation, inducing apoptosis, and

regulating

immune

inflammation

and

DNA damage

repair

[10-12].

Dihydroartemisinin (DHA) is the active metabolic derivative of artemisinin and acts as an intermediate in artemisinin-derived pharmacological effects in vivo[13]. Recent studies have revealed that DHA exhibits various anti-glioma efficacies by enhancing 4

the radiosensitivity and cytotoxic effect of temozolomide in vitro, inhibiting MMP17, Akt or caspase signaling pathways, and inducing autophagy in glioma cells [14-17]. However, studies regarding the role and mechanism of action of DHA in the migration and invasion of malignant gliomas are very limited. Reactive oxygen species (ROS) refer to a group of reactive oxidant molecules and free radicals derived from molecular oxygen. ROS can cause double-stranded DNA breaks and induce cell cycle redistribution, as well as apoptosis [18]. By activating extracellular signal-regulated kinase1/2 (ERK1/2), ROS are also responsible for the autophagy of glioma cells [19]. The antitumor activities of artemisinin and its derivatives rely on two different pathways: a ROS-dependent traditional cytotoxic effect,

and

ROS-independent

pathways

mediated

by

cell

signal

transduction–associated receptors [20]. However, the detailed molecular mechanisms involved in the inhibitory effects of DHA on the migration and invasion of glioma cells are still unknown. Herein, we hypothesize that DHA inhibits glioma cells by regulating ROS. Tumor protein p53 plays the role of a tumor suppressor under various environmental pressures. Interestingly,mutant p53 cannot be simply regarded as the loss of tumor suppressor function.Some mutant p53 can impair tumor suppressor function through a “dominant-negative” mechanism [21];another part of mutant p53 can even be regarded as a new tumor promoter in what is termed as “gain–of–function”[22]. The Wnt/β-catenin signaling pathway is involved in various biological activities such as morphogenesis, differentiation and proliferation. Aberrant activation of the Wnt/β-catenin signaling pathway plays a vital role in the proliferation, apoptosis and invasion of malignant gliomas [23]. Accumulated β-catenin is translocated into the 5

nucleus, affecting cell proliferation, differentiation, apoptosis and gene expression [24]. Wnt-dependent or -independent pathways are involved in the nuclear translocation of β-catenin. In the classic Wnt-dependent pathway, Wnt down-regulates the expression of phosphorylated β-catenin (p–β-catenin) (S45) by binding to transmembrane Frizzled receptors, resulting in suppressed ubiquitin-dependent β-catenin degradation [25]. In the Wnt-independent pathway, epidermal growth factor receptor (EGFR) activation induces the translocation of pyruvate kinase M2 (PKM2) and c-Src into the nucleus, where PKM2 binds to and transactivates c-Src–dependent p–β-catenin (Y333), promoting the binding of both proteins to the CCND1 promoter region to enhance the expression of downstream oncogenes [26]. Therefore, phosphorylated modification is a key mechanism in the β-catenin signaling pathway. In the present study, we aimed to investigate the effect of the inhibition of DHA on the migration and invasion of U87 and U251 cells, and to reveal underlying mechanisms by investigating the effects of DHA on ROS, P53, β-catenin and MMP9/7.

2. Materials and Methods 2.1. Cell culture Human U87 MG and U251 glioblastoma cell lines were purchased from Nanjing KGI Biotechnology Co., Ltd. U87 and U251 cells were cultured in Dulbecco's Modified Eagle Medium (DMEM) (Hyclone, USA) supplemented with 10% fetal bovine serum (FBS) (Hyclone, USA). Both glioblastoma cell lines were cultured in an incubator at 37°C in humidified atmosphere of 5% CO2. 2.2. Cell proliferation assay Cell proliferation was measured with CCK8 assay kit (Sigma, USA) according to 6

the according to the literature [27]. U87 and U251 cells were seeded into 96-well plates (Corning) at a density of 5×103 cells per well in standard DMEM and incubated for 24 h under standard conditions (37°C and 5 % CO2). The medium was replaced with either blank serum free DMEM or DMEM containing DHA (KPC Pharmaceuticals, China) at concentrations of 50, 100, 200, 400, 600 μmol/L. Total volume in each well was 200 μL. U87 or U251 cells were incubated in these solutions for 24 h followed by treatment with 20 μL of CCK8 in each well for another 1.5 h at 37°C. Finally, plates were shaken softly and the optical density was recorded at 570 nm (OD570) using ELISA plate reader (SYNERGY4, USA). At least three independent experiments were performed. The inhibition rate was calculated as following

formula:

(ControlOD570–experimental

group

OD570)/

ControlOD570×100%. Since the IC50 values of DHA were 210.77±18.2 μmol/L for U87 cells and 200.35±18.2 μmol/L for U251 cells in the present study, concentrations of 100, 200 and 400 μmol/L were used in the following studies. 2.3. In vitro migration assay The migratory capacity of U87 and U251 cells was evaluated by Transwell assay [28]. Cells were cultured in Transwell inserts of 8-μm pore (Corning Costar) placed in 24-well plates. U87 or U251 cells were trypsinized and seeded in serum-free DMEM at the density of 4x105/mL and 200 μL of cell suspension was added into the upper chambers. 500 μl of conditioned medium (DMEM medium supplemented with 10% FBS) was placed in the lower chambers and served as the revulsant of cell migration. Serum free DMEM served as a negative control. DHA was added in the upper chambers at the concentration of 100, 200 and 400 μmol/L. After incubated for 24 h, the inserts were taken out and cells remained on the upper surface of the filters were removed carefully with a cotton wool swab. Cells 7

that migrated to the underside surface were washed with PBS gently once and fixed with methanol and glacial acetic acid (mixed at 3:1) for 30 min at room temperature and stained in Giemsa stain for 15 min. The average number of migrating cells was counted in six random high-power fields (×400). 2.4. In vitro invasion assay Invasiveness of U87 and U251 cells was evaluated by Transwell invasion assay with inserts of 8-μm pore size (Corning Costar) as described previously [29,30]. The upper surface of Transwell inserts were coated with Matrigel (BD Biosciences, USA) diluted with medium at the ratio of 1:7. Parental U87 or U251 cells were prepared as described above. 500 μl of DMEM supplemented with 10% FBS was placed in the lower chambers. Serum free DMEM served as a negative control. 100, 200 and 400 μmol/L DHA was added in the upper chambers containing U87 cells or U251 cells. After incubated for 24 h, the inserts were taken out and prepared for observation under microscope just as mentioned above. The average number of invasive cells was counted in six random high-power fields (×400). 2.5. Detection of intracellular ROS Reactive oxygen species assay kit (Beyotime Biotechnology, China) was used to check the cellular concentrations of ROS according to the manufacturer’s instructions. Parental U87 or U251 cells were prepared as described above and when cells reached 80% confluent DMEM was replaced by DCFH-DA diluted with serum-free DMEM at the concentration of 10 μmol/L. After incubated for 20 min under standard conditions, cells were washed three times with serum-free DMEM to clear the extracellular DCFH-DA. Then U87 and U251cells were trypsinized and harvested. Flow cytometry were performed to test the fluorescence intensity in cells. 2.6. Plasmid construction and cell transfection 8

The overexpression and silencing plasmids of β-catenin, wild type p53, mutant type p53 and mutant plasmids of p-β-catenin(Y333) and p-β-catenin(S45) were constructed in pGPU6/GFP/Neo vector (GenePharma, Shanghai, China) and pIRES2-EGFP vector (GenScript Corporation, Nanjing, China). Plasmids carrying non-targeting sequence were used as negative control (NC). The sequence of shRNA-β-catenin

was

Sense:

5’-

CACCGGATGTGGATACCTCCCAAGTTTCAAGAGAACTTGGTTGGGAGGTAT CCACATCCTTTTTTG-3’.

Antisense:

5’-

GATCCAAAAAAGGATGTGGATACCTCCCAAGTTCTCTTGAAACTTGGGAGG TATCCACATCC-3’ .The sequence of sh negative control (shNC) was Sense: 5’CACCGTATGACAACAGCCTCAAGTTCAAGAGACTTGAGGCTGTTGTCATAC TTTTTTG-3’.

Antisense:

5’-

GATCCAAAAAAGTATGACAACAGCCTCAAGTCTCTTGAACTTGAGGCTGTT GTCATAC-3’. In order to enable β-catenin ,β-catenin-Mut and wild/Mutant type p53 to be over-expressed, gene of β-catenin,p-53 and related mutant type was transfected into pIRES2-EGFP expression vectors by standard procedures. An empty vector, pIRES2-EGFP, was used as a control. Parental U87 and U251 cells were seeded into 24-well plates (Corning, NY). According to the manufacturer’s instructions, stable transfection was performed with Lipofectamine 3000 and Plus Reagents (Life Technologies Corporation, Carlsbad,CA) when cells reached 50%–80% confluence. The stably transfected cells were selected by G418 (Sigma–Aldrich, St Louis, MO) and the transfection efficiency was assessed by Western blot. Cells were divided into the following groups as shown in Table 1. 2.7. Western Blot Western blot assay was carried out to detect the expression of MMP7/9, p53, 9

p-p53(S15), β-catenin, p-β-catenin(Y333) and p-β-catenin(S45). U87 or U251 cells were treated with DHA at the concentrations of 100, 200 and 400 μmol/L or DHA combined with EGFR inhibitor AG-1478 (Selleck, USA) for 24 h. Then cells were washed

three

times

with

ice-cold

PBS

and

lysed

with

ice-cold

radio

immunoprecipitation assay (RIPA) buffer (50mMTris-HCl, pH 8.0, 150mM NaCl, 0.5% sodium deoxycholate 1% NP-40, 0.1% SDS, and 1mM EDTA) supplemented with protease inhibitors (10 mg/ml aprotinin, 10 mg/ml phenyl-methylsulfonyl chloride, and 50mM sodium orthovanadate), and finally incubated on ice for 30 min. The pellet was disrupted with an ultrasonic crusher and the samples were centrifuged at 17,000 rpm for 45 min at 4°C. The supernatant was collected and transferred to a new tube. The sample tubes were heated in a boiling water bath immediately for 5 min aiming to denature the proteins. The concentration of the soluble material was determined with BCA protein assay kit (Beyotime Biotechnology, China), with bovine serum albumin used as a standard. Equal amounts of proteins (20–25 μg) were separated by 8% SDS-polyacrylamide gel electrophoresis (PAGE) and processed for immunoblotting with rabbit polyclonal antibodies against β-catenin (diluted at 1:1000; Proteintech, USA), p-β-catenin (Y333) (diluted at 1:1000; Abcam, USA), p-β-catenin (S45) (diluted at 1:1000;Cell Signal Technology, USA), MMP7/9 (diluted at 1:1000; Proteintech, USA). A mouse polyclonal anti-GAPDH antibody (diluted at 1:2000; proteintech, USA) was used as an internal control. All the protein bands were scanned with ChemiImager 5500 V2.03 software, and the integrated density values (IDV) were calculated by computerized image analysis system (Fluor Chen 2.0). 2.8. Gelatin zymography assay MMP9 and MMP7 activities were detected by gelatin zymography performed on 10% polyacrylamide gels containing 0.1% gelatin (Invitrogen, Carlsbad, CA, USA) 10

[31]. The bands were visualized by staining for 30 min with a solution containing 0.1% Coomassie R-250 in 40% ethanol and 10% acetic acid, followed by destaining in a solution containing 10% ethanol and 7.5 % acetic acid for 2 h at room temperature. Band densitometry was determined using ChemiImager 5500 V2.03 software. 2.9. Immunofluorescence Assays Immunofluorescence assay was used to detect the expression and distribution of β-catenin, p-β-catenin(Y333) and p-β-catenin(S45). Cells were cultured on glass coverslips, fixed with 4% paraformaldehyde for 20 minutes and permeated with 0.2% Triton X-100 for 10 minutes at room temperature followed by incubation in 5% BSA blocking buffer for 2 hours at room temperature. Subsequently, cells were then incubated in the blocking buffer with primary antibodies for β-catenin (diluted at 1:50; Proteintech, USA), p-β-catenin (Y333) (diluted at 1:50; Abcam, USA), p-β-catenin (S45) (diluted at 1:50; Cell Signal Technology, USA) diluted overnight at 4°C. After washed three times with PBS, cells were incubated with Alexa Fluor 555-labeled anti-rabbit IgG secondary antibody (1:500; Beyotime Institute of Biotechnology, China) for 1 h. The nuclei were counterstained with 0.5 μg/mL DAPI. The staining was analyzed using immunofluorescence microscopy (Olympus, Tokyo, Japan) and merged by the Chemi Imager 5500 V2.03 software. 2.10. Chromatin immunoprecipitation assay Simple chromatin immunoprecipitation (ChIP) Enzymatic Chromatin IP Kit (Cell Signaling Technology, Danvers, Massachusetts, USA) was used for ChIP assays according to the manufacturer's protocol. In brief, cells were crosslinked with 1% formaldehyde

and

collected

in

lysis

buffer.

Micrococcal

Nuclease.

Immunoprecipitation was used to digest Chromatin. 3 μg of anti-β-catenin antibody or 11

normal rabbit IgG was added in each immunoprecipitation sample followed by immunoprecipitation with Protein G Agarose Beads during an overnight incubation at 4 °C with gentle shaking. 2% were removed before antibody supplemental and stored at − 20°C as the input reference. The ChIP DNA was reverse-crosslinked with 5 mol/L NaCl and Proteinase K andpurified. Immunoprecipitated DNA was amplified by PCR using primers. In each PCR, the corresponding input was taken in parallel for PCR validation. 2.11. Luciferase reporter assay Then we selected a TCF4 binding site that is predicted to be located to the transcription start site on MMP9/7.We constructed luciferase reporter plasmids; Luciferase reporter Kp2000 contained a 2000 bp fragment which included TCF4 binding site in the MMP9/7 promoter region,wheareas luciferase reporter Kp700 and Kp100 contained a truncated fragment which did not include this TCF4 binding site. We then examined the effect of co-transfection of a β-catenin overexpressing vector(pIRES2-β-catenin) or the NC vector on the transcriptional activity of these MMP9/7 promoters by measurement of luciferase activity in 293T cells.

2.12. Subcutaneous xenografts in nude mice All animal experiments were approved by the the Ethics Committee of Shengjing Hospital. Athymic nude mice (BALB/C-nu/nu, 4 weeks old, male) were purchased from the Cancer Institute of the Chinese Academy of Medical Science.U87 and U251 cells stably transfected with pIRES2-β-catenin, sh-β-catenin were selected as described above.U87 and U251Cells were subcutaneously implanted into the right flanks of mice at density of 5×105 cells (n=10 each group). Animals were then treated 12

with different doses of DHA solvent (2, 10 and 50 mg/kg) by intragastric administration, once a day for 45 days. Animals were sacrificed on 45th day after injection. The tumor size was evaluated every five days until the 45th day when all the nude mice were sacrificed.The tumor volume was calculated by the formula: volume (mm3) =length×width2/2

2.13. Statistical analysis All results were described as mean ± S.D. Statistical analysis was performed with SPSS 20 software. Differences between two groups were assessed using a Student’s t-test and comparisons between multiple groups were performed using a one-way analysis of variance (ANOVA) test followed by Dunnett’s post-hoc test. P<0.05 was considered to be statistically significant.

3. Results 3.1. DHA inhibited the proliferation, migration, invasion and tumor growth of U87 and U251 cells in vitro and vivo U87 and U251 cells were treated with DHA at concentrations of 50, 100, 200, 400 and 600 μmol/L for 24 h. Results demonstrated that DHA inhibited the proliferation of both cell lines in a dose-dependent manner compared to the control group (P<0.05, Fig.1A). The IC50 of DHA were 210.77±18.2 μmol/L and 200.35±18.2 μmol/L in U87 and U251 cells respectively (P<0.05, Fig.1B). Similarly, 100, 200 and 400 μmol/L DHA inhibited the migration and invasion of U87 and U251 cells in a dose-dependent manner compared with the control group (P<0.01, Fig.1D-E). To confirm the cytotoxicity of DHA in the subsequent experiment,the 13

CCK-8 assay was used. On condition that the density of U87 and U251 cells was 5× 105 ml, 100, 200 and 400 μmol/L DHA could not change the viability of U87 and U251 cells significantly in 24h. (Fig.1C) To further evaluate the efficacy of DHA in vivo,the subcutaneous tumor animal model were established. As shown in Fig. 1F , the nude mice treated with 50 mg/kg DHA had the smallest size of tumor.DHA could inhibit the tumor growth of both cell lines in a dose-dependent manner. In this section,the results showed that DHA could inhibite the proliferation, migration, invasion both in vitro and vivo.And the effects is associated with the dose of DHA. 3.2. DHA inhibited the expression and activity of MMP9 and MMP7 in U87 cells and U251 cells Results of Western blot and zymography assays confirmed that 100, 200 and 400 μmol/L DHA significantly inhibited the protein expression and enzymatic activity of MMP9 and MMP7 compared with the control group in U87 cell lines (P<0.01, Fig.2). Similar results were observed in U251 cells that the expression and activity of MMP9 and MMP7 were also inhibited by DHA in a dose-dependent manner. The result suggested that the effects of DHA on migration and invasion in U87 and U251 cells was in connection of the expression and activity of MMP9 and MMP7. 3.3. DHA promoted ROS production and up-regulated P53 expression in U87 and U251 cells Result of flow cytometry showed that intracellular concentrations of ROS were significantly elevated in U87 cells in a dose-dependent manner compared with the control group by the treatment of 100, 200 and 400 μmol/L DHA (P<0.05, Fig.3A). 14

Results of Western blot confirmed that the expression of p-P53 (S15) and P53 was significantly up-regulated in U87 cells by DHA at the same concentrations (P<0.05, Fig.3C). Similar to U87 cells, DHA significantly promoted the production of ROS and up-regulated the protein expression of p-P53 (S15) and P53 in U251 cells in a dose-dependent manner. These results revealed the initial period of DHA’s pharmacological effect in U87 and U251 cells. 3.4. DHA affected the expression of EGFR, p-β-catenin(Y333/S45) and β-catenin in U87 and U251 cells Effects of DHA on the expression of EGFR, p-β-catenin(Y333/S45) and β-catenin showed discrepancies in U87 and U251 cells. As shown in Fig. 4, the expression levels of EGFR, p-β-catenin(Y333) and β-catenin were significantly down-regulated in U87 cells compared with the control group by the treatments of 100, 200 and 400 μmol/L DHA (P<0.05). However, p-β-catenin(S45) expression was significantly up-regulated in U87 cells (P<0.05, Fig4A). Combination of DHA with EGFR inhibitor further up-regulated the expression of p-β-catenin(S45) and down-regulated p-β-catenin(Y333) and β-catenin (P<0.01; P<0.05, Fig. 4E) compared with the DHA group. In U251 cells, EGFR, p-β-catenin(Y333) and p-β-catenin(S45) were up-regulated by DHA (P<0.05) whereas β-catenin expression did not change (Fig. 4I). Combined treatment of DHA with EGFR inhibitor significantly up-regulated p-β-catenin(S45) (P<0.05) and down-regulated β-catenin expression (P<0.05) compared with the control group, while there was no significant change in the expression of p-β-catenin(Y333) in U251 cells (Fig. 4M). As described above, the IC50 of DHA were 210.77±18.2 μmol/L in U87 cells and 200.35±18.2 μmol/L in U251 cells. Therefore, the concentration of 200 μmol/L was selected as the optimal dosage in the following experiments. As shown in Fig. 5, 15

the distribution of p-β-catenin(S45), p-β-catenin(Y333) and β-catenin was different in the two cell lines. In U87 cells, p-β-catenin(Y333) and β-catenin were highly expressed in the nuclei and p-β-catenin(S45) was mainly expressed in the cytoplasm. The treatment of DHA resulted in cell shrinkage and nuclei deviation. Compared with the control group, DHA significantly decreased the fluorescence intensity of p-β-catenin(Y333) and β-catenin and enhanced the fluorescence intensity of p-β-catenin(S45), indicating that the expression of p-β-catenin(Y333) and β-catenin was down-regulated and p-β-catenin(S45) was up-regulated by DHA in U87 cells. In U251 cells, β-catenin was highly expressed in the nuclei while p-β-catenin(Y333) and p-β-catenin(S45) were mainly expressed in the cytoplasm. 200 μmol/L DHA enhanced the fluorescence intensity of p-β-catenin(Y333) in the nuclei and p-β-catenin(S45) in the cytoplasm.

However, the fluorescence intensity of

p-β-catenin in the nuclei did not change. Aforementioned changes in the fluorescence intensity of p-β-catenin(Y333), p-β-catenin(S45) and β-catenin in U87 and U251 cells were consistent with the results of Western blot assay. These results in Fig 4 and Fig 5 revealed the underlying molecular mechanism of DHA . Several crucial molecular of β-catenin signal pathways were involved in this process.And two isolated molecular pathways, canonical and non-canonical β-catenin pathway played the different roles in U87 and U251 cells.It might be associated with the different molecular substeps of U87 and U251 cells. 3.5.The stable transfection efficiency of p53 Mut/WT, β-catenin and p-β-catenin(Y333/S45)Mut . To confirming the transfection efficiency of p53Mut/WT, β-catenin and p-β-catenin(Y333/S45)Mut in U87 and U251 cells.The expression of p53,β-catenin p-β-catenin(Y333/S45) after overexpressing or silencing in U87 and U251 cells had 16

been tested. The results showed that the expression of these genes had all changed significantly in both U87 and U251 cells.(Fig.10)

3.6. Effects of DHA on the expression of p-β-catenin(Y333/S45), β-catenin and the expression and activity of MMP9 and MMP7 by the transfection of wild or mutant P53 U87 cells were stably transfected with plasmids of mutant P53 (P53-Mut). Compared to the control group, 200 μmol/L DHA significantly down-regulated expressions of β-catenin, p-β-catenin(Y333), MMP9 and MMP7, lowered the ratio of p-β-catenin(Y333)/β-catenin (P<0.01) as well as inhibiting the activity of MMP9 and MMP7 (P<0.01), whereas it significantly up-regulated the expression of p-β-catenin(S45) and raised the ratio of p-β-catenin(S45)/β-catenin. There was no obvious difference between the DHA and DHA+NC groups. The expression of β-catenin,

p-β-catenin(Y333/S45)

and

MMP9/7,

the

ratio

of

p-β-catenin(Y333)/β-catenin and p-β-catenin(S45)/β-catenin as well as the activity of MMP9/7 were significantly promoted in the DHA+p53-Mut group compared with the DHA+NC group. U251 cells were stably transfected with plasmids of wild P53 (P53-WT). Compared to the control group, DHA showed no obvious effect on the expression of β-catenin.

However,

it

significantly

elevated

the

expression

levels

of

p-β-catenin(Y333/S45) and the ratio of p-β-catenin(Y333/S45)/β-catenin (P<0.01) and inhibited the expression and activity of MMP9/7 (Fig. 7). There was no significant difference in the above parameters between the groups of DHA and DHA+NC.

The expression of β-catenin,

p-β-catenin(Y333), the

ratio of

p-β-catenin(Y333)/β-catenin and the expression and activity of MMP9/7 were 17

significantly decreased in the DHA+p53-WT group compared with the DHA+NC group

(P<0.01).

However,

p-β-catenin(S45)

expression

and

p-β-catenin(S45)/β-catenin ratio were significantly increased (P<0.01). The results showed that wild type p53 could active p-β-catenin(S45) and suppressed p-β-catenin(Y333).But the mutant type p53 could

active both

p-β-catenin(S45) and p-β-catenin(Y333). These results suggested that p53 was a key regulator of β-catenin pathways and could influence the expression and activity of MMP9/7 in U87 and U251 cells. 3.7. Effects of DHA combined with β-catenin silence, over-expression, β-catenin(S45)-Mut or β-catenin(Y333)-Mut on the migration, invasion and MMP expression and activity in vitro and tumor growth in vivo As shown in Fig. 8A and B, DHA significantly inhibited the migration and invasion of U87 and U251 cells compared with the control group (P<0.05). There was no statistical difference between the groups of DHA and DHA+NC. Then β-catenin was silenced by transfecting with sh-β-catenin. Results showed that the migration and invasion of U87 and U251 cells were significantly inhibited in the DHA+sh-β-catenin group (P<0.05). As shown in Fig. 8C and D, cell migration and invasion were significantly

promoted

in

the

DHA+pIRES2-β-catenin

and

DHA+

pIRES2-β-catenin(S45)-Mut groups compared with the DHA+NC group (P<0.05), whereas they were inhibited in the DHA+ pIRES2-β-catenin(Y333)-Mut group (P<0.05). DHA significantly inhibited the expression and activity of MMP9 and MMP7 in U87 and U251 cells compared with the control group (P<0.05). There was no statistical difference between the groups of DHA and DHA+NC. The expression and activity of MMP9 and MMP7 in U87 and U251 cells were significantly inhibited 18

in the DHA+sh-β-catenin group compared with the DHA+NC group (P<0.05) (Fig.9A-D)

and

promoted

in

the

DHA+pIRES2-β-catenin

and

DHA+pIRES2-β-catenin(S45)-Mut groups compared with the DHA+NC group (P<0.05). DHA+ pIRES2-β-catenin(Y333)-Mut also significantly inhibited the expression and activity of MMP9 and MMP7 in U87 and U251 cells (P<0.05) (Fig.9E-H). We

also

explored

the

role

of

β-catenin

in

the

process

of

dihydroartemisinin-induced inhibition on human glioblastoma cells. As shown in Fig.8I,DHA

(10

mg/kg)

reduced

the

tumor

size.Over-expressed

β-catenin(pIRES2-β-catenin) could reduce the efficiacy of DHA,while sh-β-catenin enhanced it. This section revealed that β-catenin and p-β-catenin(S45/Y333) played important role in the expression and activity of MMP9 and MMP7 in U87 and U251 cells.And β-catenin could be regarded as a crucial regulator of MMP9 and MMP7.We would further investegate the detailed relationship between β-catenin and MMP9/7. 3.8. Expression of p53 after overexpressing or silencing β-catenin in U87 and U251 cells. To confirming the effects of p53 in β-catenin-induced changing the expression and activity of MMP9/7 in U87 and U251 cells.The expression of p53 after overexpressing or silencing β-catenin in U87 and U251 cells had been tested.The results showed that the expression of p53 had not changed significantly in both U87 and U251 cells after overexpressing or silencing β-catenin.(Fig.10)This result supported the opinion in our research that the changing of β-catenin was regulated by p53 in U87 and U251 cells which was treated with DHA. 3.9. β-catenin bound to the promoters of MMP9 and MMP7 19

Binding of β-catenin to the promoters of MMP9 and MMP7 was verified by chromatin immunoprecipitation assay. Transcription start sites (TSS) of MMP9 and MMP7 were predicted with the assistance of DBTSSHOME (http://dbtss.hgc.jp/). The binding sites between β-catenin and MMP9/7 were confirmed by the analysis of TSS upstream base sequence TTCAAAG and ACTTCAAAG that was within 1,000 bps. Binding sites between β-catenin and MMP9 or MMP7 were at 721 or 109 base sequence in the TSS upstream respectively. These binding sites were confirmed by chromatin immunoprecipitation assay.

Negative controls were designed with the

regions at 2,000 bps in the upstream of binding sites and regions where there was no potential binding site with β-catenin. Results demonstrated that β-catenin bound to the predicted binding sites of MMP9 and MMP7. No binding was conserved with the negative control.(Fig.11 AB) The luciferase reporter assay results showed that luciferase activity in the β-catenin/Kp700 or Kp100 transfected cells,whose reporter lacked the selected TCF4 binding site, was weaker than that of the β-catenin/ Kp2000 transfected cells, whose reporter included the TCF4 binding site (Fig.10 CD, *P<0.01). This result suggested that β-catenin regulated MMP7/9 transcription through activating the TCF4. 4. Discussion In the present study, we confirmed that DHA inhibited the proliferation, migration and invasion of human malignant glioma U87 and U251 cells in a dose-dependent manner. We also found that treatment with DHA combined with wild-type or mutant p53 showed different effects on the expression of β-catenin and p–β-catenin(Y333) in U87 and U251 cells. DHA activated the p53 protein by elevating the intracellular concentration of ROS. In U87 cells, DHA up-regulated the expression of p–β-catenin(S45) and down-regulated β-catenin via a Wnt-dependent 20

pathway. DHA also inhibited EGFR signaling to down-regulate p–β-catenin(Y333) by a Wnt-independent pathway. Down-regulation of β-catenin inhibited the expression and activity of MMP9 and MMP7, resulting in attenuated cell migration and invasion. In U251 cells, DHA also inhibited the expression and activity of MMP9 and MMP7. However, mutant p53 activated EGFR and up-regulated the expression of p–β-catenin(Y333) and

p–β-catenin(S45),

resulting in

unchanged

β-catenin

expression. The above results indicated that DHA may inhibit the migration and invasion of U251 cells via other signaling pathways. U87 and U251 cells were treated with 50, 100, 200, 400 and 600 μmol/L DHA for 24 h; DHA significantly inhibited cell proliferation compared with the untreated control group. Greater inhibitory effects could be obtained by treatment witha higher concentration of DHA, indicating that DHA inhibited the viability of glioma cells in a dose-dependent manner (Fig.1A). The IC50 of DHA was similar for both glioma cell lines, being 210.77±18.2 μmol/L for U87 cells and 200.35±18.2 μmol/L for U251 cells. Therefore, concentrations of 100, 200 and 400 μmol/L were used in the our studies. We have previously described how the IC50 of artemether, another derivative of artemisinin, was 958 μmol/L [32]. Obviously, DHA has even greater anti-glioma effects than artemether. Migration and invasiveness abilities contribute most to the infiltrative growth pattern and multiple metastases of gliomas. Thus, the inhibition of migration and invasiveness is an important goal in the treatment of GBM. In the present study, we confirmed that DHA inhibited the cell migration and invasion of U87 and U251 cells in a dose-dependent manner. MMPs are important proteolytic enzymes that degrade the ECM and play important roles in controlling the migration and invasion of glioma cells. MMPs belong to a super family and can be divided into four subgroups: 21

interstitial collagenases (MMP1/8), gelatinases (MMP2/9), stromelysins (MMP7), and membrane-type MMPs (MT-MMPs: MMP1/2/3/4) [33]. Our previous study revealed that MMP2, MMP7 and MMP9 played important roles in the regulation of glioma cell migration and invasion [32,34]. In this study, we discovered that DHA displayed significant inhibitory effects on the expression and activity of MMP9 and MMP7, indicating that DHA attenuated the migration and invasion of glioma cells by inhibiting MMPs. Artesunate can cause DNA double-strand breaks (DSB) via the generation of intracellular ROS [35,36],as well as increase the phosphorylation of p53 at S15 to promote p53-induced DNA damage repair [37]. In this study, we investigated changes of ROS and p53 under DHA treatment and found that DHA increased ROS and p53 in U87 and U251 cells, meaning that DHA may influence p53 expression via regulating ROS. It has been well documented that β-catenin and the downstream transcription factor, Tcf-4, are up-regulated in glioma tissue compared with normal brain tissue and that the β-catenin/Tcf-4 expression level correlates positively with pathological grades of glioma [38]. In order to verify whether DHA inhibited migration and invasion by down-regulating β-catenin expression, we investigated changes in the expression of β-catenin, p–β-catenin(Y333) and p–β-catenin(S45) under DHA treatment. It was found

that

p–β-catenin(S45)

was

up-regulated

whereas

β-catenin

and

p–β-catenin(Y333) were down-regulated. Yang et al. reported that the activation of EGFR mediated p–β-catenin(Y333) phosphorylation, resulting in increased and accumulated β-catenin [26]. Using bioinformatic methods,Chao et al. predicted that EGFR may be the direct target of DHA [20]. Therefore, we postulated that DHA may down-regulate p–β-catenin(Y333) expression via an EGFR signaling pathway. Indeed, 22

in this study, we found that DHA down-regulated EGFR expression. We also found that the combination of EGFR inhibitor and DHA significantly decreased the expression of β-catenin and p–β-catenin (Y333), indicating that DHA may inhibit β-catenin and p–β-catenin (Y333) expression by down-regulating EGFR. In U251 cells,

DHA

promoted

the

expression

of

EGFR,

p–β-catenin(S45)

and

p–β-catenin(Y333). However, β-catenin expression did not change. Combined treatment of EGFR inhibitor with DHA down-regulated β-catenin, rather than p–β-catenin(Y333). The different expression patterns of EGFR, p–β-catenin(Y333) and β-catenin between U87 andU251 cells may be due to the different p53 genotypes of the two cell lines: U87 cells are wild-type p53 glioblastoma cells and U251 cells are mutant p53cells [39,40]. Under

in

vivo

conditions,

degradation

of

β-catenin

relies

on

ubiquitination-dependent proteolysis by the formation of a multiprotein complex composed of adenomatous polyposis coli (APC), axin, and glycogen synthase kinase-3 (GSK-3) or CKIα[41]. In U87 cells, DHA may cause DNA damage by inducing ROS, and activate GSK-3 or CKIα function via a wild-type p53 pathway [42]. Meanwhile, DHA down-regulated the expression of EGFR and attenuated the phosphorylation of β-catenin(Y333) by inhibiting an EGFR signaling pathway, resulting in the decreased accumulation of β-catenin in the nucleus. In p53 mutant U251 cells, DNA damage can initiate the activation of mutant p53 transcription. Mutant p53 is known to bind to the miR-27a promoter region and suppress its expression, leading to promoted EFGR expression [43]. In this study, we showed that the phosphorylation of R273H p53 and EGFR expression were promoted by DHA in U251 cells accompanied by up-regulated p–β-catenin(Y333) expression. An EGFR inhibitor can reverse these changes. On the other hand, it is reasonable that mutant 23

p53 still retains the functions of wild-type p53 involved in the activation of phosphokinases that induce the ubiquitination-dependent degradation of β-catenin. This may explain why DHA induced p–β-catenin(S45) up-regulation but did not change β-catenin expression in U251 cells. Therefore, the inhibitory effects of DHA on cell migration and invasion may occur via other pathways, rather than via regulating β-catenin/MMP. DHA inhibited MMP9 expression through the inhibition of PKCα/Raf/ERK and JNK phosphorylation, and the reduction of NF-κB and AP-1 activation [44]. To verify whether the difference in expression of EGFR, p–β-catenin(Y333) and β-catenin was due to a different genotype of p53, U87 and U251 cells were transfected with p53-Mut or p53-WT plasmids. The proteins β-catenin and p–β-catenin(Y333) were up-regulated and down-regulated in p53-Mut–U87 and p53-WT–U251 cells, respectively. However, p–β-catenin(S45) was only regulated in p53-WT–U251 cells. The trend in the expression and activity of MMP9/7 was consistent with the change in β-catenin. These results indicated that the discrepancy in the effects of DHA on p–β-catenin(Y333) and β-catenin in two cell lines was associated with a p53 genotype. In an attempt to verify the roles of β-catenin, p–β-catenin(Y333) and p–β-catenin(S45) in the migration and invasion of glioblastoma cells, U87 and U251 cells were transfected with pIRES2–β-catenin, sh–β-catenin, p–β-catenin(Y333)-Mut and

p–β-catenin(S45)-Mut

plasmids.The

overexpression

of

β-catenin

and

p–β-catenin(S45)–Mut enhanced the migration and invasion of both cell lines by promoting the expression and activity of MMP9 and MMP7. On the contrary, sh–β-catenin and p–β-catenin(Y333)–Mut attenuated cell migration and invasion by inhibiting MMP9 and MMP7. Nuclear translocation of β-catenin promotes the 24

formation of a β-catenin/TCF4 complex, regulating MMP9/7 expression. However, the detailed mechanism is still unclear. TCF-4 is a key element in the Wnt/β-catenin signaling pathway. As the downstream activator, β-catenin is translocated from the cytoplasm into the nucleus and binds to TCF4 to generate the transcriptional active complex, which activates target genes [45]. Aberrant functions of β-catenin and TCF4 are associated with the development of a malignant tumor such as gastric cancer and glioma [38,46]. In the present study, base sequences of β-catenin/TCF4 complex were predicted to be within the promoter of MMP7/9 by bioinformatic software, including core sequences of TCF4 TTCAAAG and highly conservative TCF4 sequences ACTTCAAAG. Thus, we, for the first time, showed that there is a binding site between the β-catenin/TCF4 complex and MMP9/7, suggesting that β-catenin controls the migration and invasion of glioma cells by regulating MMP7 and MMP9 at the transcriptional level. This study demonstrated for the first time that DHA inhibited the migration and invasion of U87 or U251 cells via different pathways,in vivo and in vitro. DHA activated p53 by up-regulating ROS in both cell lines. In U87 cells, the activation of wild-type p53 up-regulated p–β-catenin(S45) and promoted the degradation of β-catenin via a Wnt-dependent pathway. Meanwhile, DHA down-regulated p–β-catenin(Y333)

by

inhibiting

EGFR

and

decreased

β-catenin

via

a

Wnt-independent pathway. Attenuated β-catenin expression inhibited the expression and activity of MMP7/9 at the transcriptional level, resulting in inhibited cell migration and invasion. However, the mechanism differed in U251 cells. The activation of mutant p53 up-regulated the expression of p–β-catenin(S45) and p–β-catenin(Y333), resulting in unchanged expression patterns of β-catenin. It is probable that the inhibitory effect of DHA on MMP7/9 may be via other pathways 25

rather than β-catenin. The results of this study have provided new ideas for the study of DHA-based therapy against glioma. 5.Conclusion DHA suppresses the proliferation, migration and invasion of human glioblastoma cells by inhibiting MMP9and MMP7.In p53 wild-type U87 cells, the mechanism of suppression by DHA is associated with β-catenin, which is regulated by Wnt-dependent and -independent ways. In p53 mutant-type U251 cells,DHA is not able to down-regulate the expression of β-catenin, while the expression and activity of MMP9 and MMP7 is also suppressed. Other molecular pathways may be involved in this process.

Conflict of interest statement We declare that all authors have no conflict of interest. Acknowledgments This work is supported by grants from the Natural Science Foundation of China (81172197, 81272564, 81372484 and 81573010), Liaoning Science and Technology Plan Project (No. 2015225007), Shenyang Science and Technology Plan Projects (Nos. F15-199-1-30 and F15-199-1-57) and outstanding scientific fund of Shengjing hospital (No. 201304).

References 1.

Q.T. Ostrom , H. Gittleman , J. Fulop , M. Liu , R. Blanda , C. Kromer , Y. Wolinsky , C. Kruchko , J.S. Barnholtz-Sloan , CBTRUS Statistical Report: 26

Primary Brain and Central Nervous System Tumors Diagnosed in the United States

in

2008-2012.

Neuro

Oncol.

17

Suppl

4(2015)

iv1–iv62.

doi:10.1093/neuonc/nov189

2.

D.N. Louis , H. Ohgaki , O.D. Wiestler , W.K. Cavenee , P.C. Burger , A. Jouvet , B.W. Scheithauer , P. Kleihues , The 2007 WHO classification of tumours of the central nervous system. Acta Neuropathol. 114(2007) 97–109. doi:10.1007/s00401-007-0243-4

3.

R. Stupp , W.P. Mason , M.J. van den Bent , M. Weller , B. Fisher , M.J. Taphoorn , K. Belanger , A.A. Brandes , C. Marosi , U. Bogdahn , J. Curschmann , R.C. Janzer , S.K. Ludwin , T. Gorlia , A. Allgeier , D. Lacombe , J.G. Cairncross , E. Eisenhauer , R.O. Mirimanoff , European Organisation for Research and Treatment of Cancer Brain Tumor and Radiotherapy Groups,National

Cancer

Institute

of

Canada

Clinical

Trials

Group,

Radiotherapy plus concomitant and adjuvant temozolomide for glioblastoma. N Engl J Med 352(2005) 987–996. doi:10.1056/NEJMoa043330

4.

V.W. Stieber, M.P. Mehta, Advances in radiation therapy for brain tumors. Neurol Clin 25(2007) 1005–33– ix. doi:10.1016/j.ncl.2007.07.005

5.

F.A. Siebzehnrubl, B.A. Reynolds, A. Vescovi, D.A. Steindler, L.P. Deleyrolle, The origins of glioma: E Pluribus Unum? Glia.59(2011) 1135–1147. doi:10.1002/glia.21143

27

6.

F.R. Lima , S.A. Kahn , R.C. Soletti ,D. Biasoli , T. Alves , A.C. da Fonseca , C. Garcia , L. Romão , J. Brito , R. Holanda-Afonso , J. Faria , H. Borges , V. Moura-Neto , Glioblastoma: therapeutic challenges, what lies ahead.

Biochim

Biophys

Acta.

1826(2012)

338–349.

doi:10.1016/j.bbcan.2012.05.004

7.

P. Lu,V.M. Weaver, Z. Werb, The extracellular matrix: a dynamic niche in cancer

progression.

J

Cell

Biol.

196(2012)

395–406.

doi:10.1083/jcb.201102147

8.

B. Psaila, D. Lyden, The metastatic niche: adapting the foreign soil. Nat Rev Cancer. 9(2009) 285–293. doi:10.1038/nrc2621

9.

M.A. van Agtmael, T.A. Eggelte, C.J. van Boxtel, Artemisinin drugs in the treatment of malaria: from medicinal herb to registered medication. Trends Pharmacol Sci. 20(1999) 199–205.

10.

I. Nakase, H. Lai,N.P. Singh,T. Sasaki,. Anticancer properties of artemisinin derivatives and their targeted delivery by transferrin conjugation. Int J Pharm. 354(2008) 28–33. doi:10.1016/j.ijpharm.2007.09.003

11.

H.H.Chen, H.J. Zhou, X. Fang, Inhibition of human cancer cell line growth and human umbilical vein endothelial cell angiogenesis by artemisinin derivatives in

vitro.

Pharmacol

Res.

48(2003)

231–236.

doi:10.1016/S1043-6618(03)00107-5

28

12.

C.Z. Zhang, H. Zhang, J. Yun, G.G.Chen, P.B.S. Lai, Dihydroartemisinin exhibits antitumor activity toward hepatocellular carcinoma in vitro and in vivo. Biochem Pharmacol. 83(2012) 1278–1289. doi:10.1016/j.bcp.2012.02.002

13.

Q. Wang, S. Wu, X. Zhao, C. Zhao, H. Zhao, L. Huo. Mechanisms of Dihydroartemisinin and Dihydroartemisinin/Holotransferrin Cytotoxicity in T-Cell

Lymphoma

Cells.

PLoS

ONE.

10(2015)

e0137331.

doi:10.1371/journal.pone.0137331

14.

J. Chen, X. Chen, F. Wang, H. Gao, W. Hu, Dihydroartemisinin suppresses glioma proliferation and invasion via inhibition of the ADAM17 pathway. Neurol Sci. 36( 2015) 435–440. doi:10.1007/s10072-014-1963-6

15.

Z.S. Zhang , J. Wang , Y.B. Shen , C.C. Guo , K.E. Sai , F.R. Chen , X. Mei , F.U. Han , Z.P. Chen , Dihydroartemisinin increases temozolomide efficacy in glioma cells by inducing autophagy. Oncol Lett 10(2015) 379–383. doi:10.3892/ol.2015.3183

16.

X.J. Huang, C.T. Li, W.P. Zhang, Y.B. Lu, S.H. Fang, E.Q. Wei, Dihydroartemisinin potentiates the cytotoxic effect of temozolomide in rat C6 glioma cells. Pharmacology.82(2008) 1–9. doi:10.1159/000125673

17.

S.J. Kim , M.S. Kim , J.W. Lee , C.H. Lee , H. Yoo , S.H. Shin , M.J. Park , S.H. Lee , Dihydroartemisinin enhances radiosensitivity of human

29

glioma cells in vitro. J Cancer Res Clin Oncol 132(2006) 129–135. doi:10.1007/s00432-005-0052-x

18.

R. Alan Mitteer , Y. Wang , J. Shah , S. Gordon , M. Fager , P.P. Butter , H. Jun Kim , C. Guardiola-Salmeron , A. Carabe-Fernandez , Y. Fan , Proton beam radiation induces DNA damage and cell apoptosis in glioma stem cells through

reactive

oxygen

species.

Sci

Rep

5(2015)

13961.

doi:10.1038/srep13961

19.

S. Pallichankandy, A. Rahman,F. Thayyullathil,S.Galadari, ROS-dependent activation of autophagy is a critical mechanism for the induction of anti-glioma effect

of

sanguinarine.

Free

Radic

Biol

Med.2015(89)

708–720.

doi:10.1016/j.freeradbiomed.2015.10.404

20.

C. Huang , Q. Ba , Q. Yue , J. Li , J. Li , R. Chu , H. Wang , Artemisinin rewires the protein interaction network in cancer cells: network analysis, pathway identification, and target prediction. Mol BioSyst. 9(2013) 3091. doi:10.1039/c3mb70342h

21

A. M. Goh, C. R.Coffill, D. P.Lane, The role of mutant p53 in human cancer. J. Pathol. 223(2011)

22

116–126.

D. Dittmer, S. Pati, G. Zambetti, S. Chu, A.K. Teresky, M. Moore, C. Finlay, A.J. Levine. Gain of function mutations in p53. Nat. Genet. 4 (1993) 42–46.

30

23.

M. Leushacke, N. Barker, Lgr5 and Lgr6 as markers to study adult stem cell roles

in

self-renewal

and

cancer.

Oncogene.

31(2011)

3009–3022.

doi:10.1038/onc.2011.479

24.

T. Shimizu , T. Kagawa , T. Inoue , A. Nonaka , S. Takada , H. Aburatani , T. Taga,

Stabilized beta -Catenin Functions through TCF/LEF Proteins and the

Notch/RBP-J Complex To Promote Proliferation and Suppress Differentiation of

Neural

Precursor

Cells.

Mol

Cell

Biol.

28(2008)

7427–7441.

doi:10.1128/MCB.01962-07

25.

C.S. Cselenyi, K.K. Jernigan, E. Tahinci, C.A. Thorne, L.A. Lee, E. Lee, LRP6 transduces a canonical Wnt signal independently of Axin degradation by inhibiting GSK3's phosphorylation of beta-catenin. Proc Natl Acad Sci USA. 105(2008) 8032–8037. doi:10.1073/pnas.0803025105

26.

W. Yang , Y. Xia , H. Ji , Y. Zheng , J. Liang ,W. Huang , X. Gao , K. Aldape , Z. Lu , Nuclear PKM2 regulates β-catenin transactivation upon EGFR activation. Nature 480(2011) 118–122. doi:10.1038/nature10598

27.

D. Ding , S. Wei , Y. Song , L. Li , G. Du , H. Zhan , Y. Cao , Osthole exhibits anti-cancer property in rat glioma cells through inhibiting PI3K/Akt and MAPK signaling pathways. Cell Physiol Biochem. 32(2013) 1751–1760. doi:10.1159/000356609

31

28.

P. Li, C. Zhou, L.Xu, H. Xiao. Hypoxia enhances stemness of cancer stem cells in glioblastoma: an in vitro study. Int J Med Sci. 10(2013) 399–407. doi:10.7150/ijms.5407

29.

S. Qi , Y. Song , Y. Peng , H. Wang , H. Long ,X. Yu , Z. Li , L. Fang , A. Wu , W. Luo,Y. Zhen , Y. Zhou , Y. Chen , C. Mai , Z. Liu , W. Fang

ZEB2

mediates multiple pathways regulating cell proliferation, migration, invasion, and

apoptosis

in

glioma.

PLoS

ONE.

7(2012)

e38842.

doi:10.1371/journal.pone.0038842

30.

J. Bai , P.J. Mei , H. Liu , C. Li , W. Li , Y.P. Wu , Z.Q. Yu , J.N.Zheng , BRG1 expression is increased in human glioma and controls glioma cell proliferation, migration and invasion in vitro. J Cancer Res Clin Oncol 138(2012) 991–998. doi:10.1007/s00432-012-1172-8

31.

H. Cai , Y. Xue , W. Liu ,Z. Li , Y. Hu , Z. Li , X. Shang , Y. Liu , Overexpression of Roundabout4 predicts poor prognosis of primary glioma patients via correlating with microvessel density. J Neurooncol. 123(2015) 161–169. doi:10.1007/s11060-015-1780-7

32.

Y.B. Wang , Y. Hu , Z. Li , P. Wang , Y.X. Xue , Y.L. Yao , B. Yu , Y.H. Liu , Artemether combined with shRNA interference of vascular cell adhesion molecule-1 significantly inhibited the malignant biological behavior of human glioma cells. PLoS ONE. 8(2013) e60834. doi:10.1371/journal.pone.0060834

32

33.

H. Sato , T. Takino , Y. Okada , J. Cao , A. Shinagawa , E. Yamamoto , M. Seiki , A matrix metalloproteinase expressed on the surface of invasive tumour cells. Nature 370(1994) 61–65. doi:10.1038/370061a0

34.

H. Xie, Y.X. Xue, L.B. Liu, P. Wang, Y.H. Liu, Ying HQ, Expressions of matrix metalloproteinase-7 and matrix metalloproteinase-14 associated with the activation of extracellular signal-regulated kinase1/2 in human brain gliomas of different pathological grades. Med Oncol. 28 Suppl 1(2011) S433–8. doi:10.1007/s12032-010-9660-7

35.

N. Berdelle, T. Nikolova, S. Quiros, T. Efferth, B. Kaina. Artesunate induces oxidative DNA damage, sustained DNA double-strand breaks, and the ATM/ATR damage response in cancer cells. Mol Cancer Ther. American Association

for

Cancer

Research.

10(2011)

2224–2233.

doi:10.1158/1535-7163.MCT-11-0534

36.

A.E. Mercer, I.M. Copple, J.L. Maggs, P.M.

O'Neill,B.K. Park,The role of

heme and the mitochondrion in the chemical and molecular mechanisms of mammalian cell death induced by the artemisinin antimalarials. J Biol Chem. 286(2011) 987–996. doi:10.1074/jbc.M110.144188

37.

M.S.Colman, C.A. Afshari, J.C. Barrett, Regulation of p53 stability and activity in response to genotoxic stress. Mutation Research/Reviews in Mutation Research. 462(2000) 179–188. doi:10.1016/S1383-5742(00)00035-1

33

38.

J. Zhang , K. Huang , Z. Shi , J. Zou , Y. Wang , Z. Jia , A. Zhang , L. Han , X. Yue , N. Liu , T. Jiang , Y. You , P. Pu ,C. Kang , High β-catenin/Tcf-4 activity confers glioma progression via direct regulation of AKT2 gene expression. Neur Oncol 13( 2011) 600–609. doi:10.1093/neuonc/nor034

39.

L. Chen , J. Zhang , L. Han , A. Zhang , C. Zhang , Y. Zheng , T. Jiang , P. Pu , C. Jiang , C. Kang . Downregulation of miR-221/222 sensitizes glioma cells to temozolomide by regulating apoptosis independently of p53 status. Oncol Rep. 27(2012)854–860. doi:10.3892/or.2011.1535

40.

Y. Kataoka, J.S. Murley, R. Patel, D.J. Grdina, Cytoprotection by WR-1065, the active form of amifostine, is independent of p53 status in human malignant glioma cell lines. Int J Radiat Biol. 76(2000) 633–639.

41.

C. Liu ,Y. Li , M. Semenov , C. Han , G.H. Baeg , Y. Tan , Z. Zhang , X. Lin , X. He , Control of beta-catenin phosphorylation/degradation by a dual-kinase mechanism. Cell 108(2002) 837–847.

42.

E. Levina, M. Oren, A. Ben-Ze'ev.

Downregulation of beta-catenin by p53

involves changes in the rate of beta-catenin phosphorylation and Axin dynamics. Oncogene 23(2004) 4444–4453. doi:10.1038/sj.onc.1207587

43.

W. Wang, B. Cheng, L. Miao, Y. Mei, M. Wu, Mutant p53-R273H gains new function in sustained activation of EGFR signaling via suppressing miR-27a expression. Cell Death Dis .4(2013) e574. doi:10.1038/cddis.2013.97

34

44.

Y.P. Hwang, H.J. Yun, H.G. Kim, E.H. Han, G.W. Lee, H.G. Jeong, Suppression of PMA-induced tumor cell invasion by dihydroartemisinin via inhibition of PKCα/Raf/MAPKs and NF-κB/AP-1-dependent mechanisms. Biochem Pharmacol. 79(2010) 1714–1726. doi:10.1016/j.bcp.2010.02.003

45.

H. Brantjes, N. Barker, J. van Es, H. Clevers, TCF: Lady Justice casting the final verdict on the outcome of Wnt signalling. Biol Chem. 383(2002) 255–261. doi:10.1515/BC.2002.027

46.

S.J. Kim , J.Y. Shin , T.C. Cheong , I.J. Choi ,Y.S. Lee , S.H. Park , K.H. Chun, Galectin-3 germline variant at position 191 enhances nuclear accumulation and activation of β-catenin in gastric cancer. Clin Exp Metastasis. 28(2011) 743–750. doi:10.1007/s10585-011-9406-8

35

Figure legends

Fig.1. DHA inhibited the proliferation, migration and invasiveness of U87 and U251 cells in dose-dependent manner. The cell viability rate was assayed by CCK-8. Transwell migration and invasiveness assays were performed to investigate the changes in the migratory capacity and invasiveness of U87 and U251cells. (A) The cell inhibition rate of U87 and U251 cells after treated with DHA at various concentrations (50, 100, 200, 400, and 600 μmol/L) for 24 h. *P<0.05, #P<0.05 compared with control group. (B) The IC50 is 210.77±18.2 μmol/L for U87 cells and 200.35±18.2 μmol/L for U251 cells. (C) On condition that the density of U87 and U251 cells was 5×105 ml, 100, 200 and 400 μmol/L DHA could not change the viability of U87 and U251 cells significantly in 24h. (D) The change in the migration of U87 and U251 cells after treated with DHA at concentration of 100, 200 and 400μmol/L. *P<0.01 compared with control group. (E) The change in the invasion of U87 and U251 cells after treated with 100, 200 and 400μmol/L DHA. *P<0.01 compared with control group. n = 5. (F)Subcutaneous xenografts in nude mice. DHA inhibited the tumor growth in vivo. *P<0.05 compared with Control group. n=10

36

Fig.2. DHA inhibited the expression and activity of MMP9 and MMP7 in a dose-dependent manner. (A, B) The changes in the expression of MMP9 and MMP7 after U87 cells were treated with 100, 200 and 400 μmol/L DHA. (C, D) The changes in the expression of MMP9 and MMP7 of U251 cells. (E, F) The changes in the activity of MMP9 and MMP7 after U87 cells were treated with 100, 200 and 400 μmol/L DHA. (G, H) The changes in the expression of MMP9 and MMP7 of U251 cells. *P<0.01 compared with control group. n = 3.

37

Fig.3. DHA increased the generation of ROS and activated p53 in U87 and U251 cells. (A)The change in the generation of ROS after U87 cells were treated with 100, 200 and 400 μmol/L DHA. n = 3. (B) The change in ROS of U251. n = 3. (C, E, F, I) The changes in the expression of p53, p-p53 (S15) and p-p53(S15)/p53 in U87 cells. n = 5. (D G H J) The changes in the expression of p53, p-p53 (S15) and p-p53(S15)/p53 in U251 cells. n = 3. *P<0.05 compared with control group.

38

Fig.4. The influence of DHA on the expression of EGFR, β-catenin, p-β-catenin(Y333) and p-β-catenin(S45). (A-D) Western blot results of U87 cells and the statistical analysis. 100, 200 and 400 μmol/L DHA significantly down-regulated the expression of

EGFR,

p-β-catenin(Y333)

and

β-catenin,

lowered

the

ratio

of

p-β-catenin(Y333)/β-catenin, up-regulated p-β-catenin(S45) and raised the ratio of p-β-catenin(S45)/β-catenin in U87 cells. (E-H) DHA combined with EGFR inhibitor further up-regulated p-β-catenin(S45) and down-regulated p-β-catenin(Y333) and 39

β-catenin compared with the DHA group. (I-L) Western blot results of U251 cells and the statistical analysis. 100, 200 and 400 μmol/L DHA significantly up-regulated the expression

of

EGFR,

p-β-catenin(Y333/S45)

and

raised

the

ratio

of

p-β-catenin(Y333S45)/β-catenin in U251 cells while β-catenin expression remained unchanged. (M-P) DHA combined with EGFR inhibitor significantly down-regulated β-catenin and p-β-catenin(Y333) and up-regulated p-β-catenin(S45) in U251 cells. *

P<0.05 compared with control group; #P<0.01 compared with DHA group. n = 3.

Fig.5. Immunofluorescence staining of β-catenin(A,B), p-β-catenin(Y333) (C,D)and p-β-catenin(S45) (E,F)in U87 and U251 cells. β-catenin, p-β-catenin(Y333) and p-β-catenin(S45) (green) were labeled with secondary antibody and nuclei (blue) were labeled with DAPI. 40

Fig.6.The

stable

transfection

efficiency

of

p53Mut/WT,

β-catenin

and

p-β-catenin(Y333/S45)Mut . *P<0.01 compared with control group. n=3

Fig.7. Effects of DHA on the expression of p-β-catenin(Y333/S45), β-catenin, and MMP9/7 activity by the transfection of wild type p53 or mutant type p53 gene. (A-E) In U87 cells, stable transfection of mutant type p53 significantly promoted the

41

expression of β-catenin, p-β-catenin(Y333/S45) and MMP9/7, the ratio of p-β-catenin(Y333)/β-catenin and p-β-catenin(S45)/β-catenin compared with the DHA+NC group. (F-J) In U251 cells, stable transfection of wild type p53 significantly decreased the expression of β-catenin, p-β-catenin(Y333), MMP9/7 and the ratio of p-β-catenin(Y333)/β-catenin and increased p-β-catenin(S45) expression and p-β-catenin(S45)/β-catenin ratio compared with the DHA+NC group. (K) Gelatin zymography results of U87 cells. The stable transfection of mutant type p53 significantly promoted the activity of MMP9 and MMP7. (L) Gelatin zymography results of U251 cells. The stable transfection of wild type p53 significantly inhibited the activity of MMP9 and MMP7. *P<0.01 compared with control group; #P<0.01 compared with DHA +NC group. n=3

42

Fig.8. The effects of sh-β-catenin, pIRES2-β-catenin, p-β-catenin(Y333)-Mut, p-β-catenin(S45)-Mut on the migration and invasion of U87 and U251 cells. (A) Results of migration in U87 and U251 cells under the treatment of sh-β-catenin. Sh-β-catenin further inhibited the migration in U87 and U251 cells compared with the 43

DHA group. (B) Results of invasion in U87 and U251 cells under the treatment of sh-β-catenin. Sh-β-catenin further inhibited the invasion in U87 and U251 cells compared with the DHA group. (C) Results of migration in U87 and U251 cells under the

treatment

of

pIRES2-β-catenin,

pIRES2-p-β-catenin(Y333)-Mut

or

pIRES2-p-β-catenin(S45)-Mut. pIRES2-p-β-catenin(Y333)-Mut further inhibited the migration in both cell lines compared with the DHA+NC group. PIRES2-β-catenin and pIRES2-p-β-catenin(S45)-Mut promoted the migration in U87 and U251 cells. (D) Results of invasion in U87 and U251 cells under the treatment of pIRES2-β-catenin, pIRES2-p-β-catenin(Y333)-Mut

or

pIRES2-p-β-catenin(S45)-Mut.

pIRES2-p-β-catenin(Y333)-Mut further inhibited the invasion in both cell lines compared

with

the

DHA

+NC

group.

PIRES2-β-catenin

and

pIRES2-p-β-catenin(S45)-Mut promoted the invasion in U87 and U251 cells. *P<0.05 compared with control group, #P<0.05 compared with DHA +NC group. n=3

44

Fig.9.

The

effects

of

sh-β-catenin,

pIRES2-β-catenin,

pIRES2-p-β-catenin(Y333)-Mut, pIRES2-p-β-catenin(S45)-Mut, on the expression and activity of MMP9 and MMP7 in U87 and U251 cells. (A) Western blot results of U87 cells under the treatment of sh-β-catenin. Sh-β-catenin further inhibited the the expression and activity of MMP9 and MMP7 in U87 cells. (B) Western blot results of U251 cells under the treatment of sh-β-catenin. Sh-β-catenin further inhibited the expression of MMP9 and MMP7 in U251 cells. (C) Gelatin zymography results of U87 cells under the treatment of sh-β-catenin. Sh-β-catenin further inhibited the activity of MMP9 and MMP7 in U87 cells. (D) Gelatin zymography results of U251 cells under the treatment of sh-β-catenin. Sh-β-catenin further inhibited the activity of 45

MMP9 and MMP7 in U251 cells. (E) Western blot results of U87 cells under the treatment

of

pIRES2-β-catenin,

pIRES2-p-β-catenin(Y333)-Mut

or

pIRES2-p-β-catenin(S45)-Mut. pIRES2-p-β-catenin(Y333)-Mut further inhibited the expression of MMP9 and MMP7 compared with the DHA+NC group. PIRES2-β-catenin and p-β-catenin(S45)-Mut promoted the expression of MMP9 and MMP7 in U87 cells. (F) Western blot results of U251 cells under the treatment of pIRES2-β-catenin,

p-β-catenin(Y333)-Mut

or

p-β-catenin(S45)-Mut.

pIRES2-p-β-catenin(Y333)-Mut further inhibited the expression of MMP9 and MMP7

compared

with

the

DHA+NC

group.

PIRES2-β-catenin

and

pIRES2-p-β-catenin(S45)-Mut promoted the expression of MMP9 and MMP7 in U251 cells. (G) Gelatin zymography results of U87 cells under the treatment of pIRES2-β-catenin, pIRES2-p-β-catenin(Y333)-Mut or pIRES2-p-β-catenin(S45)-Mut. pIRES2-p-β-catenin(Y333)-Mut further inhibited the activity of MMP9 and MMP7 compared

with

the

DHA+NC

group.

pIRES2-PIRES2-β-catenin

and

pIRES2-p-β-catenin(S45)-Mut promoted the activity of MMP9 and MMP7 in U87 cells. (H) Gelatin zymography results of U251 cells under the treatment of pIRES2-β-catenin, pIRES2-p-β-catenin(Y333)-Mut or pIRES2-p-β-catenin(S45)-Mut. pIRES2-p-β-catenin(Y333)-Mut further inhibited the activity of MMP9 and MMP7 compared

with

the

DHA+NC

group.

PIRES2-β-catenin

and

pIRES2-p-β-catenin(S45)-Mut promoted the activity of MMP9 and MMP7 in U251 cells. *P<0.05 compared with control group; #P<0.05 compared with DHA +NC group. n=3. (I) DHA (10 mg/kg)

reduced the tumor size.Over-expressed

β-catenin(pIRES2-β-catenin) could reduce the efficiacy of DHA,while sh-β-catenin enhanced it. *P<0.05 compared with Control group; #P<0.05 compared with DHA group. n=10 46

Fig.10. The expression of p53 had not changed significantly after overexpressing or silencing β-catenin in both U87 and U251 cells.n=3.

47

Fig.11. (AB)There is binding site between β-catenin and MMP9/7.Schematic representation of the human β-catenin promoter region 3,000 bp upstream of the transcription start site (TSS) which designated as +1. Putative β-catenin binding sites are indicated. Chromatin immunoprecipitation (ChIP) PCR products for binding sites and an upstream region that does not bind to β-catenin are depicted with bold lines. Dashed arrows represent the primers used for each PCR. Glioma cells were used to conduct ChIP assays. PCR was conducted with the resulting precipitated DNA. Images are representative of independent experiments. n=3. (CD) Luciferase activity in the β-catenin/Kp700 or Kp100 transfected cells,whose reporter lacked the selected TCF4 binding site, was weaker than that of the β-catenin/ Kp2000 transfected cells, whose reporter included the TCF4 binding site (Fig.10 CD, *P<0.01).n=4.

48

49

Table 1 Groups of cells Groups

Transfection

Drug treatment

Control

NA

NA

DHA group

NA

DHA (200 μmol/L)

DHA+p53-WT

Wild P53

DHA (200 μmol/L)

DHA+p53-Mut

Mutant P53

DHA (200 μmol/L)

DHA+sh-β-catenin

sh-β-catenin

DHA (200 μmol/L)

DHA+pIRES2-β-catenin

pIRES2-β-catenin

DHA (200 μmol/L)

DHA+ pIRES2-β-catenin(S45)-Mut

pIRES2-EGFP-β-catenin(S45)

DHA (200 μmol/L)

DHA+ pIRES2-β-catenin(Y333)-Mut

pIRES2-EGFP-β-catenin(Y333)

DHA (200 μmol/L)

50