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• W2072362263 abstract "Tumor suppressor p53-dependent stress response pathways play an important role in cell fate determination. In this study, we have found that glucose depletion promotes the phosphorylation of AMP-activated protein kinase catalytic subunit α (AMPKα) in association with a significant up-regulation of p53, thereby inducing p53-dependent apoptosis in vivo and in vitro. Thymocytes prepared from glucose-depleted wild-type mice but not from p53-deficient mice underwent apoptosis, which was accompanied by a remarkable phosphorylation of AMPKα and a significant induction of p53 as well as pro-apoptotic Bax. Similar results were also obtained in human osteosarcoma-derived U2OS cells bearing wild-type p53 following glucose starvation. Of note, glucose deprivation led to a significant accumulation of p53 phosphorylated at Ser-46, but not at Ser-15 and Ser-20, and a transcriptional induction of p53 as well as proapoptotic p53 AIP1. Small interference RNA-mediated knockdown of p53 caused an inhibition of apoptosis following glucose depletion. Additionally, apoptosis triggered by glucose deprivation was markedly impaired by small interference RNA-mediated depletion of AMPKα. Under our experimental conditions, down-regulation of AMPKα caused an attenuation of p53 accumulation and its phosphorylation at Ser-46. In support of these observations, enforced expression of AMPKα led to apoptosis and resulted in an induction of p53 at protein and mRNA levels. Furthermore, p53 promoter region responded to AMPKα and glucose deprivation as judged by luciferase reporter assay. Taken together, our present findings suggest that AMPK-dependent transcriptional induction and phosphorylation of p53 at Ser-46 play a crucial role in the induction of apoptosis under carbon source depletion. Tumor suppressor p53-dependent stress response pathways play an important role in cell fate determination. In this study, we have found that glucose depletion promotes the phosphorylation of AMP-activated protein kinase catalytic subunit α (AMPKα) in association with a significant up-regulation of p53, thereby inducing p53-dependent apoptosis in vivo and in vitro. Thymocytes prepared from glucose-depleted wild-type mice but not from p53-deficient mice underwent apoptosis, which was accompanied by a remarkable phosphorylation of AMPKα and a significant induction of p53 as well as pro-apoptotic Bax. Similar results were also obtained in human osteosarcoma-derived U2OS cells bearing wild-type p53 following glucose starvation. Of note, glucose deprivation led to a significant accumulation of p53 phosphorylated at Ser-46, but not at Ser-15 and Ser-20, and a transcriptional induction of p53 as well as proapoptotic p53 AIP1. Small interference RNA-mediated knockdown of p53 caused an inhibition of apoptosis following glucose depletion. Additionally, apoptosis triggered by glucose deprivation was markedly impaired by small interference RNA-mediated depletion of AMPKα. Under our experimental conditions, down-regulation of AMPKα caused an attenuation of p53 accumulation and its phosphorylation at Ser-46. In support of these observations, enforced expression of AMPKα led to apoptosis and resulted in an induction of p53 at protein and mRNA levels. Furthermore, p53 promoter region responded to AMPKα and glucose deprivation as judged by luciferase reporter assay. Taken together, our present findings suggest that AMPK-dependent transcriptional induction and phosphorylation of p53 at Ser-46 play a crucial role in the induction of apoptosis under carbon source depletion. AMP-activated protein kinase (AMPK) 3The abbreviations used are: AMPKAMP-activated protein kinaseDAPI4,6-diamidino-2-phenylindoleFACSfluorescence-activated cell sorterFBSfetal bovine serumGAPDHglyderaldehyde-3-phosphate dehydrogenaseIBimmunoblottingPARPpoly(ADP-ribose) polymerasePBSphosphate-buffered salineRTreverse transcriptionsiRNAsmall interference RNA. was originally identified as an enzyme that has an ability to inhibit hydroxymethylglutaryl-CoA reductase (1Beg G.H. Allmann D.W. Gibson D.M. Biochem. Biophys. Res. Commun. 1973; 54: 1362-1369Crossref PubMed Scopus (219) Google Scholar) and also regulate acetyl-CoA carboxylase by reversible phosphorylation (2Carlson C.A. Kim K.H. J. Biol. Chem. 1973; 248: 378-380Abstract Full Text PDF PubMed Google Scholar). Subsequent studies demonstrated that AMPK is widely expressed and exists as a heterotrimeric complex, which consists of a catalytic subunit (α) and two regulatory subunits (β and γ). The mammalian genome contains seven AMPK genes encoding two α (α1 and α2), two β (β1 and β2), and three γ (γ1, γ2, and γ3) isoforms (3Rutter G.A. Da Silva Xavier G. Leclerc I. Biochem. J. 2003; 375: 1-16Crossref PubMed Scopus (287) Google Scholar, 4Carling D. Trends Biochem. Sci. 2004; 29: 18-24Abstract Full Text Full Text PDF PubMed Scopus (963) Google Scholar, 5Hardie D.G. J. Cell Sci. 2004; 117: 5479-5487Crossref PubMed Scopus (965) Google Scholar). The catalytic α subunit is composed of three functional domains, including an NH2-terminal Ser/Thr protein kinase domain, a central auto-inhibitory region, and a COOH-terminal regulatory subunit-binding domain. AMPK acts as an intracellular energy sensor by monitoring cellular energy levels. For example, AMPK becomes activated by the tumor suppressor LKB1 complex-mediated phosphorylation at Thr-172 in response to certain energy-depleting stresses such as glucose deprivation, hypoxia, and oxidative stress, which increase the intracellular AMP:ATP ratio (6Hardie D.G. Endocrinology. 2003; 144: 5179-5183Crossref PubMed Scopus (841) Google Scholar, 7Xing Y. Musi N. Fujii N. Zou L. Luptak I. Hirshman M.F. Goodyear L.J. Tian R. J. Biol. Chem. 2003; 278: 28372-28377Abstract Full Text Full Text PDF PubMed Scopus (184) Google Scholar, 8Hawley S.A. Boudeau J. Reid J.L. Mustard K.J. Udd L. Makela T.P. Allesi D.R. Hardie D.G. J. Biol. 2003; 2: 28Crossref PubMed Google Scholar, 9Woods A. Johnstone S.R. Dickerson K. Leiper F.C. Fryer L.G. Neumann D. Schlattner U. Wallimann T. Carlson M. Carling D. Curr. Biol. 2003; 13: 2004-2008Abstract Full Text Full Text PDF PubMed Scopus (1352) Google Scholar, 10Shaw R.J. Bardeesy N. Manning B.D. Lopez L. Kosmatka M. De-Pinho R.A. Cantley L.C. Cancer Cell. 2004; 6: 91-99Abstract Full Text Full Text PDF PubMed Scopus (886) Google Scholar). AMPK can also be activated allosterically in the AMP:ATP ratio (11Hardie D.G. Carling D. Carlson M. Annu. Rev. Biochem. 1998; 67: 821-855Crossref PubMed Scopus (1281) Google Scholar). Upon activation, AMPK down-regulates the ATP consuming metabolic pathways and activates the energy-generating processes through phosphorylating the primary targets involved in energy metabolism, thereby maintaining energy balance within cells (4Carling D. Trends Biochem. Sci. 2004; 29: 18-24Abstract Full Text Full Text PDF PubMed Scopus (963) Google Scholar). AMP-activated protein kinase 4,6-diamidino-2-phenylindole fluorescence-activated cell sorter fetal bovine serum glyderaldehyde-3-phosphate dehydrogenase immunoblotting poly(ADP-ribose) polymerase phosphate-buffered saline reverse transcription small interference RNA. Pro-apoptotic p53 is a founding member of the p53 tumor suppressor family and acts as a critical regulator of many cellular processes such as cell cycle arrest and apoptosis (12Vogelstein B. Lane D. Levine A.J. Nature. 2000; 408: 307-310Crossref PubMed Scopus (5850) Google Scholar). p53 is frequently mutated in over 50% of human tumors (13Caron de Fromentel C. Soussi T. Genes Chromosomes Cancer. 1992; 4: 1-15Crossref PubMed Scopus (536) Google Scholar, 14Greenblatt M.S. Bennett W.P. Hollstein M. Harris C.C. Cancer Res. 1994; 54: 4855-4878PubMed Google Scholar, 15Chao C. Hergenhahn M. Kaeser M.D. Wu Z. Saito S. Iggo R. Hollstein M. Appella E. Xu Y. J. Biol. Chem. 2003; 278: 41028-41033Abstract Full Text Full Text PDF PubMed Scopus (118) Google Scholar), and p53-deficient mice developed spontaneous tumors (16Donehower L.A. Harvey M. Slagle B.L. McArthur M.J. Montgomery Jr., C.A. Butel J.S. Bradley A. Nature. 1992; 356: 215-221Crossref PubMed Scopus (4054) Google Scholar). p53 acts as a nuclear sequence-specific transcription factor and transactivates its numerous target genes implicated in cell cycle arrest and apoptotic cell death, including p21WAF1, Bax, Puma, Noxa, and p53 AIP1. Its pro-apoptotic function is closely linked to its DNA binding activity. Indeed, over 90% of the p53 mutation is detected within its sequence-specific DNA-binding domain (13Caron de Fromentel C. Soussi T. Genes Chromosomes Cancer. 1992; 4: 1-15Crossref PubMed Scopus (536) Google Scholar, 14Greenblatt M.S. Bennett W.P. Hollstein M. Harris C.C. Cancer Res. 1994; 54: 4855-4878PubMed Google Scholar, 15Chao C. Hergenhahn M. Kaeser M.D. Wu Z. Saito S. Iggo R. Hollstein M. Appella E. Xu Y. J. Biol. Chem. 2003; 278: 41028-41033Abstract Full Text Full Text PDF PubMed Scopus (118) Google Scholar). Intracellular p53 protein levels are tightly regulated predominantly through the ubiquitin-proteasome protein degradation pathway. MDM2, which interacts with the NH2-terminal transactivation domain of p53 and inhibits its transcriptional activity, is one of the ubiquitin-protein isopeptide ligases for p53 (17Haupt Y. Maya R. Kazaz A. Oren M. Nature. 1997; 387: 296-299Crossref PubMed Scopus (3750) Google Scholar, 18Kubbutat M.H. Jones S.N. Vousden K. Nature. 1997; 387: 299-303Crossref PubMed Scopus (2860) Google Scholar). In response to various types of cellular stress, including DNA damage, hypoxia, nucleotide pool reduction, and thermal shock, p53 is induced to be stabilized as well as activated in the cell nucleus, and thereby plays a key role in the regulation of cell fate determination (19Prives C. Hall P.A. J. Pathol. 1999; 187: 112-126Crossref PubMed Scopus (1234) Google Scholar, 20Sionov R.V. Haupt Y. Oncogene. 1999; 18: 6145-6157Crossref PubMed Scopus (503) Google Scholar, 21Vousden K.H. Lu X. Nat. Rev. Cancer. 2002; 2: 594-604Crossref PubMed Scopus (2738) Google Scholar). Recently, it has been shown that cells treated with low glucose arrest in the G1 phase of the cell cycle in association with a significant activation of AMPK (22Jones R.G. Plas D.R. Kubek S. Buzzai M. Mu J. Xu Y. Birnbaum M.J. Thompson C.B. Mol. Cell. 2005; 18: 283-293Abstract Full Text Full Text PDF PubMed Scopus (1304) Google Scholar). According to their results, AMPK-mediated cell cycle arrest in response to low glucose required the phosphorylation of tumor suppressor p53 at Ser-15. Because stress-induced phosphorylation of p53 at Ser-15, which disrupts the p53-MDM2 interaction, enhances its activity as well as stability (19Prives C. Hall P.A. J. Pathol. 1999; 187: 112-126Crossref PubMed Scopus (1234) Google Scholar, 20Sionov R.V. Haupt Y. Oncogene. 1999; 18: 6145-6157Crossref PubMed Scopus (503) Google Scholar, 21Vousden K.H. Lu X. Nat. Rev. Cancer. 2002; 2: 594-604Crossref PubMed Scopus (2738) Google Scholar), it is likely that p53 plays an important role in the regulation of cell cycle arrest caused by glucose limitation. Indeed, p53-deficient cells failed to arrest under low glucose conditions (22Jones R.G. Plas D.R. Kubek S. Buzzai M. Mu J. Xu Y. Birnbaum M.J. Thompson C.B. Mol. Cell. 2005; 18: 283-293Abstract Full Text Full Text PDF PubMed Scopus (1304) Google Scholar). Consistent with these results, activation of AMPK in human hepatocellular carcinoma-derived HepG2 cells bearing wild-type p53 resulted in G1 cell cycle arrest through stabilization of p53 (23Imamura K. Ogura T. Kishimoto A. Kaminishi M. Esumi H. Biochem. Biophys. Res. Commun. 2001; 287: 562-567Crossref PubMed Scopus (304) Google Scholar). Previously, Stefanelli et al. (24Stefanelli C. Stanic I. Bonavita F. Flamigni F. Pignatti C. Guarnieri C. Caldarera C.M. Biochem. Biophys. Res. Commun. 1998; 243: 821-826Crossref PubMed Scopus (80) Google Scholar) described that AMPK has a protective role against thymocyte apoptosis in response to dexamethasone treatment. In this study, we have found that glucose deprivation induces phosphorylation of AMPKα and promotes p53-dependent apoptotic cell death in vivo and in vitro. Under our experimental conditions, p53 was induced in response to glucose depletion at mRNA and at protein levels. Our present findings suggest that AMPK acts as a metabolic sensor to determine cell fate through the activation of pro-apoptotic p53. Mice–Six-week-old male c57BL/6 mice (23–24 g), which were purchased from Charles River Laboratories (Tokyo, Japan), were housed in an animal facility maintained on a 12-h light/dark cycle at a constant temperature of 22 ± 1 °C and given free access to food and water ad libitum. For starvation, food was withdrawn from the cages at the onset of the dark cycle for the indicated times, whereas access to water was allowed. All experiments with these mice were carried out with the approval of the Chiba Cancer Center Experimental Animal Care and Use Committee. Cell Lines and Transfection–Human osteosarcoma-derived U2OS cells were maintained in Dulbecco's modified Eagle's medium supplemented with 10% heat-inactivated fetal bovine serum (FBS; Invitrogen), 50 μg/ml penicillin, and 50 μg/ml streptomycin (Invitrogen). Human lung carcinoma H1299 cells were grown in RPMI 1640 medium supplemented with 10% heat-inactivated FBS plus antibiotics mixture. These cells were cultured in a humidified atmosphere of 5% CO2, 95% air at 37 °C. Where indicated, U2OS cells were cultured in glucose-free Dulbecco's modified Eagle's medium (Invitrogen) supplemented with 10% dialyzed FBS. For transient transfection, U2OS and H1299 cells were transfected with the indicated combinations of the expression plasmids using Lipofectamine 2000 transfection reagent (Invitrogen) according to the manufacturer's instructions. pcDNA3 (Invitrogen) was used as a blank plasmid to balance the amount of DNA introduced in transient transfection. RNA Extraction and RT-PCR–Total RNA was prepared from the indicated cells by using the RNeasy mini kit (Qiagen, Valencia, CA), according to the manufacturer's protocol, and reverse-transcribed with SuperScript II reverse transcriptase (Invitrogen). The resultant cDNA was amplified by PCR with rTaq DNA polymerase (Takara, Ohtsu, Japan) using the following primers: p53, 5′-CTGCCCTCAACAAGATGTTTTG-3′ (forward) and 5′-CTATCTGAGCAGCGCTCATGG-3′ (reverse); p21WAF1, 5′-ATGAAATTCACCCCCTTTCC-3′ (forward) and 5′-CCCTAGGCTGTGCTCACTTC-3′ (reverse); Bax, 5′-TTTGCTTCAGGGTTTCATCC-3′ (forward) and 5′-CAGTTGAAGTTGCCGTCAGA-3′ (reverse); p53 AIP1, 5′-TGGCTCCAGGAAGGAAAGGC-3′ (forward) and 5′-TGCTTTCTGCAGACAGGGCC-3′ (reverse); AMPKα1, 5′-CAGGGACTGCTACTCCACAGAGA-3′ (forward) and 5′-CCTTGAGCCTCAGCATCTGAA-3′ (reverse); AMPKα2, 5′-CAACTGCAGAGAGCCATTCACTT-3′ (forward) and 5′-GGTGAAACTGAAGACAATGTGCTT-3′ (reverse); and GAPDH, 5′-ACCTGACCTGCCGTCTAGAA-3′ (forward) and 5′-TCCACCACCCTGTTGCTGTA-3′ (reverse). The expression of GAPDH was measured as an internal control. Immunoblotting–Cells were scraped off the plates and transferred into the microcentrifuge tubes. The cells were then lysed in lysis buffer containing 10 mm Tris-HCl, pH 8.0, 150 mm NaCl, 2 mm EGTA, 50 mm β-mercaptoethanol,1% Triton X-100, a commercial protease inhibitor mixture (Sigma) and phosphatase inhibitor mixture (Sigma) for 30 min on ice, and subjected to a brief sonication for 10 s at 4 °C followed by centrifugation at 15,000 rpm at 4 °C for 10 min to remove insoluble materials. The protein concentrations were measured using the Bradford protein assay according to the manufacturer's instructions (Bio-Rad). The equal amounts of protein (20 μg) were separated by 10% SDS-PAGE and electrophoretically transferred onto polyvinylidene difluoride membranes (Immobilon-P, Millipore, Bedford, MA). The transferred membranes were blocked with Tris-buffered saline containing 5% nonfat dry milk and 0.1% Tween 20 at 4 °C overnight. After blocking, the membranes were incubated with monoclonal anti-p53 (DO-1; Oncogene Research Products, Cambridge, MA), monoclonal anti-Bax (6A7; eBioscience, San Diego, CA), polyclonal anti-phospho-p53 at Ser-15 (Cell Signaling, Beverly, MA), polyclonal anti-phospho-p53 at Ser-20 (Cell Signaling), polyclonal anti-phospho-p53 at Ser-46 (Cell Signaling), polyclonal anti-p21WAF1 (H-164; Santa Cruz Biotechnology), polyclonal anti-AMPKα (Cell Signaling), polyclonal anti-phospho-AMPKα (Cell Signaling), polyclonal anti-PARP (Cell Signaling), or with polyclonal anti-actin (20Sionov R.V. Haupt Y. Oncogene. 1999; 18: 6145-6157Crossref PubMed Scopus (503) Google Scholar, 21Vousden K.H. Lu X. Nat. Rev. Cancer. 2002; 2: 594-604Crossref PubMed Scopus (2738) Google Scholar, 22Jones R.G. Plas D.R. Kubek S. Buzzai M. Mu J. Xu Y. Birnbaum M.J. Thompson C.B. Mol. Cell. 2005; 18: 283-293Abstract Full Text Full Text PDF PubMed Scopus (1304) Google Scholar, 23Imamura K. Ogura T. Kishimoto A. Kaminishi M. Esumi H. Biochem. Biophys. Res. Commun. 2001; 287: 562-567Crossref PubMed Scopus (304) Google Scholar, 24Stefanelli C. Stanic I. Bonavita F. Flamigni F. Pignatti C. Guarnieri C. Caldarera C.M. Biochem. Biophys. Res. Commun. 1998; 243: 821-826Crossref PubMed Scopus (80) Google Scholar, 25Oda K. Arakawa H. Tanaka T. Matsuda K. Tanikawa C. Mori T. Nishimori H. Tamai K. Tokino T. Nakamura Y. Taya Y. Cell. 2000; 102: 849-862Abstract Full Text Full Text PDF PubMed Scopus (1028) Google Scholar, 26Matsuda K. Yoshida K. Taya Y. Nakamura K. Nakamura Y. Arakawa H. Cancer Res. 2002; 62: 2883-2889PubMed Google Scholar, 27Rathmell J.C. Fox C.J. Plas D.R. Hammerman P.S. Cinalli R.M. Thompson C.B. Mol. Cell. Biol. 2003; 23: 7315-7328Crossref PubMed Scopus (468) Google Scholar, 28Hofmann T.G. Moller A. Sirma H. Zentgraf H. Taya Y. Droge W. Will H. Schmitz M.L. Nat. Cell Biol. 2002; 4: 1-10Crossref PubMed Scopus (503) Google Scholar, 29D'Orazi G. Cecchinelli B. Bruno T. Manni I. Higashimoto Y. Saito S. Gostissa M. Coen S. Marchetti A. Del Sal G. Piaggio G. Fanciulli M. Appella E. Soddu S. Nat. Cell Biol. 2002; 4: 11-19Crossref PubMed Scopus (578) Google Scholar, 30Dauth I. Kruger J. Hofmann T.G. Cancer Res. 2007; 67: 2274-2279Crossref PubMed Scopus (67) Google Scholar, 31Yoshida K. Liu H. Miki Y. J. Biol. Chem. 2006; 281: 5734-5740Abstract Full Text Full Text PDF PubMed Scopus (147) Google Scholar, 32Reich N.C. Levine A.J. Nature. 1984; 308: 199-201Crossref PubMed Scopus (377) Google Scholar, 33Noda A. Toma-Aiba Y. Fujiwara Y. Oncogene. 2000; 19: 21-31Crossref PubMed Scopus (24) Google Scholar; Sigma) antibody for 1 h at room temperature. After incubation with primary antibodies, the membranes were incubated with horseradish peroxidase-coupled goat anti-mouse or anti-rabbit IgG secondary antibody (Cell Signaling) for 1 h at room temperature. Immunoblots were visualized by ECL detection reagents according to the manufacturer's instructions (Amersham Biosciences). Immunoprecipitation–At the indicated time points after glucose depletion, whole cell lysates prepared from U2OS cells were pre-cleared with 30 μl of protein G-Sepharose beads for 90 min at 4 °C, and supernatants were incubated with the indicated antibodies overnight at 4 °C. After incubation, the reaction mixtures were mixed with 30 μl of protein G-Sepharose beads and incubated for 1 h at 4 °C. The immune complexes were eluted with SDS-sample buffer, and separated by 10% SDS-PAGE followed by immunoblotting with the indicated antibodies. Indirect Immunofluorescence Microscopy–U2OS cells were cultured in the absence of glucose. At the indicated times after glucose deprivation, cells were fixed in 3.7% formaldehyde for 30 min at room temperature, permeabilized in 0.2% Triton X-100 for 5 min at room temperature, and then blocked with 3% bovine serum albumin in phosphate-buffered saline (PBS) for 1 h at room temperature. After blocking, cells were washed in PBS and incubated with polyclonal anti-phospho-AMPKα and monoclonal anti-p53 or with polyclonal anti-phospho-AMPKα and monoclonal anti-nucleolin antibodies (StressGen Biotechnologies, Cambridge, UK) for 1 h at room temperature, followed by the incubation with fluorescein isothiocyanate-conjugated anti-rabbit IgG and rhodamine-conjugated anti-mouse IgG (Invitrogen) for 1 h at room temperature. Cell nuclei were stained with DAPI. The specific fluorescence was observed by using a confocal laser scanning microscope (Olympus, Tokyo, Japan). Flow Cytometry–After glucose deprivation, both floating and attached cells were collected by low speed centrifugation and washed in PBS. The cells were treated with 500 μg/ml of RNase A (Sigma) and subsequently stained with 50 μg/ml of propidium iodide (Sigma) for 30 min at room temperature. Then the DNA content indicated by propidium iodide staining was analyzed by FACSCalibur flow cytometer (BD Biosciences). Luciferase Reporter Assay–p53-deficient H1299 cells were plated in 12-well plates at a density of 50,000 cells/well and transiently co-transfected with a constant amount of a luciferase reporter construct driven by the p53 promoter (100 ng) and 10 ng of Renilla luciferase expression plasmid (pRL-TK) together with or without the increasing amounts of the expression plasmids for AMPKα1 plus AMPK α2 (50, 100, or 200 ng). For all transfections, the total DNA amounts were kept constant (510 ng) using empty parental plasmid. Forty eight hours after transfection, cells were lysed, and their luciferase activities were measured by dual luciferase reporter assay system (Promega, Madison, WI). Results represent an average firefly luciferase value after normalization to Renilla luciferase signal. Each experiment was performed at least three times by triplicates. Construction of the Expression Plasmids for AMPKα1 and AMPKα2–To generate the expression plasmids for AMPKα1 and AMPKα2, we employed PCR-based amplification using cDNA prepared from U2OS cells as a template. For AMPKα1, the 5′-part of the entire coding region was amplified by PCR using the following primer sequences: 5′-GGAATTCCATGCGCAGACTCAGTTCCTG-3′ and 5′-CTGCAGCATATGTTTCAAAAG-3′, which include EcoRI and PstI restriction sites, respectively. The 3′-part of the entire coding region was also amplified by PCR using the following primer sequences: 5′-CTGCAGGTGGATCCCATGAAG-3′ and 5′-CCGCTCGAGCGGTTATTGTGCAAGAATTT-3′, which contain PstI and XhoI restriction sites, respectively. The resultant 5′- and 3′-parts of the entire coding regions were digested completely with EcoRI and PstI or with PstI and XhoI, respectively, and subcloned into EcoRI and XhoI restriction sites of pcDNA3 (Invitrogen) to give pcDNA3-AMPKα1. For AMPKα2, the 5′-part of the entire coding region was amplified by PCR using the following primer sequences: 5′-CGGGATCCCGATGGCTGAGAGAAGCAGAAGC-3′ and 5′-ACTAGTTCTCAGAAATTCAC-3′, which include BamHI and SpeI restriction sites, respectively. The 3′-part of the entire coding region was also amplified by PCR using the following primer sequences: 5′-ACTAGTTGCGGATCTCCAAATTATAC-3′ and 5′-GGAATTCCTCAACGGGCTAAAGTAGTAGTAATC-3′, which contain SpeI and EcoRI restriction sites, respectively. The amplified PCR products corresponding to 5′-or3′-part of the entire coding region were treated with BamHI and SpeI or with SpeI and EcoRI, respectively, and introduced into BamHI and EcoRI sites of pcDNA3 (Invitrogen) to give pcDNA3-AMPKα2. Nucleotide sequences of the PCR products were determined to verify the absence of random mutations. RNA Interference–To knock down the endogenous AMPKα, U2OS cells were transiently transfected with 10 nm of the chemically synthesized siRNAs targeting AMPKα1 and AMPKα2 or with the nonsilencing control siRNA (Invitrogen) using Lipofectamine™ RNAiMAX (Invitrogen) according to the manufacturer's recommendations. Total RNA and whole cell lysates were prepared 48 h after transfection. siRNA sequences used in the present study are available upon request. Glucose Depletion Induces Apoptotic Cell Death in Vivo and in Vitro–To examine whether glucose depletion could induce apoptotic cell death in vivo, we have prepared thymocytes from wild-type and p53-deficient mice that were maintained in the absence of glucose for 24 h, and their cell cycle distributions were analyzed by FACS. As shown in Fig. 1A, glucose deprivation resulted in a significant increase in number of thymocytes with sub-G1 DNA content in wild-type mice, whereas glucose depletion had undetectable effects on thymocytes derived from p53-deficient mice, indicating that glucose deprivation-mediated apoptotic cell death might be regulated in a p53-dependent manner. Consistent with the previous observations (22Jones R.G. Plas D.R. Kubek S. Buzzai M. Mu J. Xu Y. Birnbaum M.J. Thompson C.B. Mol. Cell. 2005; 18: 283-293Abstract Full Text Full Text PDF PubMed Scopus (1304) Google Scholar), glucose depletion promoted the extensive phosphorylation of AMPKα in wild-type thymocytes (Fig. 1B). Under our experimental conditions, p53 accumulated in response to glucose starvation. Intriguingly, the accumulation of p53 was clearly associated with a significant induction of AMPKα phosphorylation as well as pro-apoptotic Bax, which is one of the direct targets of p53 (19Prives C. Hall P.A. J. Pathol. 1999; 187: 112-126Crossref PubMed Scopus (1234) Google Scholar, 20Sionov R.V. Haupt Y. Oncogene. 1999; 18: 6145-6157Crossref PubMed Scopus (503) Google Scholar, 21Vousden K.H. Lu X. Nat. Rev. Cancer. 2002; 2: 594-604Crossref PubMed Scopus (2738) Google Scholar). Additionally, proteolytic cleavage of PARP, which is one of the substrates of the activated caspase-3, was induced in response to glucose deprivation. In contrast, the expression levels of p21WAF1, which have been shown to be implicated in p53-dependent cell cycle arrest (19Prives C. Hall P.A. J. Pathol. 1999; 187: 112-126Crossref PubMed Scopus (1234) Google Scholar, 20Sionov R.V. Haupt Y. Oncogene. 1999; 18: 6145-6157Crossref PubMed Scopus (503) Google Scholar, 21Vousden K.H. Lu X. Nat. Rev. Cancer. 2002; 2: 594-604Crossref PubMed Scopus (2738) Google Scholar), remained almost unchanged regardless of glucose deprivation. To further confirm this issue in vitro, human osteosarcoma-derived U2OS cells bearing wild-type p53 were cultured in medium completely deficient in glucose for the indicated times, and we examined their cell cycle distributions by FACS. As shown in Fig. 2A, glucose depletion led to a massive apoptotic cell death. RT-PCR analysis revealed that the expression levels of AMPKα1 and AMPKα2 remain unchanged (Fig. 2B). Unexpectedly, p53 was transcriptionally induced upon removal of glucose, which was associated with the up-regulation of pro-apoptotic Bax and p53 AIP1 (25Oda K. Arakawa H. Tanaka T. Matsuda K. Tanikawa C. Mori T. Nishimori H. Tamai K. Tokino T. Nakamura Y. Taya Y. Cell. 2000; 102: 849-862Abstract Full Text Full Text PDF PubMed Scopus (1028) Google Scholar, 26Matsuda K. Yoshida K. Taya Y. Nakamura K. Nakamura Y. Arakawa H. Cancer Res. 2002; 62: 2883-2889PubMed Google Scholar), although the expression levels of p21WAF1 remained almost constant during the glucose deprivation-mediated apoptotic cell death. Immunoblot analysis demonstrated that AMPKα is induced to be phosphorylated following glucose starvation (Fig. 2C). In contrast, total amounts of AMPKα remained unchanged even in the absence of glucose. Additionally, glucose deprivation led to an accumulation of p53 as well as an induction of p53 phosphorylation at Ser-46 but not at Ser-15 and Ser-20. As expected, the expression levels of pro-apoptotic Bax but not of p21WAF1 increased, and proteolytic cleavage of PARP was detectable in response to glucose deprivation. Effects of p53 Knockdown on Glucose Deprivation-mediated Apoptotic Cell Death–To ask whether p53 could be involved in apoptotic cell death in response to glucose removal, U2OS cells were transiently transfected with control siRNA or with siRNA against p53. Twenty four hours after transfection, cells were switched into fresh medium lacking glucose. At the indicated time points after glucose starvation, whole cell lysates were prepared and analyzed for the expression levels of p53 by immunoblotting. As shown in Fig. 3A, p53 was induced to accumulate in cells transfected with control siRNA, whereas the amounts of p53 were kept at extremely low levels in cells transfected with siRNA against p53. FACS analysis revealed that siRNA-mediated knockdown of p53 strongly reduces the number of cells with sub-G1 DNA content in response to glucose deprivation as compared with that of control cells (Fig. 3B). Similarly, p53-deficient human osteosarcoma-derived SAOS-2 cells underwent apoptotic cell death in response to glucose deprivation to a lesser degree, suggesting that p53 contributes at least in part to the induction of apoptotic cell death following glucose depletion. Because glucose deprivation resulted in the strong induction of AMPKα phosphorylation, we examined the effects of AMPKα on glucose deprivation-mediated apoptotic cell death. To this end, siRNA-mediated knockdown of AMPKα1 and AMPKα2 was performed. Twenty four hours after transfection, cells were maintained in the absence of glucose. At the indicated time points after glucose starvation, cells were harvested, and their cell cycle distributions were analyzed by FACS. As shown in Fig. 4A, siRNA-mediated knockdown remarkably inhibited apoptotic cell death caused by glucose removal relative to control cells. Immunoblot analysis demonstrated that simultaneous knockdown of AMPKα1 and AMPKα2 does not induce the phosphorylation of AMPKα as well as the accumulation of p53 in response to glucose deprivation (Fig. 4B). Intriguingly, induction of p53 phosphorylation at Ser-46 was not detectable in cells where AMPKα1 and AMPKα2 were knocked down. Additionally, RT-PCR analysis showed that knockdown of AMPKα1 and AMPKα2 significantly inhibits the transcriptional up-regulation of p53 as well as p53 AIP1 (Fig. 4C). Under our experimental condit" @default.
• W2072362263 created "2016-06-24" @default.
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• W2072362263 creator A5034252509 @default.
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• W2072362263 date "2008-02-01" @default.
• W2072362263 modified "2023-10-04" @default.
• W2072362263 title "Activation of AMP-activated Protein Kinase Induces p53-dependent Apoptotic Cell Death in Response to Energetic Stress" @default.
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