大阪府立大学　学術情報リポジトリ

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Transcription-dependent and -independent

Regulatory Mechanisms by Hypoxia-inducible

Factors in Cancer Cells

著者

東村泰希

内容記述

学位授与大学: Osaka Prefecture University(大阪

府立大学), 学位の種類: 博士(応用生命科学), 学

位記番号: 論生命第33号, 学位授与年月日:

2011-03-31, 指導教員: 乾博.

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大阪府立大学博士（応用生命科学）学位論文

Transcription-dependent and -independent Regulatory

Mechanisms by Hypoxia-inducible Factors in Cancer Cells

がん細胞における低酸素誘導因子を介した転写依存的

および非依存的な調節機構

YASUKI HIGASHIMURA

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ABBREVIATIONS

AEBSF 4-(2-aminoethyl)-benzensulfonyl fluoride bHLH basic helix-loop-helix

ChIP Chromatin immunoprecipitation C-TAD C-terminal transactivation domain DFX deferoxamine mesylate

dFBS dextran-coated charcoal-stripped FBS DMEM-HG DMEM-high glucose

DBD DNA-binding domain

DMEM Dulbecco’s modified Eagle’s medium Epo erythropoietin

ERα estrogen receptor α E2 17β-estradiol

FBS fetal bovine serum

GAPDH glyceraldehyde-3-phosphate dehydrogenase HA hemagglutinin

HIF hypoxia-inducible factor HRE hypoxia response element LBD ligand-binding domain NTD N-terminal domain

N-TAD N-terminal transactivation domain ODD oxygen-dependent degradation PAS PER-ARNT-SIM

PIP Plasmid immunoprecipitation PHDs prolyl hydroxylase domain proteins RLU relative light unit

RT-PCR reverse transcription-PCR VHL von Hippel-Lindau protein

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INTRODUCTION

Molecular oxygen acts as the terminal electron acceptor in mitochondrial oxidative phosphorylation, and involves in efficient ATP production, which is essential for most biological processes. In mammals including human, hypoxia is the biological state of oxygen starvation and is encountered acutely and chronically due to cardiopulmonary diseases, pernicious anemia, breakage of blood vessel by injury, and residence at high altitudes. Mammals have evolved specialized systems that respond to hypoxia by altering cell metabolism and survival and by rescuing oxygen delivery [1]. Hypoxia-inducible factor (HIF) is a transcription factor complex and functions as a master regulator for these hypoxic responses. Oxygen-dependent regulation of HIF activity controls the expression of a wide range of genes that allow the cell to respond to change in oxygen availability [2].

HIF is a heterodimeric complex consisting of an α subunit (HIF-1α or HIF-2α) and a β subunit (HIF-1β) [3, 4]. The latter subunit is known as aryl hydrocarbon receptor nuclear translocator, and is constitutively expressed. These subunits belong to the basic helix-loop-helix (bHLH)/PER-ARNT-SIM (PAS) family transcriptional factors. HIF-α proteins possess an oxygen-dependent degradation (ODD) domain and two transcriptional activation domains, called the N-terminal and C-terminal transactivation domains (N-TAD and C-TAD, respectively). Under normoxic conditions, the two conserved proline residues (at positions 402 and 564 in human HIF-1α, and 405 and 531 in human HIF-2α) within the ODD domain of HIF-α proteins are hydroxylated by a family of oxygen-dependent prolyl hydroxylase domain proteins (PHD1, PHD2, and PHD3). The resultant modified HIF-α proteins are recognized and polyubiquitinated by an E3 ubiquitin ligase complex, which contains the von Hippel-Lindau tumor suppressor (VHL) protein, and subsequently undergo degradation through the 26S proteasome [5]. In contrast, under hypoxic conditions, prolyl hydroxylation is inhibited, and consequently, HIF-α is stably expressed and binds to hypoxia response elements (HREs: 5´-RCGTG-3´) of the target genes together with HIF-1β (Fig. 1).

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HIF-1α and HIF-2α display the highest degree of sequence homology in the bHLH (85 %) and PAS (70 %) domain (Fig. 2). Despite these similarities, expression profiling and functional studies have revealed that the HIF-α subunits regulate both common and unique target genes [2, 6-8]. For instance, HIF-1α uniquely stimulates the expression of glycolytic enzymes [9], such as phosphoglycerate kinase and lactate dehydrogenase-A, carbonic anhydrase-9 [10], and the pro-apoptotic gene BNIP-3 [11]. In contrast, the embryonic transcription factor Oct-4 [12], Cyclin D1 [13], TWIST1 [14], transforming growth factor-α [15], and erythropoietin (EPO) [16, 17] are up-regulated by hypoxia in a HIF-2α-dependent manner. The gene expressions of vascular endothelial growth factor, adrenomedullin, and glucose transporter 1 are regulated by both α subunits [11, 18]. In contrast, the transcription-independent functions of the

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HIF-α subunits have recently been reported. HIF-1α functionally antagonizes c-Myc transcriptional activity under hypoxia, leading to cell cycle arrest. In contrast, HIF-2α promotes cell cycle progression by enhancing c-Myc transcriptional activity [19]. Furthermore, HIF-1α is stable in an acute hypoxia, and the HIF-1α protein level is decreased or disappears during prolonged hypoxia. On the other hand, HIF-2α protein level remains stabilized during chronic hypoxia [20]. Thus, HIF-1α and HIF-2α play different roles in tumor progression and do not function redundantly.

Solid tumors contain the regions which exhibit a very low oxygen concentration, because vasculature in tumors is dysfunctional and insufficient vascularization due to the rapidly growing tumor cells results in an insufficient oxygen supply [21]. In tumor cells, pathophysiological processes such as angiogenesis and glycolysis are activated to adapt to the hypoxic environment [22, 23]. HIF-1α and HIF-2α proteins are expressed in most types of tumors including bladder, breast, colon, glial, hepatocellular, ovarian, pancreatic, prostate and renal tumors [24]. The identification of novel molecular target for cancer therapy has led to a paradigm shift in drug development. HIF signaling is considered to be one of the molecular target because HIF functions play key roles not only in tumorigenesis, but also in resistance to chemotherapy and radiotherapy [25, 26] [27]. Indeed, many HIF inhibitors that target to HIF-α mRNA expression, HIF-α

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protein translation, HIF binding to DNA, and HIF transcriptional activity, are indentified [28-30]. However, most of the inhibitors are the lack of specificity, indicating the fact that HIF inhibition cannot be easily separated from other biological activities. Because of the feature of these inhibitors, HIF signaling may still have potential applications for cancer therapy. Thus, it is important to uncover the more precise and more specific to tissues molecular mechanism of HIF signaling.

In the present thesis, I have studied on the transcription-dependent and -independent functions of HIF-α subunits to provide a new approach for cancer therapy. Although the mechanisms by which glyceraldehyde-3-phosphate dehydrogenase (GAPDH) is induced by hypoxia mainly depend on the transcriptional activities of HIFs, hypoxic induction of GAPDH is specific to cell-types [31]. Therefore, in CHAPTER I, I hypothesize that the mechanisms by which hypoxia activate GAPDH expression through the transcriptional function of HIFs are different among cell types and focus on the regulatory mechanism of GAPDH gene expression under hypoxic conditions. In CHAPTER II, to provide a novel tool for inhibition of transcription-dependent function of HIF, I construct and characterize dominant negative isoform of HIF-2α. In CHAPTER III, I identify the transcription-independent function of HIF-2α as a negative regulator of estrogen receptor α (ERα) expression in breast cancer cells.

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CHAPTER I

Hypoxia-inducible Factor-1 Activity Depending on Sp1 Up-regulates

the Expression of Gyceraldehyde-3-phosphate Dehydrogenase Gene in

Hypoxic Breast Cancer Cells

GAPDH is a classical glycolytic enzyme for energy production in the cytosol and functions as a homotetramer. GAPDH reversibly catalyzes the oxidative phosphorylation of glyceraldehyde 3-phosphate to 1,3-bisphosphoglycerate. Although GAPDH is used as a housekeeping gene in many studies, GAPDH expression is up-regulated by insulin [32], calcium [33], and hypoxia [34]. Increased expression of GAPDH is observed in several tumors such as prostate, breast, lung, and cervical carcinomas [35-38], and hypoxia induces GAPDH gene expression in prostate cancer LNCaP cells and hepatoma Hep3B cells [39, 40]. In contrast, tumor cells such as hepatoma cells (HepG2 and Hep3B), colon cancer cells (HT-29 and HCT-116), lung adenocarcinoma cells (A-549), and glioblastoma cells (U373, U251, and GaMG) show no alteration of GAPDH gene expression in response to hypoxia [41, 42]. Thus, although contradictory results in hepatoma cells have been reported, hypoxia-induced transcription of GAPDH seems to be specific to tumor cell-types. If HIF involves the regulation of GAPDH gene, the molecular mechanism of HIF would be cell-type specific and novel signaling.

The nucleotide sequence from –1091 to +25 of the human GAPDH gene has six potential HREs. Previous studies [39, 40, 43] report that the HRE that is located between –125 and –121 upstream of the transcription start site of human GAPDH gene is functional in endothelial cells and hepatoma cells (Hep3B) and that two additional HREs between –217 and –203 are also functional in prostate cancer LNCaP cells. Thus, the molecular mechanisms of GAPDH gene expression by hypoxia are specific to tumor cell-types. In breast cancer, an increased expression level of GAPDH is associated with a reduced overall survival and relapse-free survival [36]. However, it remains unclear whether the expression of the GAPDH gene is increased by hypoxia in breast cancer

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cells. In this CHAPTER, I clarify the transcription-dependent mechanism of HIF-1 depending on Sp1 in hypoxic breast cancer cells.

MATERIAL AND METHODS

Cell culture. MCF-7 (human breast cancer) cells were cultured in Dulbecco’s

modified Eagle’s medium (DMEM)-high glucose (4.5 g/l glucose) (DMEM-HG) supplemented with 10% fetal bovine serum (FBS) and antibiotics (100 U/ml penicillin and 100 µg/ml streptomycin). SK-BR-3 (human breast cancer), LNCaP (human prostate cancer), PC-3 (human prostate cancer), and DU145 (human prostate cancer) cells were cultured in RPMI 1640 medium supplemented with 10% FBS and the above antibiotics. Cells were maintained at 37ºC in a 5% CO2/95% air atmosphere at 100% humidity

unless otherwise indicated. MCF-7, DU145, and SK-BR-3 cells were obtained form the RIKEN BioResource center (Ibaraki, Japan). LNCaP and PC-3 cells were obtained from the Cell Resource Center for Biomedical Research, Institute of Development, Aging and Cancer, Tohoku University (Miyagi, Japan).

Exposure to hypoxia and chemicals. Cells that had reached approximately 80%

confluence were exposed to hypoxia (1% O2) and to two agents that induce hypoxia-like

response, deferoxamine mesylate (DFX) and CoCl2, for time periods indicated. The

medium was renewed 12 h before exposure to hypoxia, DFX, or CoCl2. DFX and CoCl2

were prepared in dimethyl sulfoxide and sterile water, respectively.

Semi-quantitative reverse transcription (RT)-PCR. MCF-7 cells were exposed

to hypoxia, and total RNA was extracted and reverse transcribed. The resultant cDNAs were subjected to semi-quantitative PCR using specific primers for GAPDH (hG3PDH- F and hG3PDH-R) and β-actin (h-Beta-actin-F and h-Beta-actin-R). PCR was performed under the following conditions: denaturation at 95ºC for 1 min, primer-annealing at 55ºC for 1 min, and primer extension at 72ºC for 30 sec. The final extension was performed at 72ºC for 10 min. PCR products were subjected to agarose

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gel electrophoresis and stained with ethidium bromide. The relative expression level of mRNAs were calculated by determining the ratio of the amount of each mRNA to that of endogenous reference gene, β-actin.

Western blot analysis. For detection of endogenous GAPDH and HIF-1α,

MCF-7 and LNCaP cells were exposed to hypoxia for 24 h. Cells were harvested and sonicated in HN buffer (20 mM Hepes-NaOH, pH 7.5, 150 mM NaCl, 0.5% (w/v) Nonidet P-40, 1mM 4-(2-aminoethyl)-benzensulfonyl fluoride (AEBSF), 10 µg/mL leupeptin, and 1 µg/mL aprotinin), followed by centrifugation at 20,000×g for 20 min. For detection of endogenous Sp1, MCF-7 and LNCaP cells were exposed to hypoxia for various time periods. Cells were harvested and resuspended in TNE buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 0.5% Nonidet P-40, 10 mM sodium pyrophosphate, 2 mM EDTA, 10 mM sodium pyrophosphate, 1 mM AEBSF, 10 µg/ml leupeptin, and 1 µg/ml aprotinin), and incubated for 30 min on ice, followed by centrifugation at 20,000×g for 20 min. The supernatants were subjected to SDS-PAGE and analyzed by Western blotting with rabbit polyclonal anti-GAPDH [34] and anti-Sp1 (PEP 2, Santa Cruz Biotechnology, Santa Cruz, CA, USA) antibodies and mouse monoclonal anti-HIF-1α (Clone mgc3, Affinity Bioreagents, Golden, CO, USA) and anti-α-tubulin (clone DM 1A, Sigma Aldrich, Saint Louis, MO, USA) antibodies, followed by immunoreaction with the horseradish peroxidase-conjugated goat anti-rabbit and anti-mouse IgG, respectively. The immunoreactive proteins were visualized with Immobilon Western Chemiluminescent substrate (Millipore Corporation, Billerica, MA, USA).

Subcellular fractionation. MCF-7 cells cultured in 100-mm dishes were exposed

to hypoxia for 24 h. Cells were harvested and homogenized repeated passage in HS buffer (20 mM Hepes-NaOH, pH 7.5, 250 mM Sucrose, 1 mM EDTA, 1 mM AEBSF, 10 µg/ml leupeptin, and 1 µg/ml aprotinin) using microtube homogenizer (PT-α, I.S.O. Inc., Kanagawa, Japan). The homogenate was centrifuged at 270×g for 3 min and the supernatant was transfer to a fresh tube. The precipitation was resuspended in HS buffer,

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homogenized with the homogenizer and centrifuged at 270×g for 3 min. The supernatant obtained was mixed with the first supernatant and centrifuged at 750×g for 10 min to precipitate the nucleus. The supernatant was referred to as the post-nucleus fraction. Nucleus were resuspended and sonicated in HS buffer, followed by centrifugation at 20,000×g for 20 min. The supernatant is referred to as the nuclear fraction. Proteins in each fraction were subjected to SDS-PAGE and analyzed by Western blotting with rabbit polyclonal anti-GAPDH and anti-TPI [34] antibodies and mouse monoclonal anti-lamin B1 antibody (L-5, Zymed Laboratories Inc., San Francisco, CA, USA).

Preparation of nuclear extract. MCF-7 and LNCaP cells cultured in 100-mm

dishes were exposed to hypoxia for 3 h. Cells were harvested and resuspended in hypotonic buffer (10 mM Hepes-NaOH, pH 7.5, 1.5 mM MgCl2, 10 mM KCl, 1 mM

AEBSF 10 µg/ml leupeptin, and 1 µg/ml aprotinin), follwed by incubation for 30 min on ice. The cell suspension was centrifuged at 20,000×g for 30 sec at 4ºC. The precipitations was resuspended in hypertonic buffer (20 mM Hepes-NaOH, pH 7.5, 420 mM NaCl, 1.5 mM MgCl2, 0.2 mM EDTA, 25% glycerol, 0.5 mM DTT, 1 mM AEBSF,

10 µg/ml leupeptin, and 1 µg/ml aprotinin), followed by incubation for 30 min on ice. The cell suspension was centrifuged at 20,000×g for 2 min at 4ºC. The supernatant was referred to as the nuclear extract. The extract was subjected to SDS-PAGE and analyzed by Western blotting with anti-Sp1 antibody.

Construction of plasmid vectors. The six potential consensus HREs in human

GAPDH gene (–1091 to +25) were termed HRE1, HRE2, HRE3, HRE4, HRE5, and HRE6 from upstream to downstream (Fig. 3A). The promoter region of human GAPDH gene (GenBank Accession No. J04038) was amplified by two-sequential nested PCR. The first PCR was performed using sense primer (-1515--1492) and antisense primer (658-635), and GenomeWalker Human Kit Dra I (Clontech Laboratories, San Jose, CA, USA) as a template. The second PCR was performed using sense primer (KpnI/-1091-- 1067) and antisense primer (BglII/25-2), and the first PCR products as a template,

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followed by inserting into pCR2.1 TOPO-TA vector (Invitrogen, Carlsbad, CA, USA), termed pCR2.1-GAP(–1091). Human GAPDH gene (–230 to +25) was amplified by PCR. The PCR was performed using sense primer (NheIS-230/-207) and antisense primer (HindIIIAS+25/+2), and the EcoR I-digested fragment of pCR2.1-GAP(–1091) as a template, followed by inserting into pCR2.1 TOPO-TA vector, termed pCR2.1- GAP(–230). The Kpn I- and Bgl II-digested fragment of pCR2.1-GAP(–1091) and the

Nhe I- and Hind III-digested fragment of pCR2.1-GAP(–230) were subcloned into the

corresponding sites of pGL3-basic vector (Promega Corp., Madison, WI, USA), termed pGAP(–1091) and pGAP(–230), respectively. The mutation into HRE4 or HRE5 between –230 to +25 in the human GAPDH gene was introduced by PCR using sense primer (NheImwS-230/-192) and antisense primer (HindIIIAS+25/+2) for mutation into HRE4 or sense primer (NheIwmS-230/-181) and antisense primer (HindIIIAS+25/+2) for mutation into HRE5, and EcoR I-digested fragment of pCR2.1-GAP(–230) as a template, followed by insertion into pCR2.1 TOPO-TA vector, termed pCR2.1-GAP-5m and pCR2.1-GAP-5m, respectively. The mutation into HRE6 between –230 to +25 in the human GAPDH gene was introduced by two-step PCR using sense primer (NheIS -230/-207) and antisense primer (mAS-102AAAG-147) for amplification of 5´-terminus region or sense primer (mS-145CTTT-100) and antisense primer (HindIIIAS+25/+2) for amplification of 3´-terminus region, and EcoR I-digested fragment of pCR2.1-GAP (–1091) as a template. Each reaction products were mixed, and PCR was performed using (NheIS-230/-207) and (HindIIIAS+25/+2), followed by insertion into pCR2.1 TOPO-TA vector, termed pCR2.1-GAP-6m. The Nhe I- and Hind III-digested fragments of pCR2.1-GAP-4m, pCR2.1-GAP-5m, and pCR2.1-GAP-6m were subcloned into the corresponding sites of pGL3-basic vector, termed pGAP-4m, pGAP-5m, and pGAP-6m, respectively. The mutation into HRE1 between –1091 to +25 in the human GAPDH gene was introduced by two-step PCR using sense primer (KpnI/-1091--1067) and antisense primer (AS-Site1m) for amplification of 5´-terminus region or sense primer (S-Site1m) and antisense primer (BglII/25-2) for amplification of 3´-terminus region, and EcoR I-digested fragment of pCR2.1-GAP(–1091) as a template. Each reaction products were mixed, and PCR was performed using sense primer (KpnI/-1091--1067)

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and antisense primer (BglII/25-2), followed by insertion into pCR2.1 TOPO-TA vector, termed pCR2.1-GAP-1m. The mutation into HRE2 between –1091 to +25 in the human GAPDH gene was introduced by two-step PCR using sense primer (KpnI/-1091--1067) and antisense primer (AS-Site2m) for amplification of 5´-terminus region or sense primer (S-Site2m) and antisense primer (BglII/25-2) for amplification of 3´-terminus region, and EcoR I-digested fragment of pCR2.1-GAP-1m as a template. Each reaction products were mixed, and PCR was performed using sense primer (KpnI/-1091--1067) and antisense primer (BglII/25-2), followed by insertion into pCR2.1 TOPO-TA vector, termed pCR2.1-GAP-(1+2)m. The mutation into GC-box between –1091 to +25 in the human GAPDH gene was introduced by two-step PCR using sense primer (KpnI/-1091 --1067) and antisense primer (GAP-SP1m-AS) for amplification of 5´-terminus region or sense primer (GAP-SP1m-S) and antisense primer (BglII/25-2) for amplification of 3´-terminus region, and EcoR I-digested fragment of pCR2.1-GAP(–1091) as a template. Each reaction products were mixed, and PCR was performed using sense primer (KpnI/- 1091--1067) and antisense primer (BglII/25-2), followed by insertion into pCR2.1 TOPO-TA vector, termed pCR2.1-GAP-GCm. The Kpn I- and Bgl II-digested fragments of pCR2.1-GAP-1m, pCR2.1-GAP-(1+2)m, and pCR2.1-GAP-GCm were subcloned into the corresponding sites of pGL3-basic vector, termed pGAP-1m, pGAP-(1+2)m, and pGAP-GCm, respectively. Mammalian expression vector pcDNA3.1-HA-Myc-His was constructed by insertion of annealing a set of oligonucleotides (HA-S and HA-AS) encoding HA-tag into the Nhe I and Xho I sites of pcDNA3.1-Myc-His (-) (Invitorogen) vector. Human HIF-1α cDNA (GenBank Accession No. NM_001530) was amplified by two-sequential nested PCR. The first PCR was performed using sense primer (HIF1a (S188-209)) and antisense primer (HIF1a(AS2826-2802)), and human brain (cerebral cortex) Marathon-ready cDNA (Clontech Laboratories) as a template. The second PCR was performed using sense primer (HIF1a-Not/S285-307) and antisense primer (HIF1a Bam/AS2762-2739), and the first PCR product as a template, followed by insertion into pCR2.1 TOPO-TA vector, termed pCR2.1-HIF-1α. The Not I- and BamH I-digested fragment of pCR2.1-HIF-1α was subcloned into the corresponding sites of pcDNA3.1- HA-Myc-His vector, termed pcDNA3.1-HA-HIF-1α-Myc-His. The HIF-1αSM cDNA

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encoding mutant HIF-1α substituting Ala for Pro at position of 564 was synthesized by site-directed mutagenesis of pcDNA3.1-HA-HIF-1α-Myc-His using sense primer (HIF1aP564A/F) and antisense primer (HIF1aP564A/R), and QuikChange II XL Site-directed Mutagenesis kit (Stratagene, La Jolla, CA, USA), termed pcDNA3.1-HA- HIF-1αSM-Myc-His. The HIF-1αDM cDNA encoding mutant HIF-1α substituting Ala for Pro at positions of 402 and 564 was synthesized by site-directed mutagenesis of pcDNA3.1-HA-HIF-1αSM-Myc-His using sense primer (HIF1aP402A/F) and antisense primer (HIF1aP402A/R, and QuikChange II XL Site-directed Mutagenesis kit, termed pcDNA3.1-HA-HIF-1αDM-Myc-His. The resulting vectors encode mutants HIF-1α with the N-terminal HA-tag and C-terminal Myc- and His-tags.

Promoter analysis. Cells were cultured in appropriate medium supplemented

with 10% FBS on 24-well plates, and transiently transfected with reporter vector using HilyMax (Dojindo, Kumamoto, Japan) for 24 h. Cells were incubated in fresh medium for 12 h, followed by incubation for an additional 12 h under hypoxic conditions. For measurement of HIF-1α-mediated induction of GAPDH promoter, MCF-7 and LNCaP cells were transiently co-transfected with reporter vector and HIF-1αDM expression vector (pcDNA3.1-HA-HIF-1αDM-Myc-His) for 24 h. HIF-1αDM is a mutant form of HIF-1α, which is stably expressed even in normoxia. Cells were incubated in fresh medium for an additional 24 h. The total amount of plasmid vectors were kept constant by addition of empty vector. Cells were harvested and lysed, and firefly and Renilla luciferase activities were determined with the Dual-Luciferase reporter assay kit and GloMax 20/20 Luminometer (Promega Corp.). Transfection efficiency was normalized with Renilla luciferase expression vector, and data was expressed as relative light units (RLU, firefly luciferase divided by Renilla luciferase).

siRNA-mediated knockdown. Target sequence for siRNA duplexes were as

follows: siHIF-1α #1: 5´-CUGAUGACCAGCAACUUGAdTdT-3´ (Dharmacon, Chicago, IL, USA), siHIF-1α #2: 5´-GCCACUUCGAAGUAGUGCUdTdT-3´ (Sigma Aldrich), and siSp1: 5´-AUCACUCCAUGGAUGAAAUGAdTdT-3´ (Sigma Aldrich).

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The RISC-Free siRNA #1 (D-001220-01, Dharmacon) was used as a control siRNA. The duplexes (10 nM) were introduced into MCF-7 and LNCaP cells using Lipofectamine RNAiMAX reagent (Invitrogen) and Opti-MEM (Invitrogen) for 24 h according to the manufacturer’s protocol.

Chromatin immunoprecipitation (ChIP) analysis. MCF-7 and LNCaP cells

were exposed to hypoxia for 12 h, followed by incubation with 1% paraformaldehyde for 15 min. Cells were rinsed with ice-cold PBS and lysed in 150 µl of SDS-lysis buffer (50 mM Tris-HCl, pH 8.0, 1% SDS, 10 mM EDTA, 1 mM AEBSF, 10 µg/ml leupeptin, and 1 µg/ml aprotinin). Cell lysates were sonicated (Handy Sonic, Tomy Seiko, Tokyo, Japan), followed by centrifugation at 20,000×g for 3 min. The supernatant was diluted 10-fold with ChIP dilution buffer (16.7 mM Tris-HCl, pH 8.0, 0.01% SDS, 1.1% Triton X-100, 1.2 mM EDTA, 167 mM NaCl, 1 mM AEBSF, 10 µg/ml leupeptin, and 1 µg/ml aprotinin) and precleared with 40 µl of protein G-Sepharose (GE Healthcare, Piscataway, NJ) (50% slurry) for 1 h at 4ºC. The supernatant was reacted with rabbit polyclonal anti-Sp1 IgG, mouse monoclonal anti-HIF-1α IgG (Clone H1a67, Novus Biologicals, Littleton, CO, USA) or appropriate control IgG overnight at 4ºC and incubated with 40 µl of protein G-Sepharose (50% slurry) for 1 h. The resin was sequentially washed with low salt buffer (20 mM Tris-HCl, pH 8.0, 0.1% SDS, 1% Triton X-100, 2 mM EDTA, and 150 mM NaCl), high salt buffer (20 mM Tris-HCl, pH 8.0, 0.1% SDS, 1% Triton X-100, 2 mM EDTA, and 500 mM NaCl), LiCl buffer (10 mM Tris-HCl, pH 8.0, 0.25 M LiCl, 1% Nonidet P-40, 1% deoxycholate, and 1 mM EDTA), and TE buffer (10 mM Tris-HCl, pH 8.0, and 1 mM EDTA). Immunoprecipitated protein complexes were eluted with elution buffer (0.1 M NaHCO3, 10 mM DTT, and 1% SDS) and heated for 6

h at 65ºC to reverse the paraformaldehyde cross-linking. DNA fragments were treated with RNase A and proteinase K, extracted with phenol-chloroform, and precipitated by ethanol. The promoter region of GAPDH gene was amplified by PCR. Primer sequences were P1, forward (GAP-ChIP-F) and reverse (GAP-ChIP-R); P2, forward (GAP-ChIP2-F) and reverse (GAP-ChIP3-R).

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RESULTS

Hypoxic up-regulation of GAPDH gene expression.

To study the effect of hypoxia on GAPDH gene expression, MCF-7 cells were exposed to hypoxia, and mRNA was prepared. RT-PCR analysis showed that hypoxia resulted in increased GAPDH mRNA levels (Fig. 3A). The GAPDH mRNA expression was increased progressively as a function of time exposed to hypoxia. Furthermore, when cells were exposed to hypoxia for 24 h, hypoxia caused a 2.5-fold increase in the protein level of GAPDH (Fig. 3B). To determine the intracellular distribution of hypoxia-induced GAPDH, cells were exposed to hypoxia for 24 h. Subcellular fractionation was performed by differential centrifugation. The protein levels of GAPDH in the nuclear fractions of hypoxic cells were increased compared with those in normoxic cells (Fig. 3C). However, the GAPDH expression levels in the cytosolic and particulates fractions were unaffected by hypoxia. TPI, a cytosolic marker protein, and lamin B1, a nuclear marker protein, were detected only in each corresponding fraction. These results indicate that hypoxia up-regulates the expression levels of GAPDH mRNA and protein in MCF-7 cells.

Transcriptional activation of the GAPDH gene by hypoxia.

The expression of GAPDH is regulated by hypoxia in a cell type-specific manner [31]. The GAPDH promoter region (–1091 to +25) contains six potential consensus HREs (–989 to –985, –957 to –953, –340 to –336, –217 to –213, –207 to –203, and –125 to –121), termed HRE1–6, respectively (Fig. 4A). To identify the functional HREs of GAPDH gene in MCF-7 cells, I constructed two luciferase reporter vectors, one composed of the nucleotide sequence from –1091 to +25 containing six HREs, termed pGAP(–1091), and the other composed of the nucleotide sequence from –230 to +25 containing three HREs (HRE4–6), termed pGAP(–230). Breast cancer MCF-7 and SK-BR-3 cells were transiently transfected with either pGAP(–1091) or pGAP(–230) reporter vector and exposed to hypoxia. Hypoxia induced luciferase activity in both cell lines, and the fold induction of luciferase activity in the cells transfected with

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pGAP(–230) was less than half that in the cells transfected with pGAP(–1091) (Fig. 4B, upper panel). Likewise, hypoxia enhanced the fold induction of luciferase activity when LNCaP and DU145 cells were transiently transfected with pGAP(–1091) and exposed to hypoxia (Fig. 4B, lower panel). In PC-3 cells, the fold induction of luciferase activity

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marginally increased by hypoxia. However, deletion of the region from –1091 to –231 had no effect on the fold induction of luciferase activity by hypoxia, consistent with previous report in LNCaP cells [39]. On the other hand, when MCF-7 cells were exposed to the hypoxia mimetics DFX and CoCl2, luciferase activity was induced (Fig.

4C). The fold induction of luciferase activity by DFX and CoCl2 in the cells transfected

with pGAP(–230) was only half that in the cells transfected with pGAP(–1091). Thus, the nucleotide sequence from –1091 to –231 of the GAPDH gene in breast cancer cells has hypoxia-responsive regions, indicating that the mechanisms by which hypoxia increases GAPDH promoter activity are different between breast cancer MCF-7 cells and prostate cancer LNCaP cells.

Identification of functional HREs in the GAPDH promoter.

The nucleotide sequence from –1091 to –231 of the GAPDH promoter contains three potential consensus HREs (HRE1–3). To determine which HREs are required for the hypoxic induction of the GAPDH gene in MCF-7 cells, mutations were introduced into HRE1 alone and both HRE1 and HRE2 of the pGAP(–1091) reporter vector (Fig. 5A). The resultant mutated luciferase reporter vectors were termed pGAP-1m and pGAP-(1+2)m. Cells were transfected with the reporter vector, followed by exposure to hypoxia. In MCF-7 cells transfected with either mutated reporter vector, the fold induction of luciferase activity by hypoxia was reduced to that in cells transfected with pGAP(–230) (Fig. 5B, upper panel). In contrast, in LNCaP cells, neither mutation affected the fold induction of luciferase activity by hypoxia (Fig. 5B, lower panel). These results indicate that HRE1 is functional in MCF-7 cells, but not in LNCaP cells. On the other hand, the nucleotide sequence from –230 to +25 of the GAPDH promoter has three potential consensus HREs (HRE4–6). To identify which HREs are functional in hypoxic MCF-7 cells, mutations were introduced into each of three HREs (HRE4–6), and mutated reporter vectors (pGAP-4m, pGAP-5m, and pGAP-6m) were constructed (Fig. 5A). Hypoxia did not affect the fold induction of luciferase activity in MCF-7 and LNCaP cells transfected with pGAP-6m, indicating that HRE6, but not HRE4 and HRE5, is functional in both breast and prostate cancer cells (Fig. 5C).

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HIF-1α-mediated enhancement of GAPDH gene expression.

Hypoxia induced HIF-1α protein expression in both MCF-7 cells and LNCaP cells (Fig. 6A). To determine whether HIF-1α is involved in hypoxia-increased GAPDH promoter activity, MCF-7 and LNCaP cells were transfected with two types of HIF-1α-specific siRNAs and exposed to hypoxia. Western blot analyses showed that HIF-1α siRNAs specifically knocked down hypoxia-induced endogenous HIF-1α protein, but not α-tubulin (Fig. 6B). siRNA-mediated knockdown of HIF-1α resulted in a complete loss of hypoxia-induced luciferase activity in both cells transfected with pGAP(–1091) (Fig. 6C). Furthermore, to examine the involvement of HIF-1α in hypoxic induction of GAPDH gene expression, HIF-1αDM expression plasmid and reporter plasmid were transiently co-transfected into MCF-7 and LNCaP cells. HIF-1αDM, a mutant form of HIF-1α, can be expressed even in normoxia [44]. Exogenous HIF-1αDM increased luciferase activity in both cell lines (Fig. 6D). In contrast, the deletion of nucleotides –1091 to –231 resulted in reduction of the HIF-1αDM-increased lucifease activity in MCF-7 cells, but not in LNCaP cells. These results suggest that in MCF-7 cells, HIF-1α acts on HRE1, which was identified as a functional HRE within the nucleotide sequence from –1091 to –231. Therefore, to assess whether HIF-1 binds to HRE1, ChIP analyses with anti-HIF-1α antibody were performed. Two sets of PCR primers were used and illustrated in Fig. 7A. MCF-7 and LNCaP cells were exposed to hypoxia, followed by preparation of soluble chromatin. ChIP analysis using the primer set P1 revealed that HIF-1α bound to HRE1 in both MCF-7 and LNCaP cells under hypoxic conditions (Fig. 7B, upper panel). However, when the primer set P2 was used, no HIF-1α binding was detected in both cell lines (Fig. 7B, lower panel). These results indicate that HIF-1 binds to HRE1 in hypoxia even though it may not enhance transcription of GAPDH gene.

Participation of Sp1 in hypoxic up-regulation of GAPDH gene.

A computer analysis of the region flanking HRE1 revealed the presence of a GC-box (5´-GGGCGG-3´, nucleotide sequence between –983 and –978), which is a putative binding site of Sp family proteins [45]. To determine whether the GC-box

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affects HRE1 function in hypoxic MCF-7 cells, the nucleotide sequence of the GC-box was mutated, and mutated pGAP-GCm reporter vector was constructed (Fig. 8A). Cells were transfected with the reporter vector and exposed to hypoxia. Mutation of the GC-box reduced the fold induction of luciferase activity by hypoxia in MCF-7 cells, but not in LNCaP cells (Fig. 8B). These results indicate that the GC-box adjacent to HRE1 is essential for HRE1 to act as a functional HRE in MCF-7 cells. To assess the expression levels of Sp1 protein in MCF-7 and LNCaP cells, whole cell lysates and nuclear extracts were subjected to SDS-PAGE, followed by Western blot analysis with anti-Sp1 antibody. As shown in Fig. 9A, total and nuclear Sp1 proteins in MCF-7 cells were higher than those in LNCaP cells. When the effect of hypoxia on Sp1 expression was determined, hypoxia increased the Sp1 protein level in MCF-7 cells, but not in

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LNCaP cells (Fig. 9B). To determine whether the endogenous Sp1 binds to the GC-box adjacent to HRE1, MCF-7 and LNCaP cells were exposed to hypoxia. ChIP analysis with anti-Sp1 antibody revealed that Sp1 bound to the GC-box only in MCF-7 cells and that the Sp1 level in normoxia was the same as that in hypoxia (Fig. 10). Furthermore, the effects of knockdown of Sp1 on the hypoxia-increased luciferase activity were determined. Western blot analysis showed that Sp1 siRNA specifically decreased the

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endogenous Sp1 protein level, but not the α-tubulin level (Fig. 11A). Sp1 knockdown reduced the fold induction of luciferase activity by hypoxia in MCF-7 cells, but not in LNCaP cells (Fig. 11B). Furthermore, the increase in hypoxia-induced GAPDH protein level was inhibited by Sp1 siRNA in MCF-7 cells, but not in LNCaP cells, whereas Sp1 siRNA had no influence on GAPDH protein level in normoxia (Fig. 11C). These results

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indicate that Sp1 is essential for hypoxic up-regulation of the GAPDH gene in MCF-7 cells but that Sp1 does not affect GAPDH expression in normoxia.

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DISCUSSION

Hypoxia up-regulates the transcriptional activation of GAPDH in a cell type-specific manner [31]. Therefore, uncovering the regulatory mechanism of GAPDH gene under hypoxic conditions may lead to elucidate the novel mechanism of HIF transactivation. In this CHAPTER, I found that hypoxia increased GAPDH gene expression in a manner specific to breast cancer MCF-7 cells.

Deletion analysis of the GAPDH promoter revealed that breast cancer-specific hypoxic induction of the GAPDH promoter activity depended on the nucleotide sequence from –1091 to –231. Furthermore, introduction of mutations into the potential HREs of the GAPDH promoter showed that HRE1 (–989 to –985) was functional in addition to HRE6 (–125 to –121) in MCF-7 cells and that HRE6 alone was functional in LNCaP cells. Knockdown analysis of HIF-1α by siRNA indicated that HIF-1 contributed to up-regulation of GAPDH gene expression by hypoxia in both MCF-7 and LNCaP cells. Graven et al. [43] reported that HRE6 alone is involved in hypoxic up-regulation of the GAPDH gene in bovine aortic and pulmonary artery endothelial cells, human lung microvascular endothelial cells, and human hepatoma Hep3B cells [40]. HIF-2, but not HIF-1, preferentially binds to HRE6 in human lung microvascular endothelial cells, while both HIF-1 and HIF-2 bind to HRE6 in Hep3B cells. Thus, cell type-specific expression patterns of HIF-1α and HIF-2α seem to lead to cell type-specific regulation of GAPDH gene expression under hypoxic conditions. On the other hand, Lu et al. [39] reported that not only HRE6, but also HRE4 (–217 to –213) and HRE5 (–207 to –203), are functional in LNCaP cells, although it remains unclear which HIF isoforms are active on these HREs. These results are inconsistent with the present results that HRE6 alone was functional in LNCaP cells. However, in order for hypoxia to enhance GAPDH promoter activity, it appears that at least HRE6 needs to be functional.

Endogenous HIF-1α bound to HRE1 in both MCF-7 and LNCaP cell lines under hypoxic conditions. The GC-rich sequence to which Sp1 binds was found to be located in the 3´-flanking region of HRE1. Mutation within the GC-box resulted in a reduced

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HIF-1 activity in MCF-7 cells, but not in LNCaP cells, indicating that the GC-box is required for HRE1 to elicit HIF-1 activity in MCF-7 cells. ChIP analysis showed that endogenous Sp1, which is a ubiquitous nuclear transcriptional factor, was located on the GC-box in MCF-7 cells in normoxia and hypoxia, but not in LNCaP cells even in hypoxia. siRNA-mediated knockdown of Sp1 resulted in a reduced promoter activity and expression level of GAPDH only in hypoxic MCF-7 cells. These results indicate that the binding of Sp1 to the GC-box is not required for the binding of HIF-1 to HRE1 but that it is required for elicitation of HIF-1 activity on HRE1. In contrast, in normoxia, knockdown of Sp1 did not affect the GAPDH protein level, indicating that Sp1 itself has no effect on the regulation of GAPDH expression in normoxic cells. The mechanisms by which Sp1 binds to the GC-box may be regulated by the amounts of nuclear Sp1 or by certain factors that recruit Sp1 to the GC-box. Sp1 was increased by hypoxia in MCF-7 cells, but not in LNCaP cells, in agreement with the finding that the expression of Sp1 is induced by hypoxia in a cell type-specific manner [46-48]. However, the Sp1 level on the GC-box in normoxia was the same as that in hypoxia, indicating that the amount of Sp1 in normoxic MCF-7 cells is sufficient for HIF-1 to function on HRE1.

HIF-1 and Sp1 contributed to the hypoxic up-regulation of GAPDH gene in MCF-7 cells, but not LNCaP cells. The mechanisms by which hypoxia induces the activation of carbonic anhydrase IX and retinoic acid receptor-related orphan receptor α4 genes depend on HRE and its adjacent GC-box, and deletion of either HRE or GC-box results in loss of hypoxia-induced activation [49, 50]. On the other hand, Sp1 up-regulates the expression of β-enolase, pyruvate kinase-M, and ephrinB2 through a HIF-1-independent mechanism [48, 51] that is different from the regulatory mechanism for GAPDH gene expression in hypoxia. Thus, Sp1 play a critical role in the hypoxic up-regulation of hypoxia-inducible genes. Therefore, critical regions in the promoter regions of the hypoxia-inducible genes include not only potential HREs, but also GC-boxes, especially the GC-box adjacent to HRE.

In breast cancer MCF-7 cells, hypoxia resulted in up-regulation of GAPDH mRNA and protein, and the GAPDH expression is significantly induced in nuclear

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fraction. Recent studies show that the overall survival and relapse-free survival rates are lower in breast cancer patients in which the expression of GAPDH is increased, indicating that GAPDH expression is associated with the proliferation and aggressiveness of breast carcinoma cells [36]. GAPDH possesses diverse functions independent of its role in glycolysis and participates in many physiological and pathophysiological processes [52]. In nuclear, GAPDH involves in tRNA transport, apoptosis, DNA repair, DNA replication, and androgen receptor transactivation [52, 53]. These functions independent of glycolysis may contribute to the cancer progression.

In this CHAPTER, I clarified the novel transcription-dependent mechanism of HIF-1 in breast cancer. HIF function is considered to be one of the molecular target to cancer therapy. However, because HIF signaling is very complicated and is involved in many biological activities, most of HIF inhibitors are the lack of specificity. Therefore, repression of this breast cancer-specific HIF signaling may be an effective strategy for breast cancer treatment.

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CHAPTER II

A Variant Form Lacking Exons 12 to 15 of Human Hypoxia-inducible

Factor-2

α Functions as a Dominant Negative Form

Solid tumors have unique hypoxic microenvironments, and tumor growth in hypoxia is largely regulated by HIF-mediated gene expression. HIF is composed of α and β subunits. Two types of α-subunit isoforms, HIF-1α and HIF-2α, share a common modular structure. However, HIF-1α and HIF-2α are expressed in distinct temporal and spatial patterns, and do not function redundantly [7, 20, 54]. Thus the expression of hypoxia-responsive genes depends mainly on the amount and isoform of the α-subunit that is expressed in response to hypoxia.

Inhibitors of HIF transcriptional activity might be useful as therapeutic tools for hypoxia-related diseases such as cancers [28]. Recently, alternatively spliced isoforms of HIF-1α have been reported [55, 56]. Two of six HIF-1α isoforms, including wild-type HIF-1α function as dominant negative isoforms of HIF-1α. On the other hand, no alternatively spliced variants of HIF-2α have been found.

In this CHAPTER, to provide a novel therapeutic tool for cancer, I explored the existence of transcript variants of HIF-2α. The variant lacking exons 12 to 15 of HIF-2α mRNA is predicted to contain the bHLH/PAS, the N-terminal nuclear localization signal, and the N-terminal part of N-TAD in the ODD domain. Because these structural features suggest that the variant form can function as a dominant negative isoform, I constructed and characterized the variant.

Cell culture. HepG2 (human hepatoma) and HEK293 (human embryonic kidney)

cells were cultured in DMEM supplemented with 10% FBS and antibiotics (100 U/ml penicillin and 100 µg/ml streptomycin). Cells were maintained at 37ºC in a 5% CO2/95% air atmosphere at 100% humidity unless otherwise indicated. HepG2 and

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HEK293 cells were obtained form the RIKEN BioResource center.

Primers and probes. The nucleotides HIF2a/S472-495 and HIF2a/AS3036-3011

were labeled fluorescein-11-dUTP with Gene Images 3’-oligolabelling module (Amersham Pharmacia Biotech.) according to the manufacture’s protocols. The labeled nucleotides were used as probes A and B, respectively, in this study.

Southern blot analysis. The total RNA was extracted from HepG2 cells, and was

reverse-transcribed to synthesize cDNAs. The resultant cDNAs were subjected to two-sequential nested PCRs. For amplification of the region between exons 1 and 9, the first PCR was performed using sense primer (HIF2a/S201-222) and antisense primer (HIF2a/AS1676-1652). The second PCR was performed using sense primer (HIF2a/ S277-300) and antisense primer (HIF2a/AS1635-1610), and the first PCR product as a template. For amplification of the region between exons 9 and 16, the first PCR was performed using sense primer (HIF2a/S1567-1588) and antisense primer (HIF2a/AS 3412-3391). The second PCR was performed using sense primer (HIF2a/S1612-1637) and antisense primer (HIF2a/AS3174-3149), and the first PCR product as a template. The amplified PCR products were detected by Southern blot analysis using the probe A or probe B, respectively. Primer locations used in this section were summarized in Fig. 12A.

Construction of plasmid vectors. Constructions of pcDNA3.1-HA-HIF-1αDM-

Myc-His were described in CHAPTER I. Human HIF-2α cDNA (GenBank Accession No. NM_001430) was amplified by two-sequential nested PCR. The first PCR was performed using sense primer (HIF2a/S201-222) and antisense primer (HIF2a/AS3467- 3444), and human brain (cerebral cortex) Marathon-ready cDNA as a template. The second PCR was performed using sense primer (HIF2a-NotI/S489-511) and antisense primer (HIF2aBamAS/3098-3076), and the first PCR product as a template, followed by insertion into pCR2.1 TOPO-TA vector, termed pCR2.1-HIF-2α. The Not I- and

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sites of pcDNA3.1-HA-Myc-His and p3xFLAG-Myc-CMV-26 (Sigma Aldrich) vectors, termed pcDNA3.1-HA-HIF-2α-Myc-His and p3xFLAG-HIF-2α, respectively. The resulting vector encodes HIF-2α with the N-terminal HA-tag and C-terminal Myc- and His-tags or the three tandem repeats of N-terminal FLAG-tags, respectively. The HIF-2αSM cDNA encoding mutant HIF-2α substituting Ala for Pro at position of 531 was synthesized by site-directed mutagenesis of pcDNA3.1-HA-HIF-2α-Myc-His using sense primer (HIF2aP531A/F) and antisense primer (HIF2aP531A/R), and QuikChange II XL Site-directed Mutagenesis kit, termed pcDNA3.1-HA-HIF-2αSM-Myc-His. The HIF-2αDM cDNA encoding mutant HIF-2α substituting Ala for Pro at positions of 405 and 531 was synthesized by site-directed mutagenesis of pcDNA3.1-HA-HIF-2αSM- Myc-His using sense primer (HIF2aP405A/F) and antisense primer (HIF2aP405A/R), and QuikChange II XL Site-directed Mutagenesis kit, termed pcDNA3.1-HA-HIF- 2αDM-Myc-His. The resulting vector encodes HIF-2αDM with the N-terminal HA-tag and C-terminal Myc- and His-tags. The Not I- and BamH I-digested fragment of pcDNA3.1-HA-HIF-2αDM-Myc-His was subcloned into the corresponding sites of p3xFLAG-Myc-CMV-26 vector, termed p3xFLAG-HIF-2αDM. The resulting vector encodes HIF-2αDM with the three tandem repeats of N-terminal FLAG-tag. The HIF-2α splicing variant cDNA which is indentified in this study was generated by two-sequential nested PCR. The first PCR was performed using sense primer (HIF2a- NotI/S489-511) and antisense primer (HIF2a/AS2042-2019), and BamH I-digested fragment of pCR2.1-HIF-2α as a template. The second PCR was performed using sense primer (HIF2a-NotI/S489-511) and antisense primer (HIF2aV(ADclone1), and the first PCR product as a template, followed by insertion into pCR2.1 TOPO-TA-vector, termed pCR2.1-HIF-2α(∆E12-15). The Not I- and BamH I-digested fragment of pCR2.1-HIF- 2α(∆E12-15) was subcloned into the corresponding sites of pcDNA3.1-HA-Myc-His and p3xFLAG-Myc-CMV-26, termed pcDNA3.1-HA-HIF-2α(∆E12-15)-Myc-His and p3xFLAG-HIF-2α(∆E12-15), respectively. The resulting vector encodes HIF-2α(∆E12- 15) with the N-terminal HA-tag and C-terminal Myc- and His-tags or the three tandem repeats of N-terminal FLAG-tags, respectively. Luciferase reporter vector, p3xEpoHRE -SV40-Luc, which has three tandem repeats of a consensus HRE was constructed

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according to the methods of Kietzmann et al. [57]. Annealed oligonucleotides containing three tandem repeats of a consensus HRE driven from the human erythropoietin 3´-flanking region, (EPO3xHRE/SacF and EPO3xHRE/SacR), were inserted into the Sac I site of pGL3-promoter vector.

Reporter assay. For measurement of hypoxia-mediated luciferase activity,

HEK293 cells were transiently transfected with pRL-SV40 (0.005 µg), p3xEpoHRE- SV40-Luc (0.1 µg), and pcDNA3.1-HA-HIF-2α(∆E12-15)-Myc-His (0.1 or 0.5 µg) for 24 h, followed by incubation in fresh medium for 12 h. Cells were cultured for an additional 9 h in hypoxia (1% O2), and luciferase activity was determined. The samples

of luciferase reporter assay were separated by SDS-PAGE and subjected to Western blot analysis with anti-c-Myc antibody (A-14). For measurement of HIF-1αDM- and HIF-2αDM-mediated luciferase activity, HEK293 cells were transiently transfected with pRL-SV40 (0.005 µg), p3xEpoHRE-SV40-Luc (0.1 µg), pcDNA3.1-HA-HIF-2α (∆E12-15)-Myc-His (0.1 or 0.5 µg), and either pcDNA3.1-HA-HIF-1αDM-Myc-His or pcDNA3.1-HA-HIF-2αDM-Myc-His (0.1 µg) for 24 h, followed by incubation in fresh medium for 24 h. The total amount of plasmid vectors were kept constant by addition of empty vector. The luciferase activity was determined as described CHAPTER I. Transfection efficiency was normalized with Renilla luciferase expression vector, and data was expressed as relative light units (RLU, firefly luciferase divided by Renilla luciferase).

Immunoprecipitation. HEK293 cells were transiently transfected with p3xFLAG

-HIF-2αDM or p3xFLAG-HIF-2α(∆E12-15) (5 µg) for 6 h, followed by incubation in fresh medium for 42 h. Cells were resuspended in TNE buffer, and incubated for 30 min on ice, followed by centrifugation at 20,000×g for 20 min at 4ºC. The supernatant (500 µg) was incubated with mouse monoclonal anti-HIF-1β IgG (1 µg, BD Bioscience, San Diego, California, USA) or control mouse IgG (1 µg) for 1 h at 4ºC, followed by reaction with 40 µl of protein G-Sepharose (50% slurry) for 4 h at 4ºC. The resin was washed five times with TNE buffer, and proteins bound to the resin were analyzed by

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Western blotting with mouse monoclonal anti-FLAG M2 and anti-HIF-1β antibody, followed by immunoreaction with the horseradish peroxidase-conjugated sheep anti-mouse IgG. The immunoreactive proteins were visualized with the Immobilon Western Chemiluminescent substrate.

Plasmid immunoprecipitation (PIP) assay. HEK293 cells were transiently

transfected with p3xEpoHRE-SV40-Luc (2 µg) and pcDNA3.1-HA-HIF-2αDM-Myc- His or pcDNA3.1-HA-HIF-2α(∆E12-15)-Myc-His (1 µg) for 6 h, followed by incubation in fresh medium for 42 h. Soluble DNA was prepared as described ChIP assay in CHAPTER I, and precleared with 40 µl of protein G-Sepharose (50% slurry) for 1 h at 4ºC. The supernatant was reacted with anti-HA IgG (1 µg, 3F10, Roche Diagnostics, Penzberg, Germany) or control rat IgG (1 µg), overnight at 4ºC and incubated with 40 µl of protein G-Sepharose (50% slurry) for 1 h. Purification of DNA fragments from immunoprecipitated complexes with anti-HA antibody described in ChIP assay in CHAPTER I. The DNA, containing three HREs of p3xEpoHRE-SV40- Luc, was amplified by PCR using a sense primer (RVprimer3) and an antisense primer (GLprimer2).

RESULTS

Identification of transcript variants of human HIF-2α mRNA.

To study the existence of transcript variants of HIF-2α mRNA, total RNA was extracted from human hepatoma HepG2 cells, and was reverse-transcribed to synthesize cDNAs. Because the human HIF-2α gene is composed of 16 exons, the presence of splicing variants of HIF-2α was monitored by RT-PCR analysis between exons 1 and 9 and between exons 9 and 16 (Fig. 12A). The two RT-PCR products covered the full-length open reading frame of HIF-2α mRNA. As shown in Fig. 12B, the RT-PCR products between exons 1 and 9 were detected as a single band by Southern blot analysis, cloned, and sequenced. The nucleotide sequence analysis of cloned DNA was identical to that of the authentic form of HIF-2α mRNA, indicating that there were no

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splicing variants between exons 1 and 9 of HIF-2α mRNA. On the other hand, Southern blot analysis of the RT-PCR products between exons 9 and 16 detected seven bands. The RT-PCR products corresponding to five of the bands were cloned and sequenced. The biggest PCR fragment corresponded to the authentic form of HIF-2α mRNA, and the second-smallest PCR fragment, which was faintly detected, was identified as a variant lacking exons 12 to 15 of HIF-2α mRNA, termed HIF-2α(∆E12-15). These results suggest the occurrence of alternatively spliced HIF-2α mRNA due to deletion of exons 12 to 15 in HepG2 cells.

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Structure of HIF-2α(∆E12-15) cDNA.

The HIF-2α(∆E12-15) was cloned and sequenced as described in above, and its structure is summarized in Fig. 13. In HIF-2α(∆E12-15), the joining of exons 11 and 16 resulted in the generation of a new reading frame for exon 16 (the shaded box in Fig. 13). Hence the deduced amino acid sequence of HIF-2α(∆E12-15) was composed of 518 amino acids encoded by exons 1 to 11 and an additional 18 amino acids, Ala-Trp-Gln-Ala-Gly-Cys-Ser-Gly-Pro-His-Leu-Ser-Pro-Thr-Cys-Cys-Pro-Asn, encoded by exon 16. HIF-2α(∆E12-15) conserved the bHLH/PAS, the N-terminal nuclear localization signal, and the N-terminal part of N-TAD in the ODD domain. However, it has lost the C-terminal part of N-TAD and C-TAD. These structural features suggest that HIF-2α(∆E12-15) can bind to DNA, interact with HIF-1β, and translocate to the nucleus, but cannot induce HIF transcriptional activity.

Inhibitory effect of HIF-2α(∆E12-15) on hypoxic response.

To determine whether HIF-2α(∆E12-15) inhibits endogenous HIF transcriptional activity, HEK293 cells were transiently transfected with pcDNA3.1-HA-HIF-2α(∆E12- 15)-Myc-His and p3xEpoHRE-SV40-Luc, followed by exposure to hypoxia. HIF-2α(∆E12-15) suppressed hypoxia-induced HIF transcriptional activity in a dose-dependent manner (Fig. 14A). In contrast, HIF-2α(∆E12-15) had no HIF transcriptional activity, although HIF-2α(∆E12-15) has the N-terminal part of N-TAD, which plays a dominant role in transactivation of hypoxia-responsive genes [58]. Next, to determine the inhibitory effect of HIF-2α(∆E12-15) on HIF-1α- or HIF-2α-induced HIF transcriptional activity, HIF-1αDM or HIF-2αDM expression vector and p3xEpoHRE-SV40-Luc were transiently co-transfected into HEK293 cells. HIF-1αDM and HIF-2αDM activated HIF transcriptional activity in normoxia, and HIF-2α(∆E12-15) inhibited exogenous HIF-1α- or HIF-2α-activated HIF transcriptional activity in a dose-dependent manner when co-expressed (Fig. 14B). These results indicate that HIF-2α(∆E12-15) functions as a dominant negative form.

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HIF-2α(∆E12-15)•HIF-1β complex on HRE.

To examine the association of HIF-2α(∆E12-15) with HIF-1β in vivo, HIF-2α(∆E12-15) or HIF-2αDM expression vector was transiently transfected into HEK293 cells, and immunoprecipitation analysis with anti-HIF-1β IgG was performed.

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Anti-HIF-1β IgG co-immunoprecipitated HIF-2α(∆E12-15) or HIF-2αDM with HIF-1β, whereas control IgG immunoprecipitated neither HIF-1β, HIF-2α(∆E12-15), nor HIF-2αDM (Fig. 15A). Next, to assess whether HIF-2α(∆E12-15) binds to HREs, HIF-2α(∆E12-15) or HIF-2αDM expression vector was co-transfected with p3xEpoHRE-SV40-Luc, and PIP assay was performed [59]. As shown in Fig. 15B, HIF-2α(∆E12-15) interacted with DNA containing HREs in p3xEpoHRE-SV40-Luc. These results indicate that HIF-2α(∆E12-15) forms a protein complex with HIF-1β on HRE.

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DISCUSSION

Repression of HIFs function is proposed as a strategy for cancer prevention. Recently, alternatively spliced isoforms of HIF-1α have been reported and function as dominant negative isoforms of HIF-1α [55, 56]. On the other hand, no alternatively spliced variants of HIF-2α have been found. In this CHAPTER, I explored transcript variants of HIF-2α to provide a novel tool for cancer therapy. Because the variant form of HIF-2α indentified in this CHAPTER was predicted to function as a dominant negative form, I constructed and characterized the variant form.

Southern blot analysis of the RT-PCR products suggested the occurrence of alternatively spliced HIF-2α mRNA in human hepatoma HepG2 cells. The variant form was indentified as a variant lacking exons 12 to 15, which was termed HIF-2α(∆E12-15). The deduced structure of HIF-2α(∆E12-15) allowed us to predict that HIF-2α(∆E12-15) functions as a dominant negative form of HIF-2α. Reporter analyses showed that HIF-2α(∆E12-15) suppressed hypoxia- and exogenous HIFs-induced luciferase activities, and HIF-2α(∆E12-15) was effective at inhibiting HIF-2 activity at a lower dose, suggesting that HIF-2α(∆E12-15) functions as a dominant negative form, which competes with the α subunits of HIF, especially HIF-2α. Although, HIF-1α and HIF-2α exhibit significant homology in their DNA binding and dimerization domains, HIF-1α and HIF-2α regulate both unique and common target genes [9]. These results suggest that HIF-2α(∆E12-15) are expected to be useful as therapeutic tools for HIF-2α-related diseases, including cancers.

Immunoprecipitation analyses showed that HIF-2α(∆E12-15) formed a protein complex with endogenous HIF-1β. Furthermore, PIP analysis revealed that the complex composed of HIF-2α(∆E12-15) and HIF-1β bound to HREs, indicating that the complex inhibits the binding of authentic HIFs, especially HIF-2, to HREs. In contrast, two HIF-1α dominant negative isoforms, HIF-1αZ [55] and HIF-1α516_{[56], interact}

with HIF-1β in the cytosol and appear to block the nuclear translocation of HIF-1β. Thus the mechanisms by which HIF-2α(∆E12-15) and HIF-1α variants inhibit HIF transcriptional activity might be different. Because HIF-1α and HIF-2α regulate unique

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target gene, the HIF-α subunits do not function redundantly in cancer progression. Therefore, HIF-2α(∆E12-15) is expected to be useful as a therapeutic tool for cancers (e.g., breast cancer and renal cancer) that preferentially express HIF-2α.

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CHAPTER III

The von Hippel-Lindau Protein-dependent Degradation of

Hypoxia-inducible Factor-2

α Regulates Estrogen Receptor α

Expression in Breast Cancer Cells

Estrogens regulate not only physiological processes such as cell growth, development and tissue-specific gene expression in the reproductive tract, brain and cardiovascular tissues, but also pathophysiological processes including breast, endometrial, ovarian and prostate tumorigenesis, osteoporosis and cardiovascular disease [60]. The biological functions of estrogens are exerted through binding to ERα, which is a member of the nuclear receptor superfamily of ligand-mediated transcriptional factors [61]. The nuclear receptors share common structural and functional domains referred to as domains A to E (or F, which is present at the C-terminus in some, but not all nuclear receptors). ERα is composed of domains A to F: the variable N-terminal domain (NTD; also termed the A/B domains), the highly conserved central DNA-binding domain (DBD; also termed the C domain), the non-conserved hinge (D domain), the C-terminal ligand-binding domain (LBD; also termed the E domain), and the F domain. ERα possesses two distinct regions that contribute to transcriptional activity, one of which is a ligand-independent transcriptional activation function (AF-1) located in the N-terminal A/B domain [62]. The other is a ligand-dependent transcriptional activation function (AF-2) located within the LBD (E domain). ERα is localized as an inactivated form in nucleus and cytosol in the absence of ligand, and binding of estrogens to the LBD leads to the transcriptionally active form of ERα (Fig. 16).

The recruitment of coactivators to ERα is required for ERα-mediated transcriptional activity, and a number of ERα coactivators have been indentified and characterized [63-70]. Although the differential regulation of coactivator activity modulates ERα transactivation via direct or indirect interactions, the magnitude of estrogen-induced response depends on the initial levels of ERα [71]. Thus, ERα

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expression is a molecular predictor of tumor responsiveness to antiestrogens during breast cancer therapy. ERα is degraded in response to various stimuli such as estrogen, hypoxia, and dioxin [71-73]. Recently, it has been reported that overexpressed HIF-1α is involved in the proteasome-dependent ERα degradation under hypoxic mimetic conditions in breast cancer MCF-7 cells [74]. In contrast, increased HIF-1α levels in mammary tumors correlate with increased ERα expression [75]. Thus, the detailed regulation of ERα expression by HIF-1α remains unclear.

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Under hypoxia conditions, stabilized HIF-α subunits bind with HIF-β subunit and function as transcription factors in nucleus. In contrast, in normoxia, HIF-α subunits are unable to function as a transcription factors due to the proteasome-dependent degradation controlled by PHDs and VHL. Therefore, to indentify transcription- independent functions of HIF, I focus on the effect of HIF-α subunits on ERα expression mainly in normoxia. In this CHAPTER, I report that siRNA-mediated knockdown of HIF-2α, but not of HIF-1α, up-regulates the ERα expression level in breast cancer cells. Furthermore, I demonstrate that HIF-2α forms a protein complex with ERα in the presence and absence of E2.

Cell culture. MCF-7 (ERα-positive human breast cancer) and SK-BR-3

(ERα-negative human breast cancer) cells were cultured as described in CHAPTER I. COS-7 (African green monkey kidney) cells were cultured in RPMI 1640 medium supplemented with 10% FBS and antibiotics (100 U/ml penicillin and 100 µg/ml streptomycin). Cells were maintained at 37ºC in a 5% CO2/95% air atmosphere at 100%

humidity unless otherwise indicated. COS-7 cells were obtained from American Type Culture Collection (Manassas, VA, USA).

Construction of plasmid vectors. Mammalian expression vector, pcDNA3.1-

Myc-Myc-His, was constructed by insertion of annealing a set of oligonucleotides (F- NheI-Myc-XhoI and R-NheI-Myc-XhoI)) encoding Myc-tag into the Nhe I and Xho I sites of pcDNA3.1-Myc-His (-) vector. Escherichia coli (E. coli) expression vector, pET30a(+)-Myc, was constructed by insertion of annealing a set of oligonucleotides (myc-F/NspV and myc-R/KpnI) encoding Myc-tag into the Nsp V and Kpn I sites of pET30a(+) (Novagen, Madison, WI, USA) vector. Constructions of pcDNA3.1-HA-HIF -2αDM-Myc-His, p3xFLAG-HIF-2α, and p3xFLAG-HIF-2αDM were described in CHAPTER II. The HIF-2αDM(∆N-TAD) cDNA encoding mutant HIF-2αDM lacking N-TAD (amino acids 496-545) was amplified by two-step PCR, using sense primer

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(HIF2a-NotI/S489-511) and antisense primer (AS2a-delNTAD) for amplification of N-terminus region or sense primer (S2a-delNTAD) and antisense primer (HIF2aBam/ AS3098-3076) for amplification of C-terminus region, and BamH I-digested fragment of pcDNA3.1-HA-HIF-2αDM-Myc-His as a template. Each reaction products were mixed, and PCR was performed using sense primer (HIF2a-NotI/S489-511) and antisense primer (HIF2aBam/AS3098-3076), followed by insertion into pCR2.1 TOPO-TA vector, termed pCR2.1-HIF-2αDM(∆N-TAD). The HIF-2αDM(∆C-TAD), HIF-2αDM(1-495), HIF-2αDM(1-396), HIF-2αDM(396-823), and HIF-2αDM(581- 823) cDNAs encoding amino acids 1-823, 1-495, 1-396, 396-823, and 581-823, respectively, were generated by PCR using sense primer (HIF2a-NotI/S489-511) and antisense primer (BamHIF2a(2469-2450)) for amplification of HIF-2αDM(∆C-TAD) cDNA, sense primer (HIF2a-NotI/S489-511) and antisense primer (BamHIF2a(1485- 1465)) for amplification of HIF-2αDM(1-495) cDNA, sense primer (HIF2a-NotI/S489- 511) and antisense primer (BamHIF2a(1188-1168)) for amplification of HIF-2αDM(1- 396) cDNA, sense primer (BamHIF2a(1186-1206)) and antisense primer (BamHIF2a (2469-2450)) for amplification of HIF-2αDM(396-823) cDNA or sense primer (BamHIF2a(1741-1761)) and antisense primer (BamHIF2a(2469-2450)) for amplification of HIF-2αDM(581-823) cDNA, and BamH I-digested fragment of pcDNA3.1-HA-HIF-2αDM-Myc-His as a template, followed by insertion into pCR2.1 TOPO-TA vector, termed pCR2.1-HIF-2αDM(∆C-TAD), pCR2.1-HIF-2αDM(1-495), pCR2.1-HIF-2αDM(1-396), pCR2.1-HIF-2αDM(396-823), and pCR2.1-HIF-2αDM (581-823), respectively. The Not I- and BamH I-digested fragments of pCR2.1-HIF- 2αDM(∆N-TAD), pCR2.1-HIF-2αDM(∆C-TAD), pCR2.1-HIF-2αDM(1-495), and pCR2.1-HIF-2αDM(1-396) were subcloned into the corresponding sites of pcDNA3.1- HA-Myc-His and p3xFLAG-Myc-CMV-26 vectors, termed pcDNA3.1-HA-HIF-2αDM (∆N-TAD)-Myc-His, pcDNA3.1-HA-HIF-2αDM(∆C-TAD)-Myc-His, pcDNA3.1-HA- HIF-2αDM(1-495)-Myc-His, pcDNA3.1-HA-HIF-2αDM(1-396)-Myc-His, p3xFLAG- HIF-2αDM(∆N-TAD), p3xFLAG-HIF-2αDM(∆C-TAD), p3xFLAG-HIF-2αDM(1- 495), and p3xFLAG-HIF-2αDM(1-396), respectively. The resulting vectors encode mutants HIF-2α with the N-terminal HA-tag and C-terminal Myc- and His-tags or the

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three tandem repeats of N-terminal FLAG-tags, respectively. The Not I- and Kpn I-digested fragments of pcDNA3.1-HA-HIF-2α-Myc-His, pcDNA3.1-HA-HIF-2αDM- Myc-His, pcDNA3.1-HA-HIF-2αDM(∆N-TAD)-Myc-His, pcDNA3.1-HA-HIF-2αDM (∆C-TAD)-Myc-His, pcDNA3.1-HA-HIF-2αDM(1-495)-Myc-His, and pcDNA3.1-HA- HIF-2αDM(1-396)-Myc-His were subcloned into the corresponding sites of pACT (Promega Corp.) vector, termed pACT-HIF-2α, pACT-HIF-2αDM, pACT-HIF-2αDM (∆N-TAD), pACT-HIF-2αDM(∆C-TAD), pACT-HIF-2αDM(1-495), and pACT-HIF- 2αDM(1-396), respectively. The BamH I-digested fragments of pCR2.1-HIF-2αDM (396-823) and pCR2.1-HIF-2αDM(581-823) were subcloned into the corresponding site of pACT vector, termed pACT-HIF-2αDM(396-823) and pACT-HIF-2αDM(581- 823), respectively. The resulting vectors encode mutants HIF-2α fused with herpes simplex virus VP16 activation domain at the N-terminus. To construct of E. coli expression vector, pGEX-HIF-2αDM(396-823), the BamH I-digested fragment of pCR2.1-HIF-2αDM(396-823) was subcloned into the corresponding site of pGEX5X-1 (Amersham Pharmacia Biotech., Piscataway, NJ, USA) vector. The resulting vector encodes mutant HIF-2α fused with glutathione S-transferase (GST) at the N-terminus. Human VHL cDNA (GenBank Accession No. NM_000551) was amplified by two-sequential nested PCR. The first PCR was performed using sense primer (VHL(74-93)F) and antisense primer (VHL(950-927)R), and human brain (cerebral cortex) Marathon-ready cDNA as a template. The second PCR was performed using sense primer (EcoVHL(213-233)F) and antisense primer (BamVHL(857-838)R), and the first PCR product as a template, followed by insertion into pCR2.1 TOPO-TA vector, termed pCR2.1-VHL. The VHL(-ATG) cDNA was amplified by PCR using sense primer (EcoVHL(-ATG)) and antisense primer (BamVHL(857-838)R), and EcoR I-digested fragment of pCR2.1-VHL as a template. The EcoR I- and BamH I-digested fragments of pCR2.1-VHL(-ATG) was subcloned into the corresponding sites of pcDNA3.1-Myc-Myc-His vector, termed pcDNA3.1-Myc-VHL. The resulting vector encodes the mutant VHL with the N-terminal Myc-tag. Human ERα mammalian expression vector, pCAGGS-ERα, was kindly provided by Dr. Testuya Adachi (Kyoto Prefecture University of Medicine, Kyoto, Japan). The ERα, ERα-AB, ERα-C, and

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ERα-DEF cDNAs encoding full-length ERα, NTD (A/B domains; amino acids 1-185), DBD (C domain; amino acids 185-250), and C-terminal domain (D/E/F domains; amino acids 250-596) were generated by PCR using sense primer (BamER(1-21)-F) and antisense primer (BamER(1788-1768)-R) for amplification of ERα cDNA, sense primer (BamER(1-21)-F) and antisense primer (BamER(555-535)-R) for amplification of ERα-AB cDNA, sense primer (BamER(553-573)-F) and antisense primer (BamER(750 -730)-R) for amplification of ERα-C cDNA, and sense primer (BamER(748-768)-F) and antisense primer (BamER(1788-1768)-R) for amplification of ERα-DEF cDNA, and EcoR I-digested fragment of pCAGGS-ERα as a template, followed by insertion into pCR2.1 TOPO-TA vector, termed pCR2.1-ERα, pCR2.1-ERα-AB, pCR2.1-ERα-C, and pCR2.1-ERα-DEF, respectively. The BamH I-digested fragments of pCR2.1-ERα, pCR2.1-ERα-AB, pCR2.1-ERα-C, pCR2.1-ERα-DEF, and pCR2.1-ERα-E were subcloned into the corresponding site of pBIND (Promega Corp.) vector, termed pBIND -ERα, pBIND-ERα-AB, pBIND-ERα-C, pBIND-ERα-DEF, and pBIND-ERα-E, respectively. The resulting vectors encode mutants ERα fussed with yeast GAL4 DNA- binding domain (GAL4DBD) at the N-terminus. The ERα-E cDNA encoding LBD (E domain; amino acids 298-554) was generated by PCR using sense primer EcoRI-ERaF (892-) and Xho-ERaR(-1662STOP), and EcoR I-digested fragment of pCAGGS-ERα as a template, followed by insertion into pCR2.1 TOPO-TA vector, termed pCR2.1-ERα-E. To construct of E. coli expression vectors, pGEX-ERα-E and pET30a(+)-His-Myc-ERα -E, the EcoR I- and Xho I-digested fragment of pCR2.1-ERα-E was subcloned into the corresponding sites of pGEX5X-1 and pET30a(+)-Myc vectors, respectively. The resulting vectors encode mutant ERα fused with GST at the N-terminus or with His- and Myc-tags at the N-terminus. The BamH I- and Not I-digested fragment of pGEX-ERα-E was subcloned into the corresponding sites of pBIND vector, termed pBIND-ERα-E. The resulting vector encodes mutant ERα fused with yeast GAL4DBD at the N-terminus. Luciferase reporter vector, p3xERE-SV40-Luc, which has three tandem repeats of an estrogen response element was constructed according to the methods of Legler et al. [76]. Annealed oligonucleotides containing three tandem repeats of a consensus estrogen response element, (3xERE Nhe/Bgl-F and 3xERE

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Nhe/Bgl-R), were inserted into the Nhe I and Bgl II sites of pGL3-promoter (Promega Corp.) vector.

siRNA-mediated knockdown. Target sequences of siHIF-1α #1, siHIF-1α #2,

and RISC-Free siRNA #1 were described in CHPTER II. Target sequence for siRNA duplexes were as follows: siHIF-2α #1: 5´-CAGCAUCUUUGAUAGCAGUdTdT-3´ (Dharmacon), siHIF-2α #2: 5´-GCGACAGCUGGAGUAUGAAdTdT-3´ (Sigma Aldrich), and siVHL: 5´-AAGAGUACGGCCCUGAAGAAGdTdT-3´ (Sigma Aldrich). The duplexes (10 nM) were introduced in MCF-7 and SK-BR-3 cells using Lipofectamine RNAiMAX reagent and Opti-MEM for 24 h according to the manufacture’s protocol.

Western blot analysis. Rabbit polyclonal anti-HIF-2α antibody was kindly

provided by Dr. Futoshi Shibasaki (Tokyo Metropolitan Institute of Medical Science, Tolyo, Japan). For detection of endogenous HIF-1α, HIF-2α, and ERα, MCF-7 cells

were cultured in DMEM-HG or phenol red-free DMEM-HG supplemented with 5% dextran-coated charcoal-stripped FBS (dFBS) (steroid-free DMEM-HG), followed by exposure to hypoxia for time periods indicated. For detection of exogenous proteins, MCF-7 and SK-BR-3 cells were cultured in steroid-free DMEM-HG or RPMI1640 medium, respectively. Cells were transiently transfected using HilyMax for 24 h, followed by incubation for an additional 12 h in appropriate fresh medium. Cells were harvested and sonicated in HN buffer, followed by centrifugation at 20,000×g for 20 min. The supernatants were subjected to SDS-PAGE and analyzed by Western blot analysis with mouse monoclonal anti-HIF-1α (Clone mgc3), anti-α-tubulin, anti-c-Myc (9B11, Cell Signaling Technology, Beverly, MA, USA), anti-FLAG M2 (Sigma Aldrich) and rabbit polyclonal anti-HIF-2α, anti-ERα antibodies, followed by immunoreaction with the horseradish peroxidase-conjugated goat anti-rabbit IgG and goat anti-mouse IgG, respectively. The immunoreactive proteins were visualized with Immobilon Western Chemiluminescent substrate.

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