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Introduction

Glioblastoma multiforme (GBM; WHO grade IV glioma) is the most malignant brain tumor in adults. Even following treatment with surgical resection, radiotherapy and concomitant chemotherapy, the median survival time of patients with GBM is only 14.6 months (1). A number of molecular markers for GBM have been identified and are associated with diagnosis, prognosis and treatment. For example, somatic mutations in isocitrate dehydrogenase 1 (IDH1) have been identified in GBM patients, particularly in secondary GBM, which evolves from lower-grade gliomas (2). In 90% of IDH1 mutations in gliomas, the arginine at position 132 is replaced by a histidine (R132H mutation) (3). Patients with IDH1R132H gliomas also have significantly improved survival rates (4). However, the mechanism responsible for this improved survival rate remains to be elucidated.

MicroRNAs (miRNAs) are a class of non-coding, single-stranded RNAs comprised of ~22 nucleotides (5). miRNAs are post-transcriptional regulators that bind complementary sequences on target mRNA transcripts. Target binding usually results in gene regulation by translational repression or mRNA degradation, thereby modulating a variety of biological processes (6). Several miRNAs are known to associate with clinical outcomes in GBM (5,7,8), including miR-128a, a crucial regulator in brain development (9-11). Aberrant expression of miR-128a has been detected in glioma (12) and is involved in cancer-associated biological processes, including cell proliferation, differentiation and apoptosis (13,14).

In order to evaluate the effect of the IDH1R132H mutation and its association with miR-128a, a clonal U87 cell line overexpressing the IDH1R132H mutant protein was generated. The present study also investigated the functional molecules upstream and downstream of miR-128a in IDH1R132H overexpressing cells.

Materials and methods

Cell culture and reagents

U87 glioma cells and HEK 293T cells were obtained from the American Type Culture Collection (Manassas, VA, USA) and were cultured in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% heat-inactivated fetal bovine serum (FBS; Invitrogen Life Technologies, Carlsbad, CA, USA). SYBR Green PCR master mix and the TaqMan microRNA reverse transcription kit were purchased from Applied Biosystems (Foster City, CA, USA). YC-1 was obtained from Sigma-Aldrich (St. Louis, MO, USA) and dissolved in dimethyl sulfoxide.

Generation of constructs

The full-length human IDH1 coding sequence was amplified from HEK 293T cells (Sangon Biotech Co., Ltd., Shanghai, China). The cDNA was fused in-frame with a FLAG tag at the N-terminus using the following synthesized primers: Forward primer with MluI site: TTTCGTACGATGGATTACAAGGACGACGAT GACAA GTCCAAAAAAAT and reverse primer with BsiWI site: TTTACGCGTGGTATGAACTTAAAGTTTGG. The amplified target was inserted into the MluI- and BsiWI-linearized pHR-SIN vector. The IDH1R132H mutation was generated in the vector described above by QuickChange (Stratagene, Santa Clara, CA, USA) with the following primer sequences: R132H, forward 5′-ACCTATCATCATAGGTCATCATGCTTA TGGG-3′ and reverse 5′-TGACCTATGATGATAGGTT TTACCCATCCAC-3′.

Stable overexpression of IDH1WT and IDH1R132H constructs in U87 cells

HEK 293T cells were seeded in 60 mm plates (4×106 cells/plate) in DMEM with 10% heat-inactivated FBS 1 day prior to transfection. Cells were transfected with 5.2 μg of either pHR-SIN-IDH1WT or pHR-SIN-IDH1R132H, along with 2.36 μg of pSPAX2 and 0.8 μg of pMD2 G plasmids using Lipofectamine 2000 in DMEM. After 6 h, the transfection media was removed and replaced with antibiotic-free DMEM with 10% heat-inactivated FBS. Lentiviral particles were harvested at 72 h post-transfection. U87 glioma cells were plated 1 day prior to transduction, at 20% confluency, in 60 mm plates with DMEM containing 10% phosphate-buffered saline (PBS). To transduce cells, 3 ml of conditioned media containing viral particles and 6 μg/ml polybrene (Sigma-Aldrich) were added to each culture. After 16 h, the conditioned media was removed and replaced with DMEM containing 15% FBS.

Protein extraction and western blotting

Cells prepared for protein extraction were immediately placed on ice and washed with ice-cold PBS solution. Total protein extraction was achieved using RIPA lysis buffer containing protease inhibitors (Roche Diagnostics, Mannheim, Germany) and PMSF. Proteins were resolved on 8-12% SDS-polyacrylamide gels (Sangon Biotech Co. Ltd., Shanghai, China) and transferred onto nitrocellulose membranes (Bio-Rad Laboratories, Inc., Hercules, CA, USA). The membranes were blocked in Tris-buffered saline and Tween 20 (TBST) with 5% non-fat dry milk for 1 h and incubated overnight at 4°C with the following respective primary antibodies in TBST with 5% bovine serum albumin: Rabbit monoclonal anti-human IDH1 (1:1,000; cat. no. 8137; Cell Signaling Technology, Inc., Danvers, MA, USA), mouse monoclonal anti-human IDH1R132H (1:200; cat. no. SAB4200548; Sigma-Aldrich), mouse monoclonal anti-FLAG M2 (1:500; cat. no. F3165; Sigma-Aldrich) or mouse monoclonal anti-human HIF-1α (1:1,000; cat. no. AH339; Beyotime Institute of Biotechnology, Shanghai, China). Immunospecific bands were detected using an ECL substrate (Sigma-Aldrich). β-actin immunoblotting was performed as a loading control.

Reverse transcription quantitative polymerase chain reaction (RT-qPCR) analysis

Total RNA was isolated from cells using TRIzol reagent (Invitrogen Life Technologies). Total RNA (1 μg) was used as a template for a Moloney murine leukemia virus-RT reverse transcriptase reaction, which was performed according to the manufacturer's instructions (Fermentas, Vilnius, Lithuania). qPCR was performed using SYBR Green Master mix (Applied Biosystems). β-actin amplification was used as an internal control. To quantitate the expression level of miR-128a, 1 μg of total RNA was reverse transcribed in 50 μl reaction using TaqMan Reverse Transcription Reagents (Applied Biosystems) with the stem loop primer. Reverse transcribed cDNA (2 μl) was subjected to SYBR Green mix and the results were normalized to RNU48.

The following primers were used: RNU48, forward 5′-TGATGATGACCCCAGGTAACTC-3′ and reverse 5′-GAG CGCTGCGGTGATG-3′; Bmi-1, forward 5′-CTGCAGCTC GCTTCAAGATG-3′ and reverse 5′-CACACACATCAGGT GGGGAT-3′; β-actin, forward 5′-ATGACTTAGTTGCGTTAC ACC-3′ and reverse 5′-TGCTGTCACCTTCACCGTTC-3′; miR-128a stem loop primer: 5′-GTCGTATCCAGTGCAGGG TC CGAGGTATTCGCACTGGATACGACAAAGAG-3′; miR-128a, forward 5′-TCGCGTTCACAGTGAA-3′ and reverse 5′-GTGCAGGGTCCGAGG-3′.

Chromatin immunoprecipitation (ChIP) assays

ChIP assays were performed using a ChIP kit (cat. no. 17-371; EMD Millipore, Billerica, MA, USA) according to the manufacturer's instructions. Cells were crosslinked with 1% formaldehyde for 20 min at 37°C and quenched in 0.125 M glycine. DNA was immunoprecipitated from sonicated cell lysates and quantified using SYBR Green Real-time PCR analysis. The following primer sequences were used: Forward: 5′-GTAATAGAATTTTCATATTG-3′ and reverse: 5′-ATTTTGCCATGTTGAAGAAC-3′. Fold enrichment was calculated based on Ct as 2−Δ(ΔCt), where ΔCt = CtIP – CtInput and Δ(ΔCt) = ΔCtantibody − ΔCtIgG.

Cell proliferation

Briefly, 5,000 cells per well were plated in 96-well plates and cultured for 96 h. Following incubation, CCK-8 (10 μl) was added to each well and allowed to incubate at 37°C for 2 h. Cell proliferation was then quantified by measuring the optical density at a wavelength of 450 nm (SpectraMax M2 Multimode Microplate Reader; Molecular Devices, LLC, Sunnyvale, CA, USA). Each experiment was performed in triplicate.

Statistical analysis

The data are presented as the mean ± standard deviation. The samples were analyzed using a two-tailed unpaired Student's t-test, unless otherwise noted. P<0.05 was considered to indicate a statistically significant difference.

Results

Overexpression of IDH1R132H decreases U87 cell proliferation

To determine the effects of glioma-associated IDH1R132H, N-terminal FLAG-tagged IDH1WT, IDH1R132H and control expression constructs were generated for stable expression in U87 cells (Fig. 1A). Western blot analysis with FLAG, IDH1 and IDH1R132H specific antibodies confirmed the expression of the appropriate IDH1 protein products in U87 cell lines (Fig. 1B). Previous clinical studies demonstrated that patients with IDH1R132H-positive tumors have a significant survival advantage (4,15). However, the role of IDH1R132H in glioma cell proliferation is not fully understood. To assess the role of IDH1R132H in U87 cell proliferation, CCK-8 proliferation assays were performed. The proliferation of U87 cells overexpressing IDH1R132H was significantly decreased compared with the control and IDH1WT-expressing cells over a 4-day incubation period (Fig. 1C; P<0.01).

Figure 1 Overexpression of IDH1R132H decreases U87 cell proliferation. (A) Schematic diagram of the FLAG-IDH1 fusion protein amino acid sequence. The FLAG tag was linked to the N-terminus of IDH1. Red indicates the FLAG tag and mutation at amino acid residue 132 of IDH1. (B) Representative western blot analysis for FLAG-tagged IDH1WT and IDH1R132H in stably transfected U87 cells. (C) CCK-8 proliferation assays were performed after a 4 day incubation of U87 cells containing IDH1WT, IDH1R132H or control expression vectors. **P<0.01. IDH1, isocitrate dehydrogenase 1; OD, optical density.

Overexpression of IDH1R132H in U87 cells induces the expression of miR-128a

Several miRNAs associated with IDH1 mutations have been revealed via miRNA expression profiling and the miRNA expression signature has been identified as a prognostic biomarker for GBM patients (7,11). However, whether IDH1R132H induces a specific miRNA that controls cell proliferation remains to be elucidated. According to the IDH1 mutation-specific 23-miRNA signature analyzed by Wang et al (11), miR-128a was significantly overexpressed in IDH1 mutant patient samples compared with wild-type samples. miRNA expression data was analyzed using The Cancer Genome Atlas (TCGA) portal (https://tcga-data.nci.nih.gov/tcga/), which contains data from seven IDH1R132H and 133 wild-type IDH1 patients. In this dataset, miR-128a was overexpressed in IDH1R132H compared with wild-type IDH1 patients (Fig. 2A; P=0.0078). To confirm this expression pattern, qPCR was performed to detect miR-128a levels in IDH1-expressing U87 cells. In agreement with the clinical samples, U87 cells overexpressing IDH1R132H had significantly higher levels of miR-128a compared with control and IDH1WT cells (Fig. 2B; P<0.01).

Figure 2 Overexpression of IDH1R132H in U87 cells induces the expression of miR-128a. (A) Analysis of miR-128a expression from The Cancer Genome Atlas in IDH1R132H patients compared with wild-type IDH1 patients. (B) Quantitative polymerase chain reaction analysis of miR-128a expression in IDH1WT, IDH1R132H and control overexpressing U87 cells. **P<0.01. IDH1, isocitrate dehydrogenase 1; miR-128a, microRNA-128a.

HIF-1α is required for miR-128a expression induced by IDH1R132H

The ectopic expression of mutant IDH1 in cultured cells also increases the levels of HIF-1α (16). It was hypothesized that increased HIF-1α expression may therefore control IDH1R132H-induced miR-128a expression. In agreement with this hypothesis, HIF-1α protein levels were found to be significantly higher in U87 cells overexpressing IDH1R132H compared with the control and IDH1WT cells (Fig. 3A). In order to assess whether HIF-1α is required for miR-128a overexpression in IDH1R132H cells, cells were treated with YC-1, an inhibitor of HIF-1α activity (17). YC-1 significantly decreased the miR-128a level in IDH1R132H cells, however, similar effects were not observed in the control and IDH1WT cells (Fig. 3B; P<0.01). Notably, there is a conserved HIF-1α binding sequence GCGTG (hypoxia response element; HRE) ~2.5 kilobases upstream of the gene encoding pri-miR-128a (Fig. 3C). Therefore, HIF-1α could potentially regulate miR-128a expression by binding to this HRE. To investigate whether HIF-1α binds this element in the promoter of the miR-128a gene, ChIP assays were performed. In IDH1R132H overexpressing U87 cells, HIF-1α was in fact enriched at this HRE suggesting that it regulates miR-128a expression (Fig. 3D; P<0.01). Taken together, the data demonstrated that HIF-1α binds the HRE in the pri-miR-128a promoter and is required for IDH1R132-induced miR-128a expression.

Figure 3 HIF-1α is required for miR-128a expression induced by IDH1R132H. (A) Western blot analysis of HIF-1α expression in IDH1WT, IDH1R132H and control overexpressing U87 cells. (B) U87 cells were treated with 5 μM YC-1, a HIF-1α inhibitor, and the level of miR-128a was determined by quantitative polymerase chain reaction. (C) Diagram of the putative HRE site in the promoter of miR-128a. (D) Chromatin immunoprecipitation was performed in IDH1R132H-overexpressing U87 cells with an HIF-1α-specific antibody. **P<0.01. IDH1, isocitrate dehydrogenase 1; HIF-1α, hypoxia inducible factor-1α; HRE, hypoxia response element; miR-128a, microRNA-128a; DMSO, dimethyl sulfoxide; IgG, immunoglobulin G; NC, negative control.

miR-128a regulates the decreased proliferation rate induced by IDH1R132H

Previous studies have reported that miR-128a is downregulated in human glioblastoma (18) and its expression inhibits cell growth in vitro and in vivo (19). To examine the biological function of miR-128a expression, U87 cells were transfected with miR-128a mimics or the miR-128a inhibitor and CCK-8 proliferation assays were performed. Overexpression of miR-128a decreased the cell proliferation rate, whereas, knockdown of miR-128a with the miR-128a inhibitor enhanced cell proliferation (Fig. 4A; P<0.01). IDH1R132H overexpressing cells were also treated with the miR-128a inhibitor, which rescued the decreased proliferation rate induced by IDH1R132H (Fig. 4B; P<0.01). miR-128a causes a marked decrease in the expression of the oncogene Bmi-1 in U87 cells by direct regulation of the Bmi-1 mRNA 3′-untranslated region (19). Therefore, it was hypothesized that Bmi-1 may be involved in the decreased proliferation of IDH1R132H cells. qPCR was performed to detect the Bmi-1 level in IDH1R132H overexpressing cells. The results demonstrated that it was decreased compared with the control and IDH1WT cells (Fig. 4C; P<0.01). Furthermore, overexpression of miR-128a decreased the mRNA expression of Bmi-1, whereas inhibition of miR-128a enhanced Bmi-1 expression (Fig. 4D; P<0.01). Taken together, it can be concluded that IDH1R132H restricts cell proliferation by inducing the expression of miR-128a, which downregulates Bmi-1 expression.

Figure 4 miR-128a regulates the decreased proliferation rate induced by IDH1R132H. (A) CCK-8 proliferation assay of U87 cells treated with 50 nM miR-128a mimics or the miR-128a inhibitor. (B) CCK-8 proliferation assay of IDH1WT, IDH1R132H and control overexpressing U87 cells treated with the miR-128a inhibitor or scramble control. (C) qPCR analysis of Bmi-1 expression in IDH1WT, IDH1R132H and control overexpressing U87 cells. (D) qPCR analysis of Bmi-1 expression in U87 cells treated with miR-128a mimics, the miR-128a inhibitor or scramble control. **P<0.01. IDH1, isocitrate dehydrogenase 1; miR-128a, microRNA-128a; qPCR, quantitative polymerase chain reaction; Bmi-1, B-cell-specific Moloney murine leukemia virus integration site 1 protein.

Discussion

Clinical data demonstrate that patients with the IDH1R132H mutation have an improved outcome compared with those with the wild-type sequence. However, the mechanistic role by which IDH1R321H regulates tumor development is not fully understood. In the present study, the functional impact of the tumor-associated IDH1R132H mutation was examined in U87 glioma cells in vitro. Following establishing stable expression of IDH1R132H in U87 cells, it was found that it reduced cell proliferation, which is consistent with the findings of Bralten et al (20). This lends support to the theory that IDH1 mutations protect against tumor proliferation and may explain why IDH1R132H is an independent favorable prognostic marker in glioma patients (15,21).

Wang et al reported that the IDH1 mutation-specific microRNA signature predicted a favorable prognosis in glioblastoma patients with the IDH1 wild type (11). However, it remained to be determined whether the IDH1R132H mutation regulated cell growth through microRNAs. miR-128a expression is markedly reduced in human glioblastoma specimens compared with adjacent brain samples without tumors (18,19,22). A previous study suggested that loss of miR-128 expression is an early event in gliomagenesis (23). By analyzing the TCGA data, miR-128a was found to be overexpressed in human IDH1R132H GBM patients compared with wild-type IDH1 patients. In addition, miR-128a expression was increased in IDH1R132H overexpressing U87 glioma cells. The result demonstrating that miR-128a is upregulated in IDH1R132H gliomas is noteworthy given that miR-128a is downregulated in IDH1WT glioblastomas (11). This demonstrates that increased miR-128a expression is an important characteristic of mutant IDH1R132H gliomas.

miR-128a is downregulated in GBM, suggesting that it may function as a tumor suppressor in brain tumor progression (24) and may be associated with the survival of GBM patients (11). In U87 cells, overexpression of miR-128a suppressed cell proliferation. By contrast, knockdown of miR-128a increased cell proliferation. Our findings are consistent with a study on miR-128a in medulloblastoma cells (25). Furthermore, the present study illustrated that inhibition of miR-128a in IDH1R132H overexpressing U87 cells reversed the cell proliferation induced by IDH1R132H, suggesting that miR-128a is responsible for IDH1R132H-induced cell proliferation.

HIF-1α, an important transcription factor of the cellular response to hypoxia, is known to facilitate tumor growth when oxygen is low (26). The stability of HIF-1α is regulated by α-ketoglutaric acid, which is an important enzyme product of IDH1 (27). The results demonstrated that overexpression of the IDH1R132H mutant increased HIF-1α protein levels in U87 glioma cells, which is consistent with a former study by Zhao et al (16). In the present study, miR-128a expression was significantly decreased when the activity of HIF-1α was inhibited, which indicated that HIF-1α may regulate miR-128a in IDH1R132H cells. In addition, an HRE located in the promoter upstream of the gene encoding pri-miR-128a was identified and HIF-1α was found to specifically bind the site in IDH1R132H over-expressing U87 cells. This suggests that the transcription factor HIF-1α regulates miR-128a expression. Currently, to the best of our knowledge, the present study is the first to demonstrate that HIF-1α acts as a regulator of IDH1R132-dependent miR-128a expression by directly binding an HRE within its promoter.

Bmi-1, a direct target of miR-128a, is upregulated in several types of cancer, including gliomas and is a positive regulator of neural stem cell renewal (19,28–31). The present study demonstrated that Bmi-1 was downregulated by miR-128a in IDH1R132H overexpressing cells. Bmi-1 is an oncogene involved in regulating cell proliferation as a transcriptional repressor (19), and studies in transgenic mice revealed a critical role for Bmi-1 in driving glioma growth (32). The present data indicated that inhibition of Bmi-1 may be a mechanism of miR-128a-mediated growth restriction in IDH1R132H over-expressing cells.

In conclusion, the present study described the function and mechanism of action of miR-128a in IDH1R132H glioma cells. The results of the present study demonstrated that the IDH1R132H mutation leads to an increase in the levels of HIF-1α protein, which binds to an HRE in the miR-128a promoter to induce its expression. The enhanced expression of miR-128a subsequently inhibits Bmi-1 expression, thereby decreasing cell proliferation in IDH1R132H glioma cells. The present study provides novel insight into the pathophysiology of miR-128a in IDH1 mutant glioma. Finally, our characterization of the IDH1R132H-HIF-1α-miR-128a-Bmi-1 pathway may have therapeutic implications for the treatment of glioma.

Acknowledgments

This study was supported by the Shanghai Science and Technology Committee (grant no. 13XD1402600), Shanghai Health and Family planning Commission (grant no. 2013SY024) and the State Key Laboratory of Oncogenes and Related Genes (grant no. 90-14-01), Shanghai Cancer Institute, Ren Ji Hospital, School of Medicine, Shanghai Jiao Tong University.

References