Transcription Factors: Methods and Protocols [1 ed.] 1607617374, 9781607617372

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METHODS

IN

MOLECULAR BIOLOGY™

Series Editor John M. Walker School of Life Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK

For other titles published in this series, go to www.springer.com/series/7651

Transcription Factors Methods and Protocols

Edited by

Paul J. Higgins Center for Cell Biology and Cancer Research, Albany Medical College, Albany, New York, USA

Editor Paul J. Higgins, Ph.D. Center for Cell Biology and Cancer Research Albany Medical College Albany New York USA [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-60761-737-2 e-ISBN 978-1-60761-738-9 DOI 10.1007/978-1-60761-738-9 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2010929390 © Springer Science+Business Media, LLC 2010 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. While the advice and information in this book are believed to be true and accurate at the date of going to press, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper Humana Press is a part of Springer Science+Business Media (www.springer.com)

Preface In compiling “Transcription Factors: Methods and Protocols,” it was the intent to present a methodological resource that highlights several of the most exciting recent developments that have expanded our appreciation for the complexity of transcriptional controls in mammalian cells. This volume is divided into four major sections and includes chapters that specifically focus on the newest experimental approaches that investigators can utilize to: (1) probe mechanisms underlying transcription factor nuclear-cytoplasmic trafficking (i.e., “shuttling”) and (2) assess the impact of post-translational modifications on transcription factor function including regulation of the coupled ubiquitination/degradation process. In each instance, the protocols highlight specific transcription factor family members with particular relevance to human disease. The aim of this book is to present a compilation of state-of-the-art techniques and concepts important to elucidating controls on transcription factor intracellular localization and activity. The topics selected, therefore, are distinct from those covered in other methodologically oriented texts on transcriptional regulation. Each chapter is contributed by prominent experts in their respective fields who, in many cases, not only had significant input into the development of the techniques/methods detailed but currently utilize the described technologies in their own research on transcription factor function. This volume begins with three comprehensive overviews. Part I details a review of biologically critical shuttling and post-translational controls on specific transcriptional proteins that set the stage for the subsequent protocol-focused chapters. Part II presents a collection of techniques to assess a major consideration in transcription factor activity, namely nuclear translocation, controls on nuclear export with high resolution imaging of intracellular trafficking. The chapters in Part III consist of a methodological tour-de-force to evaluate defined post-translational modifications (hydroxylation, phosphorylation, ubiquitination among others), and the involved pathways and enzymes, in function regulation. The protocols in Part IV describe methods for the optimization of transcription factor functional assessments and are a unique contribution to this work. I wish to thank all the authors for their outstanding and cutting-edge contributions to this book. In certain cases, this book represents the first publication of the relevant techniques and the underlying biological contexts. This fact, framed in a presentation of the most current insights into critical molecular events in transcriptional regulation, underscores the generosity of the participants in sharing with the readers their “tricks of the trade” in a benchside reference format. I would also like to take this opportunity to acknowledge the editor of the “Methods in Molecular Biology” series, Dr. John Walker, for his guidance during the early editing process; it should also be recognized that the original concept of a methods volume on transcription factors was, in fact, Dr. Walker’s idea. It is our hope that this work will serve as a frequent reference text for the current and next generation of scientists working to decipher the intricate nature of transcriptional regulation. The support of NIH (GM057242) is greatfully acknowledged. Albany, NY

P.J. Higgins

v

Contents Preface. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

v xi

PART I TRANSCRIPTION FACTOR TRAFFICKING AND POST-TRANSLATIONAL MODIFICATIONS: OVERVIEWS OF MECHANISMS 1 A Review of Post-translational Modifications and Subcellular Localization of Ets Transcription Factors: Possible Connection with Cancer and Involvement in the Hypoxic Response . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Céline Charlot, Hélène Dubois-Pot, Tsvetan Serchov, Yves Tourrette, and Bohdan Wasylyk 2 Regulation of Transcription Factor Function by Targeted Protein Degradation: An Overview Focusing on p53, c-Myc, and c-Jun . . . . . . . . . . . . . . Jukka Westermarck 3 Review of Molecular Mechanisms Involved in the Activation of the Nrf2-ARE Signaling Pathway by Chemopreventive Agents . . . . . . . . . . . . . Aldo Giudice, Claudio Arra, and Maria C. Turco

PART II

3

31

37

CYTOPLASMIC-NUCLEAR TRAFFICKING OF TRANSCRIPTION FACTORS

4 Subnuclear Localization and Intranuclear Trafficking of Transcription Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77 Sayyed K. Zaidi, Ricardo F. Medina, Shirwin M. Pockwinse, Rachit Bakshi, Krishna P. Kota, Syed A. Ali, Daniel W. Young, Jeffery A. Nickerson, Amjad Javed, Martin Montecino, Andre J. van Wijnen, Jane B. Lian, Janet L. Stein, and Gary S. Stein 5 Analysis of Ligand-Dependent Nuclear Accumulation of Smads in TGF-b Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 Douglas A. Chapnick and Xuedong Liu 6 Raf/MEK/MAPK Signaling Stimulates the Nuclear Translocation and Transactivating Activity of FOXM1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113 Richard Y. M. Ma, Tommy H. K. Tong, Wai Ying Leung, and Kwok-Ming Yao 7 Coupling of Dephosphorylation and Nuclear Export of Smads in TGF-b Signaling. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 125 Fangyan Dai, Xueyan Duan, Yao-Yun Liang, Xia Lin, and Xin-Hua Feng 8 Assessing Sequence-Specific DNA Binding and Transcriptional Activity of STAT1 Transcription Factor. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139 Thomas Meyer and Uwe Vinkemeier

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Contents

9 Analysis of Nuclear Export Using Photoactivatable GFP Fusion Proteins and Interspecies Heterokaryons. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kerry-Ann Nakrieko, Iordanka A. Ivanova, and Lina Dagnino 10 Determination of Nuclear Localization Signal Sequences for Krüppel-Like Factor 8 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tina S. Mehta, Farah Monzur, and Jihe Zhao 11 Methods to Measure Nuclear Export of b-Catenin Using Fixed and Live Cell Assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Manisha Sharma and Beric R. Henderson 12 Imaging of Transcription Factor Trafficking in Living Cells: Lessons from Corticosteroid Receptor Dynamics . . . . . . . . . . . . . . . . . . . . . . . . . Mayumi Nishi

PART III

171

187

199

POST-TRANSLATIONAL MODIFICATIONS AND IMPACT ON FUNCTION

13 Hypoxia-Inducible Factors: Post-translational Crosstalk of Signaling Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Elitsa Y. Dimova and Thomas Kietzmann 14 The Basic Helix-Loop-Helix-Leucine Zipper Gene Mitf : Analysis of Alternative Promoter Choice and Splicing . . . . . . . . . . . . . . . . . . . . . . Kapil Bharti, Julien Debbache, Xin Wang, and Heinz Arnheiter 15 Phosphorylation Control of Nuclear Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . Sébastien Lalevée, Christine Ferry, and Cécile Rochette-Egly 16 Regulation of Krüpple-Like Factor 5 by Targeted Protein Degradation . . . . . . . . . Ceshi Chen 17 Post-translational Control of ETS Transcription Factors: Detection of Modified Factors at Target Gene Promoters . . . . . . . . . . . . . . . . . . . Li Li, Janice Saxton, and Peter E. Shaw 18 Integration of Protein Kinases into Transcription Complexes: Identifying Components of Immobilised In Vitro Pre-initiation Complexes . . . . . Hong-Mei Zhang, Stéphanie Vougier, Glenn Hodgson, and Peter E. Shaw 19 Post-translational Modification of p53 by Ubiquitin . . . . . . . . . . . . . . . . . . . . . . . Chunhong Yan 20 Phosphorylation-Dependent Regulation of SATB1, the Higher-Order Chromatin Organizer and Global Gene Regulator . . . . . . . . . . Dimple Notani, Amita S. Limaye, P. Pavan Kumar, and Sanjeev Galande

PART IV

161

215

237 251 267

279

291 305

317

PROTOCOLS FOR OPTIMIZATION OF FUNCTIONAL ASSAYS

21 In Vivo and In Vitro Tools to Identify and Study Transcriptional Regulation of USF-1 Target Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 339 Marie-Dominique Galibert and Sébastien Corre 22 Measuring the Absolute Abundance of the Smad Transcription Factors Using Quantitative Immunoblotting. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 357 David C. Clarke and Xuedong Liu

Contents

ix

23 Flow Cytometry Analysis of Transcription Factors in T Lymphocytes . . . . . . . . . . 377 Diana I. Albu, Danielle Califano, and Dorina Avram 24 Identification of Specific Protein/E-Box-Containing DNA Complexes: Lessons from the Ubiquitously Expressed USF Transcription Factors of the b-HLH-LZ Super Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 391 Marie-Dominique Galibert and Yorann Baron Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 407

Contributors DIANA I. ALBU • Center for Cell Biology and Cancer Research, Albany Medical College, Albany NY, USA SYED A. ALI • Department of Cell Biology, University of Massachusetts Medical School, Worcester MA, USA HEINZ ARNHEITER • Mammalian Development Section, National Institutes of Neurological Disorders and Stroke, National Institutes of Health, Bethesda MD, USA CLAUDIO ARRA • Pascale Foundation National Cancer Institute, Naples, Italy DORINA AVRAM • Center for Cell Biology and Cancer Research, Albany Medical College, Albany NY, USA RACHIT BAKSHI • Department of Cell Biology, University of Massachusetts Medical School, Worcester MA, USA YORANN BARON • Department of Medical Genomics, Rennes Hospital, France KAPIL BHARTI • Mammalian Development Section, National Institutes of Neurological Disorders and Stroke, National Institutes of Health, Bethesda MD, USA DANIELLE CALIFANO • Center for Cell Biology and Cancer Research, Albany Medical College, Albany NY, USA DOUGLAS A. CHAPNICK • Department of Chemistry and Biochemistry, University of Colorado at Boulder, Boulder CO, USA CELINE CHARLOT • Department of Cancer Biology, Institute de Genetique et de Biologie, Moleculaire et Cellulaire, France CESHI CHEN • Center for Cell Biology and Cancer Research, Albany Medical College, Albany NY, USA DAVID C. CLARKE • Department of Chemistry and Biochemistry, University of Colorado at Boulder, Boulder CO, USA SEBASTIEN CORRE • Department of Medical Genomics, Rennes Hospital, France LINA DAGNINO • Departments of Physiology and Pharmacology, and Pediatrics, Child Health Research Institute and Lawson Health Research Institute, University of Western Ontario, London ON, Canada FANGYAN DAI • Department of Molecular & Cellular Biology, Michael E. DeBakey, Department of Surgery, The Dan L. Duncan Cancer Center, Baylor College of Medicine, Houston TX, USA JULIEN DEBBACHE • Mammalian Development Section, National Institutes of Neurological Disorders and Stroke, National Institutes of Health, BethesdaMD, USA ELITSA Y. DINOCA • Department of Chemistry/Biochemistry, University of Kaiserlautern, Kaiserlautern, Germany

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Contributors

XUEYAN DUAN • Department of Molecular & Cellular Biology, Michael E. DeBakey, Department of Surgery, The Dan L. Duncan Cancer Center, Baylor College of Medicine, Houston TX, USA HELENE DUBOIS-POT • Department of Cancer Biology, Institute de Genetique et de Biologie Moleculaire et Cellulaire, France XIN-HUA FENG • Department of Molecular & Cellular Biology, Michael E. DeBakey, Department of Surgery, The Dan L. Duncan Cancer Center, Baylor College of Medicine, Houston TX, USA CHRISTINE FERRY • Department of Functional Denomics, Institut de Genetique et de Biologie Molecularie et Cellulaire, France SANJEEV GALANDE • National Centre for Cell Science, Ganeshkhind Pune Maharashtra, India MARIE-DOMINIQUE GALIBERT • Genetic and Development Institute of Rennes, Transcriptional Regulation and Oncogenesis Team, Rennes University, France ALDO GUIDICE • G. Pascale Foundation National Cancer Institute, Naples, Italy BERIC R. HENDERSON • Westmead Institute for Cancer Research, Westmead Millennium Institute at Westmead Hospital, The University of Sydney, Westmead NSW, Australia GLENN HODGSON • Queen’s Medical Centre, School of Biomedical Sciences, University of Nottingham, Nottingham, UK IORDANKA A. IVANOVA • Departments of Physiology and Pharmacology, University of Western Ontario, London ON, Canada AMJAD JAVED • Department of Cell Biology, University of Massachusetts Medical School, Worcester MA, USA THOMAS KIETZMANN • Department of Chemistry/Biochemistry, University of Kaiserlautern, Kaiserlautern, Germany KRISNA P. KOTA • Department of Cell Biology, University of Massachusetts Medical School, WorcesterMA, USA PAVAN KUMAR • National Centre for Cell Science, Ganeshkhind Pune Maharashtra, India SEBASTIEN LALEVEE • Department of Functional Denomics, Institut de Genetique et de Biologie Molecularie et Cellulaire, France WAI YING LEUNG • Department of Biochemistry, Faculty of Medicine, The University of Hong Kong, Pokfulam Hong Kong, China LI LI • Queen’s Medical Centre, School of Biomedical Sciences, University of Nottingham, Nottingham, UK JANE B. LIAN • Department of Cell Biology, University of Massachusetts Medical School, Worcester MA, USA YAO-YUN LIANG • Michael E. DeBakey Department of Surgery, Baylor College of Medicine, Houston TX, USA AMITA S. LIMAYE • National Centre for Cell Science, Ganeshkhind Pune Maharashtra, India XIA LIN • Michael E. DeBakey Department of Surgery, The Dan L. Duncan Cancer Center, Baylor College of Medicine, Houston TX, USA

Contributors

XUEDONG LIU • Department of Chemistry and Biochemistry, University of Colorado at Boulder, Boulder CO, USA RICHARD Y.M. MA • Faculty of Medicine, Department of Biochemistry, The University of Hong Kong, Pokfulam Hong Kong, China RICARDO F. MEDINA • Department of Cell Biology, University of Massachusetts Medical School, Worcester MA, USA TINA S. MEHTA • Center for Cell Biology and Cancer Research, Albany Medical College, Albany NY, USA THOMAS MEYER • Department of Cardiology, University of Marburg, BaldingerstrasseMarburg, Germany MARTIN MONTECINO • Department of Cell Biology, University of Massachusetts Medical School, Worcester MA, USA FARAH MONZUR • Center for Cell Biology and Cancer Research, Albany Medical College, Albany NY, USA KERRY-ANN NAKRIEKO • Departments of Physiology and Pharmacology, University of Western Ontario, London ON, Canada JEFFERY A. NICKERSON • Department of Cell Biology, University of Massachusetts Medical School, WorcesterMA, USA MAYUMI NISHI • Department of Anatomy and Neurobiology, Kyoto Prefectural University of Medicine, Kyoto, Japan DIMPLE NOTANI • National Centre for Cell Science, Ganeshkhind Pune Maharashtra, India SHIRWIN M. POCKWINSE • Department of Cell Biology, University of Massachusetts Medical School, Worcester MA, USA CECILE ROCHETTE-EGLY • Department of Functional Denomics, Institut de Genetique et de Biologie Molecularie et Cellulaire, France JANICE SAXTON • Queen’s Medical Centre, School of Biomedical Sciences, University of Nottingham, Nottingham, UK TSVETAN SERCHOV • Department of Cancer Biology, Institute de Genetique et de Biologie Moleculaire et Cellulaire, France MANISHA SHARMA • Westmead Institute for Cancer Research, Westmead Millennium Institute at Westmead Hospital, The University of Sydney, Westmead NSW, Australia PETER E. SHAW • Queen’s Medical Centre, School of Biomedical Sciences, University of Nottingham, Nottingham, UK GARY S. STEIN • Department of Cell Biology, University of Massachusetts Medical School, Worcester MA, USA JANET L. STEIN • Department of Cell Biology, University of Massachusetts Medical School, Worcester MA, USA TOMMY H.K. TONG • Faculty of Medicine, Department of Biochemistry, The University of Hong Kong, Pokfulam Hong Kong, China YVES TOURRETTE • Department of Cancer Biology, Institute de Genetique et de Biologie Moleculaire et Cellulaire, France MARIA C. TURCO • Pascale Foundation National Cancer Institute,, Naples, Italy; DiFarma, University of Salerno, Italy

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Contributors

ANDRE J. VAN WIJNEN • Department of Cell Biology, University of Massachusetts Medical School, Worcester MA, USA UWE VINKEMEIER • Queen’s Medical Centre, School of Biomedical Sciences, University of Nottingham Medical School, Nottingham, UK STEPHANIE VOUGIER • Queen’s Medical Centre, School of Biomedical Sciences, University of Nottingham, Nottingham, UK XIN WANG • Mammalian Development Section, National Institutes of Neurological Disorders and Stroke, National Institutes of Health, Bethesda MD, USA BOHDAN WASYLYK • Department of Cancer Biology, Institute de Genetique et de Biologie Moleculaire et Cellulaire, France JUKKA WESTERMARCK • Institute of Medical Technology, University of Tampere, Finland CHUNHONG YAN • Center for Cell Biology and Cancer Research, Albany Medical College, Albany NY, USA KWOK-MING YAO • Faculty of Medicine, Department of Biochemistry, The University of Hong Kong, PokfulamHong Kong, China DANIEL W. YOUNG • Department of Cell Biology, University of Massachusetts Medical School, Worcester MA, USA SAYYED K. ZAIDI • Department of Cell Biology, University of Massachusetts Medical School, Worcester MA, USA HONG-MEI ZHANG • Queen’s Medical Centre, School of Biomedical Sciences, University of Nottingham, Nottingham, UK JIHE ZHAO • Center for Cell Biology and Cancer Research, Albany Medical College, Albany NY, USA

Chapter 1 A Review of Post-translational Modifications and Subcellular Localization of Ets Transcription Factors: Possible Connection with Cancer and Involvement in the Hypoxic Response Céline Charlot, Hélène Dubois-Pot, Tsvetan Serchov, Yves Tourrette, and Bohdan Wasylyk Abstract Post-translational modifications and subcellular localizations modulate transcription factors, generating a code that is deciphered into an activity. We describe our current understanding of these processes for Ets factors, which have recently been recognized for their importance in various biological processes. We present the global picture for the family, and then focus on particular aspects related to cancer and hypoxia. The analysis of Post-translational modification and cellular localization is only beginning to enter the age of “omic,” high content, systems biology. Our snap-shots of particularly active fields point to the directions in which new techniques will be needed, in our search for a more complete description of regulatory pathways. Key words: Ets factors, Cancer, Hypoxia, Phosphorylation, Acetylation, Sumoylation, Ubiquitination, Glycosylation, Subcellular localization, Caenorhabditis elegans, Drosophila melanogaster, Mouse, Human

1. Introduction In this review, we will focus on the E26 transformation specific (Ets) family of transcription factors. Ets factors have often served as a paradigm for the analysis of Post-translational modification and subcellular localization. After a short introduction about the Ets family, we will broadly describe these processes in the family, and then focus on how they are integrated into biological processes, in particular cancer and the response to hypoxia.

Paul J. Higgins (ed.), Transcription Factors: Methods and Protocols, Methods in Molecular Biology, vol. 647, DOI 10.1007/978-1-60761-738-9_1, © Springer Science+Business Media, LLC 2010

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In vivo studies will become increasingly important. We will discuss the tools that are available to investigate these modifications in living animals. 1.1. The Ets Factors

The Ets family is one of the largest families of transcription factors that controls various cellular functions. To date, approximately 30 mammalian genes homologous to Ets-1 have been identified. Ets-1 was the first cellular homolog of the viral oncogene v-ets from the avian transforming retrovirus E26 to be discovered. All Ets transcription factors retain a highly conserved 85 amino acid motif called the Ets domain, which belongs to the superfamily of winged helix-turn-helix DNA-binding domains and recognizes a core GGAA/T sequence, referred to as the Ets-binding site (EBS). The sequences located around the core EBS are variable and define the target gene specificity of individual Ets transcriptional factors. Ets transcription factors are divided into several subfamilies based on homology within the Ets domain. Some subfamilies have the Ets domains at the C-terminal end, and some at the N-terminal end. One-third of Ets transcription factors also contain a conserved N-terminal domain called the Pointed Domain (PD) (see Fig. 1). Several studies indicate that PDs of Ets transcription factors are involved in homo-oligomerization, heterodimerization, and transrepression (for reviews see ref. (1, 2)). Ets transcription factors play essential roles throughout development and adulthood, functioning as downstream effectors of many signal transduction pathways. They are known to regulate the expression of genes involved in various biological processes, including control of cellular proliferation, differentiation, hematopoiesis, apoptosis, metastasis, tissue remodeling, angiogenesis, and transformation (3). Most Ets factors were characterized as either transcriptional activators or repressors, but it has become evident that several Ets factors can function as either activators or repressors, depending upon the type of promoter and/ or Post-translational modifications (2). Various strategies have evolved to regulate transcription factor function and activity, providing the temporal and spatial specificity of the transcriptional output. This specificity is particularly important, as misregulation of the transcriptional response is a fundamental contributor to and consequence of many human diseases, including cancer (4). In this first part of the review, we will consider Post-translational modifications known to regulate Ets factors activity, focusing on phosphorylation, glycosylation, acetylation, ubiquitination, and sumoylation. Then we will discuss specific examples of how such modifications influence their function, more specifically their transcriptional activity, subcellular localization, and stability.

5

A Review of Posttranslational Modifications and Subcellular Localization

Subfamily

M embers

ETS

ETS-1 ETS-2

ERG

ERG FLI-1/ERGB FEV

TAD

ELG

GABPa

TAD

ELF

ELF-1 ELF-2/NERF2 MEF/ELF-4/ELFR

ESE

ESE-1/ESX/ELF-3 ESE-2/ELF-5 ESE-3/EHF

ERF

ERF/PE2 ETV3/PE-1

TEL

TEL/ETV6 TEL-2/ETV7

PEA3

Structure TAD

ND ETS

PD

ETS

PD

ND

TAD

ETS

TAD

ETS

TAD

PD

ETS

ETS

PD

PEA3/E1AF/ETV4 ERM/ETV5 ER71/ETV2 ER81/ETV1

SP I

PU.1/SPI SPIB SPIC

TCF

ELK-1 SAP-1a/ELK-4 NET/SAP-2/ELK-3/ERP

PDEF

PD

RD

TAD

PDE F/SPDE F/PSE

ETS

ND

TAD

ETS

ND

ETS

ETS

PD

RD

RD

TAD

ETS

Fig. 1. The Ets transcription factors family. The 11 subfamilies and their members are presented. One characteristic structure is represented for each subfamily. The boxes correspond to the conserved domains. ETS DNA-binding domain, PD pointed domain, TAD transcriptional activator domain, ND negative domain (negatively control DNA binding), RD repression domain.

2. Regulation of Ets Transcriptional Factors by the Post-translational Modifications

Ets functional activity is modulated at different levels. Ets factors are dependent on interaction with other factors for precise transcriptional regulation. Second, multiple intracellular signaling

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Charlot et al.

pathways and Post-translational modifications directly affect the activity of several Ets proteins by regulating their subcellular localization, DNA-binding activity, transactivation potential, or stability (4). As presented in Table 1, many Ets family members are subject to Post-translational modifications in response to a variety of upstream signals, and these modifications exert a broad spectrum of effects on their activity. One of the best-studied Post-translational modification, phosphorylation, plays a key role in modulating the activity of many proteins, including transcription factors. Phosphorylation occurs by addition of a phosphate group to the hydroxyl group of serine (S), threonine (T), or tyrosine (Y) residues by two families of kinases, S/T protein kinases and Y protein kinases (5). Most Ets phosphorylation occurs via the MAPK (mitogen-activated protein kinase) pathway. In contrast to phosphorylation, glycosylation has been recently investigated as a means of influencing transcription factor activity. Glycosylation of cellular proteins occurs by the addition of the simple monosaccharide O-linked b-N acetylglucosamine (O-GlcNAc) to the hydroxyl group of either S or T residues. Many of the glycosylation sites are identical or closely adjacent to those recognized by S/T protein kinases, suggesting that glycosylation and phosphorylation play competing and antagonistic roles (6). Acetylation, sumoylation, and ubiquitination are Posttranslational modifications that target lysine (K) residues. In addition to phosphorylation, acetylation can regulate Ets gene function. Acetyltransferases, a diverse family of enzymes with the most prominent being p300, transfer an acetyl group to the specific K on the target protein with the reverse reaction mediated by histone deacetylases (HDACs). HDACs recruit a variety of corepressor proteins, and thus are frequently found associated with transcriptional repressors (7). Sumoylation and ubiquitination are also reversible modifications of K residues that affect the stability, activity, and localization of many transcription factors, including those of the Ets family. Ubiquitin and SUMO are both small polypeptides, 9 and 11 kDa, respectively, which are added to a protein by three different enzymes: E1-activating enzymes, E2-conjugating enzymes, and E3 ligases. Ubiquitin and sumoylation-mediated processes have extremely diverse functions with respect to transcriptional regulation. For example, ubiquitination plays key role in regulating transcription factor activity, both indirectly by inducing proteasome-mediated degradation of the protein and directly by altering its transcriptional activity (8). Sumoylation has been shown to affect the stability, activity, and localization of its targets (9). Below, we present several examples of how Ets transcription factors are regulated by Post-translational modifications, to underline the integration of multiple mechanisms to specificity of transcriptional modulation.

A Review of Posttranslational Modifications and Subcellular Localization

7

Table 1 Post-translational modifications and their functional effect on the human Ets transcription factors Modification

Effect

References

Phosphorylation: ERK, JNK, p38

Increases DNA binding and activation

(5, 10–14)

SUMO

Inhibition, nucleo-cytoplasmic shuttling

(13, 14)

Ras-ERK

Converts repressor to activator

(15–17)

JNK

Nuclear export

(18)

SUMO

Increases repression

(19)

Ubiquitination

Degradation

(20)

Sap1a

Phosphorylation ERK

Transcriptional activation

(21)

Tel/ETV6

Phosphorylation ERK

Inhibits repression

(22)

p38

Nuclear export

(23, 24)

SUMO

Nuclear export

(24, 25)

Ubiquitination

Degradation

(26)

MAPK, PKC

Increases activation

(27–29)

MLCK, CaMKII

Autoinhibition, decreases stability, converts to repressor

(30, 31)

Acetylation

Inhibition

(32)

SUMO

Represses transcriptional activity

(33)

Ubiquitination

Degradation

(33)

Ets-2

Phosphorylation MAPK, PKC

Increases activation and protein stability

(28, 34)

Pea3

Phosphorylation MAPK

Increases activation

(35, 36)

SUMO

Inhibition

(36)

Phosphorylation PKA, MAPK

Increases activation

(37, 38)

SUMO

Transactivation

(39)

Ubiquitination

Degradation

(40)

Elk-1

Net/Elk3

Ets-1

Erm/ETV5

Phosphorylation

Phosphorylation

(continued)

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Charlot et al.

Table 1 (continued) Modification

Effect

References

PKA, Msk1, Rsk1

Increases activation

(41, 42)

Mk2

Decreases activation

(43)

SUMO

Transactivation

(36)

Acetylation

Stabilize, alters interactions

(44)

Elf-1

Phosphorylation/ glycosylation

Nuclear translocation, increases activation

(45)

Mef

Phosphorylation CyclinA

Decreases activation

(46)

Cdk2

Ubiquitination/degradation

(47)

SUMO

Transactivation

(48)

Phosphorylation

Unknown

(49)

SUMO

Unknown

(50)

Erg

Phosphorylation PKC

Unknown

(51)

Pu.1/Spi1

Phosphorylation CKII, p38

Increases activation and alters protein interactions

(52–54)

Spi-B/Spi2

Phosphorylation CKII, MAPK

Increases activation and alters protein interactions

(54, 55)

Erf

Phosphorylation MAPK

Nuclear export and loss of repression

(56–58)

GABPa/ELG

Phosphorylation ERK, JNK

Increases activation and stabilize protein complex formation

(59–61)

Er81/ETV1

Fli-1

Phosphorylation

2.1. Cooperation of Phosphorylation and Sumoylation as Mechanisms of Downregulation of Tel Repressor Function

Tel (Translocation Ets Leukemia) is one of the best-characterized transcriptional repressor within the Ets superfamily. It plays an important role in the development and maintenance of vasculature and for adult hematopoiesis, and is frequently rearranged or deleted in several cancers. Tel is regulated by specific MAPKmediated phosphorylation that leads to removal of its transcriptional repressive activity and induction of nuclear export. ERK (Extracellular signal-Regulated Kinase) and p38 kinases phosphorylate Tel removing its transcriptional repression by decreasing its DNA-binding ability (22, 23). In addition to being regulated by phosphorylation, Tel is also sumoylated. The E2 SUMO-conjugating enzyme UBC9 (Ubiquitin-like protein SUMO-1 conjugating enzyme) interacts with the PD of Tel.

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9

SUMO-modified Tel localizes to nuclear bodies termed Tel-bodies, which are transient structures formed during S phase and are disrupted during the rest of the cell cycle (62). Mutated K99R Tel, which cannot be sumoylated, cannot be exported from the nucleus or localized to Tel-bodies, and functions as a better transcriptional repressor than wild-type Tel (62). These results suggest that SUMO modification contributes to the abrogation of transcriptional repression and nuclear export of Tel (24). 2.2. Antagonism Between Phosphorylation and Sumoylation Regulates Elk-1 Activity

Elk-1 belongs to the ternary complex factor (TCF) subfamily of the Ets transcription factors. Their characteristic property is the ability to form a ternary nucleoprotein complex with the serum response factor over the serum response element on the regulated promoter (1). In vitro and cell culture studies have demonstrated that Elk-1 functions as a transcriptional activator, and is regulated by phosphorylation and sumoylation (13). Members of all three MAPK subgroups, ERK, JNK, and p38, phosphorylate Elk-1 at several sites within the transactivation domain (TAD). Multiple phosphorylation events on Elk-1 lead to conformational changes that alter intramolecular interactions between the Ets domain and the TAD, resulting in increased DNA binding and transcriptional activation (for review see ref. (12)). When the MAPK pathway is not activated, both the Ets domain and an inhibitory domain, called the R motif, recruit corepressors and suppress the activity of the Elk-1 TAD, maintaining the TCF in an inactive state. Several SUMO consensus sites have been identified within the R motif. Blocking sumoylation, using different strategies, increases Elk-1 transcriptional activity in the absence of MAPK activation, suggesting that sumoylation plays a role in repressing the basal level of the Elk-1 transcriptional activity. Simultaneous activation of the ERK pathway and inhibition of sumoylation produce a synergistic increase in transcriptional response, indicating that the ERK and SUMO pathways function antagonistically to control Elk-1 transactivation potential. Thus MAPK-mediated phosphorylation of Elk-1 both directly and indirectly enhances transcriptional activation, by potentiating activity of the TAD and by inhibiting sumoylation of the R motif, respectively (13). In addition, sumoylation has been shown to influence the nucleo-cytoplasmic transport of Elk-1, thereby regulating its nuclear retention and potentially affecting transcriptional output (13, 14).

2.3. Diverse Posttranslational Modifications Regulate the Dual Role of Net as a Repressor and/or Activator

The transcriptional factor Net (New Ets factor; also called Elk-3, SAP2, and ERP) is another member of the TCF subfamily. It is regulated by mechanisms involving complex patterns of multisite Post-translational modifications that influence DNA-binding ability, protein–protein interactions, subcellular localization, stability, and transcriptional activation and/or repression. Under

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basal serum conditions, Net, in contrast to the other TCF members Elk-1 and Sap-1a, is a strong repressor of transcription. Two autonomous repression domains of Net, the NID and the CID, mediate repression. Interestingly, Net is rapidly converted to an activator by the growth factor/Ras/MAPK pathway induced phosphorylation of its TAD (15). ERK and p38 bind to the D box of Net and binding is required for phosphorylation of the adjacent TAD (16). Net is regulated by nuclear-cytoplasmic shuttling in response to specific signaling pathways. Net is mainly nuclear under both normal and basal serum conditions. Net contains two nuclear localization signals and one nuclear export signal in the conserved Ets DNA-binding domain. Consequently, Net is exported from the nucleus in response to stress stimuli transduced through the JNK pathway, leading to relief from repression. JNK binds to the J box in the middle of the protein, and binding is required for phosphorylation of the adjacent export motif. Nuclear exclusion relieves transcriptional repression by Net (18). In conclusion, Ras signaling and JNK phosphorylation are crucial factors which regulate Net function as transcription activator or repressor and connect two important pathways involved in cell transformation. Net transcriptional repression is regulated by the NID domain and involves sumoylation. Net sumoylation increases its repression and decreases the positive activity of Net (19). Another mechanism of regulation of Net transcriptional repressor function involves hypoxia induces Net ubiquitination, and proteasomal degradation (20). The effect of hypoxia on Net function will be discussed in detail below. 2.4. Multiple Phosphorylations and Acetylation Regulates Er81 Activity

The transcriptional activator Er81 (Ets relative protein 81) is another example of how coordinated and/or antagonistic phosphorylation, acetylation, and ubiquitin-mediated degradation modulate protein activity. Er81 transcriptional activation is enhanced by phosphorylation at multiple sites in response to signaling downstream of the HER2/Neu RTK (Human Epidermal Growth factor Receptor 2) by ERK and p38 MAPK, and also by a MAPK-stimulated protein kinase, Msk1 (or Rsk1) (41–43). In contrast, protein kinase A (PKA) recognizes sequences similar to those recognized by Msk1 and phosphorylates Er81, resulting in a reduction of its DNA-binding ability but also an increase of its transcriptional activation (37). Decreased DNA binding could prevent activation of low-affinity promoters, but have no effect on those with high affinity. Thus changing DNA affinity may be an important strategy for determining target specificity. Er81 is also negatively regulated by phosphorylation. MAPK-activated protein kinase 2 (Mk2), phosphorylates Er81 on its inhibitory domain and suppresses basal transcriptional activity (43).

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A second consequence of Her2/Neu signaling is acetylation of Er81 in its TAD. Er81 is acetylated by two coactivators/acetyltransferases, p300 and p300- and CBP-associated factor (P/CAF) in vitro and in vivo (44). Acetylation of Er81 not only enhances its transactivation, but also increases its DNA-binding activity and in vivo half-life. Furthermore, the oncogenic HER2/Neu, which induces phosphorylation and thereby activation of Er81, is less able to activate acetylation-deficient Er81 mutants, indicating that both acetyltransferase and protein kinase-specific regulatory mechanisms control Er81 activity (44). Thus Her2/Neu signaling activates Er81 at multiple levels, which results in context-specific differential expression of target genes. 2.5. Synergism Between Phosphorylation and Glycosylation Regulates Elf-1 Activity, Subcellular Localization, and Degradation

Elf-1 (E74 like factor-1) is one of the few proteins known to be phosphorylated and glycosylated at the same time. Elf-1 is involved in the transcriptional regulation of many hematopoietic cell genes. Several studies of Elf-1 reveal that differential phosphorylation and glycosylation regulate subcellular localization, protein–protein interactions and protein–DNA interactions. Elf-1 exists primarily as a 98-kDa form in the nucleus and as an 80-kDa form in the cytoplasm. Phosphorylation by PKC and O-linked glycosylation contribute to the increased Post-translational molecular mass of Elf-1. The 98-kDa Elf-1 is released from the cytoplasm, dissociates from its cytosolic binding partner retinoblastoma protein and moves to the nucleus, where it binds to a target promoter. Both modifications are required for maximal activation of the promoter, indicating that they target distinct residues and function cooperatively. Interestingly, the cytoplasmic 98-kDa form enters the proteasome pathway and undergoes degradation. In conclusion, different Post-translational modifications, glycosylation and phosphorylation, cooperatively influence Elf-1 transcription factor activity (45).

2.6. Phosphorylation Tightly Regulates Fli-1 and Mef Half-Life and Activity

Phosphorylation of Ets factors, including Fli-1 and Mef, appears to precisely regulate their function in a short time frame, simultaneously determining protein half-life and activity. The Fli-1 (Friend leukemia virus integration-1) transcription factor is involved in the regulation of several developmental processes and becomes oncogenic when overexpressed or mutated. Fli-1 is a short-lived phosphoprotein in the human T cell line Jurkat. Fli-1 is expressed as two isoforms, p51 and p48, which are both phosphorylated. Interestingly, Fli-1 phosphorylation increases by a Ca(2+)-mediated process, but it is not stimulated by protein kinase C activation. The p51 isoform has a half-life of 105 min, and p48 80 min. In addition, newly synthesized Fli-1 rapidly decreases during human T cell activation (49). Another example of a short-lived phosphoprotein is the Ets transcription factor Mef (Myeloid Elf-1 like factor), which acts as

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a strong transcriptional activator of cytokine gene expression and plays an important role in hematopoietic stem cell behavior and normal development of NK T cells. Interestingly, the transcriptional activity of Mef is largely restricted to the G1 phase of the cell cycle. Mef expression peaks during late G1 phase. Mefdependent transcription is suppressed by Cyclin A-mediated phosphorylation (46). The rapid turnover of Mef in S phase is dependent on specific phosphorylation of its C-terminus by cdk2 and on ubiquitination and proteolysis by SCFSkp2 (a Skp1/ Cul1/F-box (SCF) E3 ubiquitin ligase complex) (47). The tight regulation of Mef levels by phosphorylation during the cell cycle establishes a novel link between the Ets family of proteins and the cell cycle machinery. 2.7. Sumoylation and Ubiquitination Negatively Regulate the Activity of Ets-1, Pea3, and Erm

Several recent reports revealed the role of sumoylation and ubiquitination in the regulation of Ets-1, Pea3 (Polyomavirus enhancer activator 3), and Erm (Ets-related molecule) Ets transcriptional factors. Sumoylation of Ets-1 by Ubc9 and PIASy on two K sites in the Synergistic Control motifs leads to reduced transactivation, but does not affect its stability. Ets-1 is modified by K48-linked polyubiquitinylation independently of the sumoylation acceptor sites and is consequently degraded through the 26S proteasome pathway (33). These data indicate that Ets-1 can be modified by sumoylation and/or ubiquitinylation, with sumoylation repressing the transcriptional activity of Ets-1 and having no clear antagonistic action on the ubiquitin-proteasome degradation pathway. Two independent publications recently demonstrated that the transactivation function of Pea3 and Erm is regulated by sumoylation. Erm and Pea3 belong to the PEA3 group of Ets transcription factors (36, 39). They are involved in many developmental processes and are transcriptional regulators in metastasis. The PEA3 group members have similar N-terminal TADs whose activity is inhibited by the negative regulatory domain (NRD). For Erm, the NRD is a SUMO dependant repression domain. In addition, SUMO sites outside the NRD also play a role in the negative regulation (39). There are similar effects of sumoylation on transactivation of Pea3 (36). Collectively, these observations suggest that the activity of Pea3 and its paralog Erm are negatively regulated by sumoylation. Interestingly, Erm is almost undetectable in a variety of human breast cancer cell lines, suggesting that it is rapidly degraded. Endogenous and ectopically expressed Erm are shortlived. Erm is ubiquitylated on its C-terminal region and is degraded via the 26S proteasome pathway. This mechanism plays an important role in the regulation of Erm transcriptional activity (40).

A Review of Posttranslational Modifications and Subcellular Localization

3. Ets Posttranslational Modifications in Cancer

The Ets family is implicated in the regulation of genes involved in cell proliferation, differentiation, transformation, and apoptosis. The control of such important biological processes put the Ets factors in a key position in normal cell homeostasis and in mechanisms of disease. Ets factors are overexpressed in different diseases; for example, Ets-2 is overexpressed in Down syndrome (63, 64) and in rheumatoid arthritis (65). Post-translational modifications of Ets factors are altered in different human pathologies (Table 2). Strikingly, deregulation of Ets proteins is frequently observed in human cancers. We will describe some examples of Ets deregulation and their involvements in various diseases, in particular cancer.

3.1. Ets Posttranslational Modifications in Human Diseases

3.2. Ets Factor Deregulation and Post-translational Modification in Cancer

Cancer is the result of several genetic alterations leading to deregulation of normal cell physiology, escape from apoptosis or loss of growth control (3) (72). Cancer cells share common features that have been listed in six main sets of functional capabilities (73): self sufficiency in growth signals, insensitivity to growth arrest signals, escape from apoptosis, unlimited replicative potential, angiogenesis (new blood vessels formation), promotion of tissue invasion and metastasis.

3.2.1. Ets Deregulation

Table 2 Involvement of Ets Post-translational modifications in human diseases Modifications

Ets factors

Disease

References

Phosphorylation

Elf-1

Lupus

(66)

Elk-1

Cervical cancer (HeLa)

(67)

Long term behaviors to THC

(68)

Addictive properties of cocaine

(69)

Ets-2

Down syndrome Leukemia

(64)

Net

Cervical cancer

(70)

Kaposi sarcoma Head and neck squamous cell carcinoma Prostate cancer

(71)

Ets-1/Ets-2

Invasive breast cancer

(29)

Er81

Neoplasia

(44)

Ets-1

Arthritis, cancer

(32)

Glycosylation

Elf-1

Lupus

(66)

Sumoylation

TEL/AML

Leukemia

(62)

Acetylation

13

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Cancer is responsible for approximately 13% of mortality in the world and this percentage is continually rising. Increased knowledge will provide earlier detection, more efficient treatment, and prevention strategies. Interestingly Ets factors play a key role in this pathology. Ets transcription factors can activate or repress genes implicated in differentiation, proliferation, transformation, angiogenesis, or apoptosis (74), biological processes that are implicated in the development of cancer. The first clue for the role of Ets factors in cancer came with the discovery of the avian retrovirus E26 oncogene v-ets (2). v-ets induces erythroid-myeloid and lymphoid leukemia in mice. Protooncogenes are activated by overexpression, point mutation, activation by insertions of new regulatory sequences (for complete review see ref. (3)). In most cases, Ets factors are amplified or overexpressed. For example, high levels of Ets-1 have been found in thyroid cancer cells (75), and in many invasive and metastatic solid tumors. However, Net, Ets-1, and Fli-1 are expressed at a low level in breast cancer cells (both at the mRNA and protein level), suggesting that they can act as suppressors during mammary tumorigenesis (76). Interestingly, there is additional evidence for tumor suppressor activity for Net, in cervical and pancreatic cancer cells. The cervical cancer study (70) used a model generated by the fusion of HeLa and IMR90 cells. The fusion gave two HPV positive cell lines: the nontumorigenic line (444) and the spontaneous revertant tumorigenic line (CGL3) (77). In this model, increased tumorigenicity is associated with decreased Net expression and increased expression of the protooncogene c-fos, which is involved in cell proliferation and other processes (78). In pancreatic cancer cells, overexpression of Net leads to inhibition of proliferation (79). Net is expressed at a low level in tumor tissues and at a high level in normal pancreatic cells. Net may have a physiological role in controlling the expression of the immediate early gene c-fos in pancreatic cells. 3.2.2. Ets Posttranslational Modifications

Ets factors are affected by phosphorylation, sumoylation, acetylation, glycosylation, and ubiquitination (4). A number of studies describe specific links between Ets Post-translational modifications and cancer.

3.2.3. Phosphorylation

The MAPK family is represented by three kinases subfamily: ERK, JNK, and p38. JNK and p38 are mainly activated by stress signals and ERK respond to mitogenic signals. Several oncogene products are upstream effectors of the ERK signaling pathway, including Ras (80). Ras gene mutations are found in multiple tumor types. The Ras signaling pathway induces phosphorylation of Net and other factors (71, 81). Under basal conditions, Net is a strong repressor of transcription and it is converted to an activator when it is phosphorylated in its TAD. Net is involved in angiogenesis (43). Mutated oncogenic Ras elevates expression

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of VEGF (vascular endothelial growth factor), stimulates angiogenesis and leads to the promotion of tumor growth and metastasis. Phosphorylated Net is coexpressed with VEGF in human tumors (head and neck squamous cell carcinoma and prostate cancer) and both are highly expressed in transformed cells, highlighting the fact that phosphorylated Net could be a positive regulator of angiogenesis through effects on VEGF. In the cervical carcinoma 444/CGL3 cell model (82), we have evidence that Net phosphorylation is cell-type specific (unpublished data). The nontumorigenic cells (444) have a high level of nonphosphorylated Net that acts as a powerful repressor of transcription. In contrast, in the tumorigenic segregants (CGL3) Net is expressed at a lower level but is mainly phosphorylated. Phosphorylation of Net may promote tumorigenesis in cervical cancer cells. The related ternary complex factor, Elk-1, has recently been shown to be phosphorylated in a model of arsenic-transformed prostate epithelial cells (83). Ets factors are also phosphorylated by Protein Kinase C (PKC), which form a family that can be divided between conventional, novel and atypical types. The conventional PKCs are composed of several isoforms (a, b1, b2, and g), and require calcium, diacylglycerol, and phospholipids to be activated. The novel PKCs need diacylglycerol for activity, whereas atypical PKCs are independent of calcium and diacylglycerol. PKCa phosphorylates Ets-1 without calcium mobilization. Phosphorylation occurs on the exon IV domain that has four potential phosphorylatable serines. PKC dependent phosphorylation could contribute to invasion by breast cancer cells (29). 3.2.4. Acetylation

Ets factor acetylation in cancer is poorly investigated, but has been reported for Ets-1 and Er81. Ets-1 acetylation is stimulated by the transforming growth factor b (TGFb) pathway (32). TGFb can induce prolonged acetylation of Ets-1, leading to increased matrix degradation that promotes tumorigenesis. Er81 acetylation appears to have a pleiotropic effect in tumor formation (44). Expression of Er81 is necessary for normal development, especially of the spinal cord, whereas deregulation of Er81 expression is linked to neoplasia. Er81 expression is elevated in some human breast tumor cell lines. Interestingly, Er81 acetylation increases its transactivation, DNA-binding activity, and halflife. Er81 interacts with two acetyltransferases: p300- and p300/ CBP-associated factor. Interestingly, Er81 regulation by acetylation and phosphorylation is linked. The HER2/Neu pathway is activated by gene amplification in some breast carcinomas. Several studies have shown that the HER2/Neu-Ras-MAPK pathway phosphorylates and activates Er81. In addition, this pathway promotes acetylation by p300, which further stimulates Er81 transactivation (44).

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3.2.5. Sumoylation and Subcellular Transport

Sumoylation with SUMO-1 of TEL in the TEL/AML1 fusion protein (leukemia-associated fusion protein TEL/AML1) has been reported to have a role in leukemogenesis (62). Tel is located in the nucleus in almost all tissues, where it acts as a transcriptional repressor by inhibiting gene expression through the histone-deacetylase pathway. Tel interacts with UBC9, leading to SUMO-1 modifications of Tel and localization to Tel-bodies. Acute Myeloblastic Leukemia (AML) is associated with the translocation t (12, 21) that fuses TEL to AML1 (gene cbfa2). The TEL/AML1 fusion protein is also modified by SUMO-1 and located in the Tel-bodies. The hypothesis is that SUMO Posttranslational modification of TEL/AML1 could lead to abnormal localization of the fusion protein that may contribute to leukemogenesis (62).

3.3. Ets Factors as Targets for Cancer Diagnosis and Therapy

Aberrant expression and modification of Ets transcriptional factors is observed in numerous cancers, which could be exploited to develop markers and targeted treatments. Overexpression appears to be frequent because many transcription factors are inactive under normal physiological conditions (84). Overexpression of Erm and Er81 (PEA3 group) is linked with breast cancer (85). Elevated levels of Ets-1 correlate with the degree of malignancy of breast and lung cancers (86, 87). Multiple strategies have already been used to decrease expression levels, including RNA interference, antisense oligonucleotides, and negative mutants (for a general review see ref. (60)). Most recently, antisense oligonucleotides targeting Elk-1 have been shown to suppress tumorigenicity of human hepatocellular carcinoma cells (17, 88). Post-translational modification of Net has been used as a target to isolate an antineoplastic agents (17). XRP44X was identified in a large-scale screen for small-molecule inhibitors of Ras-activated Net transcriptional activation. XRP44X is a pyrazole, a chemical family that is used conventionally as analgesic, anti-inflammatory, or antipyretic drugs. XRP44X inhibits FGF2induced Net phosphorylation by the Ras-ERK signaling pathway. Interestingly, XRP44X also induces depolymerization of microtubules and affects the morphology of the actin skeleton. It shares this property with combretastatin A-4, an activated metaboliteprodrug derived from an African bush willow that is used for its tumor vascular-targeting activity. These classes of therapeutical agents differ from others that target microtubules and who have little (vincristine) or no (nocodazole, docetaxel) effect on the Ras-Net pathway. Another exciting feature of XRP44X is that it inhibits the growth of transformed cells in culture and angiogenesis in an ex vivo assay on endothelial cell sprouting. Our study identified a novel inhibitor of the Ras-ERK pathway that inhibits efficiently Net activation via inhibition of Net phosphorylation,

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and also act as inhibitor for several genes induced by the Ras-ERK pathway as c-fos or egr-1. We propose that XRP44X is a groundbreaking class of drug which combines two anticancer mechanisms, by acting as an antimitotic drug and by inhibiting a signaling pathway.

4. Involvement of Ets Factors in the Hypoxic Response

4.1. Transcriptional Regulation of Ets Factors in Hypoxia

In order to efficiently exploit Ets factors for cancer therapy, we need to understand better their role in tumor progression. Most solid tumors are sensitive to the properties of their microenvironment, including the availability of nutrients and oxygen. Decreased oxygen tension, so called hypoxia, stimulates cell migration and angiogenesis, in order to provide oxygen and nutrients required for tumor growth. The regulation of Ets factors by hypoxia is a growing area of interest. Hypoxia has many effects on cells, of which induction of the HIF-1a transcription factor (or its paralog HIF-2a) is considered to be of particular importance, and is extensively studied in many laboratories. In the presence of oxygen, HIF-1a is hydroxylated by the Prolyl Hydroxylase Domain proteins (PHDs). Hydroxylated HIF-1a binds to the tumor suppressor Von Hippel–Lindau (VHL) protein that targets HIF-1a for proteasomal degradation. In the absence of oxygen, HIF-1a is not degraded and translocates into the cell nucleus where it heterodimerises with HIF-1b and binds to the DNA on hypoxia-response elements (HREs) of hypoxia-response genes (89). Hypoxia induces genes involved in angiogenesis, cell migration, cell growth, cell proliferation, cell invasion, and apoptosis (for reviews see ref. (89–91)). Interestingly, the Ets transcription factors are known to regulate genes involved in these processes (3), suggesting that Ets factors could play a role in hypoxic conditions. We will review recent studies that highlight the emerging role of Ets transcription factors in the hypoxic response. One of the first indications for the involvement of Ets in the hypoxic response was the study of Ets-1 expression under hypoxic conditions (92). In a human bladder cancer cell line atmospheric hypoxia induces Ets-1 gene transcription. The Ets-1 promoter contains one consensus hypoxia-responsive element that binds HIF-1, suggesting that hypoxia induces Ets-1 expression via HIF-1. Hypoxia and HIF-1 have also been shown to decrease the expression of Elk-1 in arterial endothelial cells (93). Hypoxia regulation is not always HIF1 dependent. In hepatoma cell lines, the expression of Ets-1 is strongly increased in hypoxia, but this induction is not affected when HIF-1a is repressed (94). Similarly, expression of Nerf2

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(New Ets-related factor 2), which is involved in blood vessel development, is increased in a HIF-1a independent manner under hypoxia in human umbilical vein endothelial cells (95). Angiopoietin-1 protein, a growth factor that promotes angiogenesis, regulates this induction. Changes in Ets protein levels affect the hypoxic response in various ways. 4.2. Cooperation of Ets Factors and HIF in Hypoxic Induction of Target Genes

HIF-1 and HIF-2 have been shown to interact with various transcription factors. A recent report shows that cooperation between HIF-2a and Elk-1 is required to activate specific target genes, such as Epo and Pai-1 (96). Interestingly, ten genes that were found to be regulated by HIF-2, but not HIF-1, have at least one HRE in proximity to an EBS on their promoter (97). HIF-2a and Elk-1 have been shown to physically interact and cooperate in the hypoxic induction of three genes: Cited-2, Wisp2, and Igfbp3. HIF-2a has also been shown to interact with Ets-1 in various studies. For instance, the Flk-1 (vascular endothelial growth factor receptor-2) promoter contains two functional binding sites for HIF-2a, located in close proximity to functional EBS in the proximal region of the promoter. HIF-2a physically interacts with Ets-1 and this cooperation activates the transcription of Flk-1 that is indispensable for angiogenesis (98). Endosialin is a transmembrane glycoprotein that is induced in hypoxia and is implicated in angiogenesis. The endosialin promoter contains two distinct regions: a distal region that contains a functional HRE site and an adjacent Ets-1-binding site, and a proximal region with two EBS but without HRE site. In the distal region, HIF-2a and Ets-1 bind to their respective recognition sites and interact, thereby contributing to the hypoxic induction of the endosialin gene. The proximal region of the promoter is also sensitive to hypoxia despite the absence of HREs. The two EBS elements bind Ets-1, which in turn recruits and interacts with HIF-2a, allowing the hypoxic induction of the endosialin gene (99). Interestingly, although the VE-cadherin gene is not sensitive to hypoxia, HIF-2a and Ets-1 specifically activate the promoter in synergy (100). The Vegfr1 promoter is regulated by functional interactions of Ets-1 and HIF-2. They alter nucleoprotein structure by interacting with the transcriptional coactivators CBP/p300, leading to the recruitment of Pol II and transcriptional induction in endothelial cells (101). Ets-1 is not specific for HIF-2a, and can also act with HIF1a. Ets-1 cooperates with HIF-1a during the density-dependent upregulation of the hypoxia-inducible gene Ndrg1 (102). Ets-1 and HIF-1a bind to their closely located binding sites on this promoter, interact together, and cooperate in regulating its transcription. However, this interaction is not obligatory, since in the absence of HIF-1a or HIF-2a, Ets-1 alone is sufficient for the upregulation of hypoxia-inducible genes.

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These data raise important questions about the importance of Ets factors in the hypoxic response, relative to the “master” regulator, the HIFs, that have been mostly studied. 4.3. Involvement of Ets Factors in the Hypoxic Induction of Target Genes

A recent study with a breast cancer cell line has shown that 83% of the hypoxia-responsive genes are regulated by HIF-1a, 4% by HIF-2a, and 6% by both factors. Seven percent of hypoxiaresponsive genes are independent of HIF-1a and HIF-2a (97). Supporting this model, another study has shown that HIF-1a modulates only about half of the hypoxia-induced genes in arterial endothelial cells (93). These results suggest that other transcription factors, such as Ets transcription factors, might be involved in regulating hypoxia-inducible genes. HIF transcription factors undergo Post-translational modifications, such as hydroxylation, polyubiquitination, acetylation, phosphorylation, sumoylation, or S-nitrosylation (103) that are important for their regulation and function in the hypoxic response. Interestingly, two TCF subfamily members, Elk-1 and Net, have been shown to be regulated by protein modification in hypoxia, but in distinct ways. Elk-1 is phosphorylated in response to hypoxia via the MAPK pathway, leading to the hypoxic induction (without HIF involvement) of target genes such as c-fos (67) and Egr-1 (104). In contrast, hypoxia enhances Net ubiquitination, nuclear export, and subsequent proteasomal degradation. Phosphorylation by ERK is not necessary for the decrease of Net levels in response to hypoxia. Furthermore, the level of the two TCFs, Elk-1 and Sap-1a remained stable under hypoxia, suggesting that the downregulation in hypoxia is specific for Net. Net degradation leads to loss of Net repression and induction of c-fos, Egr-1, and Pai-1, genes that are known to be involved in migration, proliferation, or angiogenesis (20). In a larger-scale analysis, we identified 78 other genes whose hypoxic induction is dependent on Net but not HIF-1a (105). These genes are mainly involved in cell cycle, cancer, and cell-to-cell signaling. In summary, under hypoxic conditions some Ets factors are essential for the induction of genes that respond to hypoxia, with or without HIF involvement. The hypoxic regulation of Ets factors, at transcriptional or Post-translational level, also contributes to the hypoxic response (see Fig. 2).

4.4. The Ets Factor Net is a Regulator of HIF-1a Protein Stability

We have compared the HIF-1 and Net pathways that are activated by hypoxia. Surprisingly, we found that Net participates in the transcriptional hypoxic regulation of a large set of genes, of which three are known to be important for HIF-1a protein stability, PHD2, PHD3, and Siah2 (105). Net appears to be an indirect modulator of HIF-1a protein stability, suggesting that it has a key role in the hypoxic response. All these studies highlight the close link between hypoxia-inducible factors and Ets transcription

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Hypoxia Activation

H IF -1a

H IF -2a

Elk-1

Net

Ets-1

Cooperation

Hypoxic target฀genes

Degradation

Ets-1

N drg1

Elk-1

F lk-1 Endosialin Vegfr1

Ep o Pai-1 Cited-2 Wisp 2 Igfbp 3

C-fos Egr-1

C-fos Egr-1 Pai-1 …

Fig. 2. Ets factors in the hypoxic induction of target genes. Hypoxia activates HIF transcription factors which in turn cooperate with Ets-1 or Elk-1 to induce target genes. Hypoxia also activates Elk-1 (phosphorylation by MAPK) leading to induction of two target genes. Hypoxia enhances Net degradation, leading to the loss of the repression and induction of target genes.

factors under hypoxic conditions, suggesting that Ets factors are involved in responses to oxygen in the tumor environment during tumorigenesis.

5. Studying Ets Factors Posttranslational Modifications in Living Animals

5.1. Strategies for Studies of Ets Factors Phosphorylation in Mouse Cancer Models

Studies of Post-translational modifications of Ets transcription factors have been performed in in vitro and in various cell lines. We reviewed their involvement in human diseases and especially in cancer and in biological processes important for these diseases. These different studies can help us to understand fundamental molecular and cellular mechanisms, but investigating Ets protein modifications in well-characterized living animal models remains essential. Here we will review studies of Post-translational modifications of Ets factors in different models: mice, rats, fruit flies, and worms. Post-translational modifications of Ets factors are poorly documented at the level of the animal. Several studies compare null mutants with in vitro studies to deduce the roles of Ets protein modification. However, some interesting publications have

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21

used more appropriate tools to study Ets Post-translational modifications: –

Alleles specifically mutated at modification sites

–

Imaging technology

–

Antibody specifically targeting modified Ets sites

5.1.1. Ets-2 Phosphorylation: Mutation of the Modification Site

The importance of Ets-2 activation by MAPK phosphorylation in mammary tumor development has been investigated with a specific mutated allele of Ets-2 (Ets-2A72), in which Thr-72 is substituted by an Ala residue (106). In the Ets-2A72 knock-in mouse line, mRNA expression and protein stability are not affected, suggesting that the observed decrease in Ets-2 activity is due to the absence of Thr-72 residue. Moreover, adult Ets-2A72/A72 animals do not have abnormalities in fertility, longevity or mammary gland development. Two mouse models of mammary tumorigenesis were used, MMTV-pyMT and MMTV-NeuYD, in which the MMTV (mouse mammary tumor virus) promoter is linked to the polyomavirus Middle T oncogene (pyMT) or the Neu protooncogene mutated allele (the Neu EGFR family member is implicated in human breast cancer). In both models, Ets-2A72 mutation restricts development of diverse mammary tumors, showing that Ets-2 phosphorylation is important for tumor development. The role of Ets-2 in blood vessel formation was investigated with a vascularization mutant of VEGF, VEGF-25. An Ets-2A72/VEGF25/NeuYD mouse line does not have altered tumor vascularization, suggesting that Ets-2 acts downstream of VEGF.

5.1.2. Elk-1 Phosphorylation: Imaging with Bioluminescence

Another interesting and innovative study in cancer research used imaging in the context of xenograft models of prostate cancer (107). The technique uses a modified version of two-step transcriptional amplification (TSTA). In this system, the GAL4 DNAbinding domain is fused to the activation domain of Elk-1, leading to the expression of a GAL4-Elk-1 element. When the Elk-1 domain is activated by phosphorylation, the GAL4 DNA-binding domain is able to interact with a GAL4-responsive reporter gene that generates high levels of firefly luciferase (Fluc), whose bioluminescent activity can be detected. The GAL4-Elk-1 element is under the control of a modified prostate-specific antigen (PSA) promoter, which responds strongly to the androgen receptor (AR). These constructs were inserted in a replication-defective adenovirus (adTSTA-Elk-1) that can be used in xenographts. This system was used to detect EGF-activated MAPK in two prostate cancer xenographts generated with CWR22-AI (an androgen-independent prostatic-carcinoma-derived cell line that expresses PSA and mutant AR), and LAPC9 (an androgen-dependent bone-metastasis-derived cell line that expresses PSA and wild-type AR). adTSTA-Elk-1-mediated luciferase expression was

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found to be EGF inducible in the androgen-dependent and independent xenographts models. Interestingly, MAPK activity and Elk-1 phosphorylation were found to be greater in the CWR22 tumors than in the LAPC9 tumors in vivo. 5.1.3. Net Phosphorylation: Antibodies Designed to Target Modification

It is also possible to detect the phosphorylated and nonphosphorylated forms of Ets proteins with appropriate antibodies designed to specifically bind to phosphorylated or nonphosphorylated sites of the protein. In the study of the in vivo roles of Net, we have demonstrated in wound healing assays that activated phosphorylated Net (P-Net) and VEGF are coexpressed in the angiogenic process (71). Furthermore, Net downregulation inhibits angiogenesis and VEGF expression in vivo, whereas P-Net stimulates the mouse VEGF promoter in cells. This study illustrates the role of a phosphorylated form of an Ets factor in a fundamental aspect of tumor progression, by combining in vitro, cell line, and in vivo studies, with appropriate Net and P-Net antibodies. The role of Ets factors is not restricted to tumorigenesis, as they have been shown to be involved in other functions including immunity, or neuronal process.

5.2. Ets-2 Phosphorylation in Macrophages in Mouse

Several Ets transcription factors are expressed in the immune system, including Ets-1, Sap-1a, and Fli-1 in T cells and Ets-2 in macrophages, which are key elements of both innate and adaptive immune responses. The biological role of Ets-2 phosphorylation on Thr-72 has been studied in macrophages and the inflammatory response using the mouse line with the Ets-2A72 allele (see above) (108). Ets-2 is constitutively phosphorylated on Thr-72 in a model of acute inflammation generated with the hemopoietic cell phosphatase (Hcph) allele, Hcphme-v. Crosses between mice with the mutant alleles were used to show that Ets-2 phosphorylation has a positive role in the severe inflammatory response of the me-v model, by mediating macrophage survival and expression of inflammatory genes such as TNF-a, CCL-3, MMP-9 in macrophages. Since metalloproteases and macrophages are involved in cancer development and in VEGF expression, MMP-9 and MMP-3 expression were also studied in macrophages (106). A dramatic deficiency in the context of Ets-2A72 indicates that Ets-2 phosphorylation at Thr-72 is essential for MMP-9 and MMP-3 expression. In conclusion, mammary cancer progression could be due in part to the regulation of macrophage metalloproteases via this Ets-2 modification.

5.3. Elk-1 Phosphorylation in Neurons in the Rat Model

The most studied Ets factor for Post-translational modifications in animals is probably the ternary complex factor Elk-1 in brain and neuronal processes. They rely on a specific antibody that targets Elk-1 phosphorylated on Ser-383.

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In adult rat striatum, Elk-1 is nuclear and cytoplasmic, and phosphorylated Elk-1 (P-Elk-1), activated by electrostimulation, is localized in the nucleus, cytoplasm, and dendrites (109). Erk phosphorylation is involved in glutamate signaling in the rat striatum in neuronal cells in vivo (110). P-Elk-1 (targeted with P-ElkSer-383 antibody) was investigated in striatal slices prepared from adult rat brains stimulated by superfused solutions containing the excitatory neurotransmitter glutamate. P-Elk-1 levels were found to be increased by glutamate treatment. Slices treated with glutamate and the ERK inhibitor PD98059 did not exhibit increased P-Elk-1. Elk-1 and ERK-1/2 phosphorylation in the striatum is also increased by stimulation of group I mGluRs (metabotropic glutamate receptors) (111). Protein phosphatase 1/2A (PP1/2A) regulates mGluRs activity by dephosphorylating Elk-1: injection of okadaic acid (a PP1/2A inhibitor), and DHPG (a GluRs I agonist), lead to increased P-Elk-1 (112). These studies also help us to understand how drugs act on brain. Amphetamine injection leads to an increase in P-ERK and P-Elk-1 in rat striatum via the group I mGluRs (113). Cocaine injection (69) and THC administration (68) in mice results in Elk-1 hyperphosphorylation by ERK signaling. Nicotine modulates Elk-1 in the rat hippocampus in a spatially and temporally specific manner. In vivo acute nicotine activates Elk-1 in the CA1 area but not in the dentate gyrus. Chronic nicotine for 14 days changes the level of total Elk-1 but not its phosphorylation state. Thus, Elk-1 regulation of transcriptional events may contribute to nicotine-induced changes in the hippocampus (114). Interestingly, unfamiliar taste induces Elk-1 phosphorylation via P-ERK, whereas familiar taste has no incidence (115). These studies help us to understand mechanisms leading to the phosphorylation and dephosphorylation of the TCF Elk-1 in neurons and the role of these modifications in drug or taste habit-formation. 5.4. Ets Posttranslational Modifications in Invertebrate Animal Models

Some organisms are frequently used as models for fundamental biology, such as the fruit fly Drosophila melanogaster and the worm Caenorhabditis elegans. Generally speaking, Ets studies in these organisms combine in vitro studies to understand molecular and biochemical interactions with in vivo studies to verify the role and function of these interactions at the level of the organism.

5.4.1. Drosophila melanogaster

Various Ets transcription factors have been investigated in flies, especially YAN and Pointed, which are homologs of human TEL and Ets-1, respectively. YAN is a repressor and Pointed is an activator of transcription that is encoded by two alternative transcripts, PntP1 and PntP2. YAN and PntP2 are phosphorylated by MAPK (reviewed by Hsu and Schulz (116)). The fate of eye R7 photoreceptor cells of Drosophila is controlled by a proneural signaling cascade involving Ras and Raf, and Rolled/MAPK that phosphorylates YAN and PntP2.

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The modulation of R7 cells fate can cause different eye disc phenotypes, which can be used to study the roles of YAN and PntP2 in vivo. YAN is an inhibitor of cell fate specification during fly development that has eight phosphorylation sites. Each site was mutated in vitro and the corresponding transgenic lines were produced (117). Ser-127 was found to be absolutely necessary for the response to signaling, whereas the other sites only modulate or amplify this response. YAN phosphorylation at the Ser-127 site, as well as PntP2 phosphorylation, is mediated by Mae (118). A combined study with Drosophila cells (S2) and fly eye disc phenotype demonstrated that the C-terminal 162 amino acids are necessary for YAN translocation from the nucleus to the cytoplasm (117). In vitro studies have also demonstrated that Rolled/ MAPK phosphorylates PntP2 at an unique site, which is required for PntP2 function in vivo (119). The repressor functions of YAN and TEL are regulated through protein stability and ubiquitinylation. In mammalian and Drosophila Schneider cells, the conserved F-box protein Fbl6 interacts with TEL and YAN via their SAM (PD) domains, induces ubiquitinylation and degradation. This result was confirmed for YAN in Drosophila embryos by measuring the level of YAN in the presence or absence of Fbl6 (26). 5.4.2. Caenorhabditis elegans

Ten Ets factors have been identified in C. elegans (reviewed by Hart et al. (120)). Post-translational modification of Ets factors has been studied during vulval cell fate. The most studied Ets factor is Lin-1, which belongs to the TCF subgroup, acts as a repressor and is an inhibitor of vulval cell fates. Vulval cell fate is regulated by a kinase cascade that includes Ras, Raf, MEK, MAP and leads to Lin-1 activation (121). Lin-1 works in association with the winged helix transcription factor Lin-31. In vitro experiments show that both Lin-1 and Lin-31 are phosphorylated by MAPK and that Lin-1 phosphorylation disrupts the Lin-1/Lin-31 complex, which allows Lin-31 to promote cell fate in vivo (122). Lin-1 is therefore an inhibitor whose phosphorylation leads to a decrease in its negative role. But Lin-1 also has a positive role in vivo: the egl-17::GFP marker, that is known to be activated by Ras/MAPK, has a reduced or undetectable expression in a mutant C. elegans line with lin-1 alleles with defective phosphorylation sites (123). The Lin-1 Ets factor therefore has both positive and negative functions, depending on phosphorylation.

6. Perspectives We have described analyses of increasing level of complexity, going from the molecular to the pathological process (cancer), to the signaling pathway (hypoxia), and to the animal (in vivo).

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The analysis of Post-translational modification and cellular localization is only beginning to enter the age of “omic,” high content, systems biology. They are part of the proteome and “locali-some,” that are not yet analyzable by high throughput techniques, and require low-throughput hypothesis-driven research. These snap-shots of particularly active fields point to the directions in which new techniques will be needed, in our search for a more complete description of the living system and a unified theory of biology.

Acknowledgments We would like to thank Christophe Bleunven, Jan Brants, and Catherine Fromental for critical reading of the review. We would like to thank, for fellowships: INCa (DKFZ-CGE project) for Céline Charlot; the Ministère de l’Enseignement Supérieur et de la Recherche for Hélène Dubois-Pot; the Région Alsace (DKFZCGE project) for Tsvetan Serchov; and AICR (05-390) and PRIMA (#504587) for Yves Tourrette. We would like to thank for financial support the Ligue Nationale Française contre le Cancer, the Ligue Régionale (Bas-Rhin) contre le Cancer and the Ligue Régionale (Haut-Rhin) contre le Cancer, the Association pour la Recherche contre le Cancer, the Centre National de la Recherche Scientifique, the Institut National de la Santé et de la Recherche Médicale, the EU (FP6 Prima project #504587), INCa (the Axe IV and DKFZ-CGE projects), and AICR (05-390). References 1. Buchwalter G, Gross C, Wasylyk B (2004) Ets ternary complex transcription factors. Gene 324:1–14 2. Oikawa T, Yamada T (2003) Molecular biology of the Ets family of transcription factors. Gene 303:11–34 3. Seth A, Watson DK (2005) Ets transcription factors and their emerging roles in human cancer. Eur J Cancer 41:2462–2478 4. Tootle TL, Rebay I (2005) Post-translational modifications influence transcription factor activity: a view from the ETS superfamily. Bioessays 27:285–298 5. Whitmarsh AJ, Davis RJ (2000) Regulation of transcription factor function by phosphorylation. Cell Mol Life Sci 57:1172–1183 6. Slawson C, Housley MP, Hart GW (2006) O-GlcNAc cycling: how a single sugar

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102. Salnikow K, Aprelikova O, Ivanov S, Tackett S, Kaczmarek M, Karaczyn A, Yee H, Kasprzak KS, Niederhuber J (2008) Regulation of hypoxia-inducible genes by Ets1 transcription factor. Carcinogenesis 29:1493–1499 103. Ke Q, Costa M (2006) Hypoxia-inducible factor-1 (HIF-1). Mol Pharmacol 70: 1469–1480 104. Yan SF, Lu J, Zou YS, Soh-Won J, Cohen DM, Buttrick PM, Cooper DR, Steinberg SF, Mackman N, Pinsky DJ, Stern DM (1999) Hypoxia-associated induction of early growth response-1 gene expression. J Biol Chem 274:15030–15040 105. Gross C, Dubois-Pot H, Wasylyk B (2008) The ternary complex factor Net/Elk-3 participates in the transcriptional response to hypoxia and regulates HIF-1a. Oncogene 27:1333–1341 106. Man AK, Young LJ, Tynan JA, Lesperance J, Egeblad M, Werb Z, Hauser CA, Muller WJ, Cardiff RD, Oshima RG (2003) Ets2dependent stromal regulation of mouse mammary tumors. Mol Cell Biol 23:8614–8625 107. Ilagan R, Pottratz J, Le K, Zhang L, Wong SG, Ayala R, Iyer M, Wu L, Gambhir SS, Carey M (2006) Imaging mitogen-activated protein kinase function in xenograft models of prostate cancer. Cancer Res 66: 10778–10785 108. Wei G, Guo J, Doseff AI, Kusewitt DF, Man AK, Oshima RG, Ostrowski MC (2004) Activated Ets2 is required for persistent inflammatory responses in the motheaten viable model. J Immunol 173:1374–1379 109. Sgambato V, Vanhoutte P, Pages C, Rogard M, Hipskind R, Besson MJ, Caboche J (1998) In vivo expression and regulation of Elk-1, a target of the extracellular-regulated kinase signaling pathway, in the adult rat brain. J Neurosci 18:214–226 110. Vanhoutte P, Barnier JV, Guibert B, Pages C, Besson MJ, Hipskind RA, Caboche J (1999) Glutamate induces phosphorylation of Elk-1 and CREB, along with c-fos activation, via an extracellular signal-regulated kinase-dependent pathway in brain slices. Mol Cell Biol 19:136–146 111. Choe ES, Wang JQ (2001) Group I metabotropic glutamate receptor activation increases phosphorylation of c-AMP responses element-binding protein, Elk-1 and extracellular signal-regulated kinases in rat dorsal striatum. Brain Res Mol Brain Res 94:75–84

112. Choe ES, Parelkar NK, Kim JY, Cho HW, Kang HS, Mao L, Wang JQ (2004) The protein phosphatase 1/2A inhibitor odakaic acid increases CREB and Elk-1 phosphorylation and c-fos expression in the rat striatum in vivo. J Neurochem 89:383–390 113. Choe ES, Wang JQ (2002) CREB and Elk-1 phosphorylation by metabotropic glutamate receptors in striatal neurons. Int J Mol Med 9:3–10 114. Nuutinen S, Barik J, Jones IW, Wonnacott S (2007) Differential effects of acute and chronic nicotine on Elk-1 in rat hippocampus. NeuroReport 18:121–126 115. Berman DE, Hazvi S, Rosenblum K, Seger R, Dudai Y (1998) Specific and differential activation of mitogen-activated protein kinase cascades by unfamiliar taste in the insular cortex of the behaving rat. J Neurosci 18:10037–10044 116. Hsu T, Schulz RA (2000) Sequence and functional properties of Ets genes in the model organism Drosophila. Oncogene 19:6409–6416 117. Rebay I, Rubin GM (1995) Yan functions as a general inhibitor of differentiation and is negatively regulated by activation of the Ras1/MAPK pathway. Cell 81:857–866 118. Baker DA, Mille-Baker B, Wainwright SM, Ish-Horowicz D, Dibb NJ (2001) Mae mediates MAP kinase phosphorylation of Ets transcription factors in Drosophila. Nature 411:330–334 119. Brunner D, Ducker K, Oellers N, Hafen E, Scholz H, Klambt C (1994) The Ets domain protein pointed-P2 is a target of MAP kinase in the sevenless signal transduction pathway. Nature 370:386–389 120. Hart AH, Reventar R, Bernstein A (2000) Genetic analysis of Ets genes in C. elegans. Oncogene 19:6400–6408 121. Beitel GJ, Tuck S, Greenwald I, Horvitz HR (1995) The Caenorhabditis elegans gene lin-1 encodes an Ets-domain protein and defines a branch of the vulval induction pathway. Genes Dev 9:3149–3162 122. Tan PB, Lackner MR, Kim SK (1998) MAP kinase signaling specificity mediated by the LIN_1 Ets/LIN-31 WH transcription factor complex during C. elegans vulval induction. Cell 93:569–580 123. Tiensuu T, Larsen MK, Vernersson E, Tuck S (2005) Lin-1 has both positive and negative functions in specifying multiple cell fates induced by Ras/MAP kinase signaling in C. elegans. Dev Biol 286:338–351

Chapter 2 Regulation of Transcription Factor Function by Targeted Protein Degradation: An Overview Focusing on p53, c-Myc, and c-Jun Jukka Westermarck Abstract Regulation of protein degradation is an important mechanism by which concentrations of proteins is controlled in cells. In addition to proteins involved in cell cycle regulation or mitosis, protein levels of many transcription factors are regulated by targeted proteosomal degradation. Regulation of protein degradation and stability is usually linked to post-translational modification of the target protein by phosphorylation. The resulting phosphoaminoacid in the context of the adjacent protein sequence is then recognized by E3 ubiquitin ligase enzymes that covalently attach small ubiquitin protein to the target protein and thereby direct them to be degraded by the proteosomes. Here, we present an overview of mechanisms regulating stability of p53, c-Myc, and c-Jun transcription factors. Especially, the purpose is to highlight the role of protein phosphorylation in the regulation of stability of these transcription factors. We also present examples where phosphorylation can either enhance or inhibit protein degradation. Lastly, we discuss the common theme among p53, c-Myc, and c-Jun proteins that the N-terminal phosphorylation both increases the transactivation capacity of the protein and protects the protein from proteolytic degradation. Key words: c-Jun, c-Myc, p53, Phosphorylation, Proteosomal degradation, Ubiquitin

1. Introduction Protein stability is an important mechanism by which concentrations of biologically active proteins is controlled in cells. Changes in protein stability occurs usually when rapid fluctuations in protein amounts are needed. Some proteins are intrinsically very stable but become degraded in response to specific physiological signals or cellular state. One example of such proteins are those involved in cell cycle regulation. Other proteins are usually very short lived and degraded at all times, but they Paul J. Higgins (ed.), Transcription Factors: Methods and Protocols, Methods in Molecular Biology, vol. 647, DOI 10.1007/978-1-60761-738-9_2, © Springer Science+Business Media, LLC 2010

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become stabilized when a rapid increase in protein amount is required and induction of protein expression by increased gene transcription is not sufficient. This type of mechanism is best exemplified by the very rapid increase in p53 transcription factor stability in response to cellular stress (1). However, often both gene transcription and protein stability are coupled synchronously to either repress or increase protein amounts. c-myc gene transcription and stabilization of MYC protein, for example, are modulated in response to phosphorylation by ERK kinase (2, 3). Control of protein stability is therefore a powerful mechanism to regulate protein amount and therefore this mechanism has been hijacked by cancer cells to promote oncogenic behavior. In general, alterations linked to protein stabilization in cancer tend to accelerate degradation of tumor suppressor proteins and in turn protect oncoproteins from degradation (4, 5). Regulation of protein stability is very often linked to protein phosphorylation. The resulting phosphoaminoacid in the context of the adjacent protein sequence (phospho-degron) is recognized by E3 ubiquitin ligase enzymes that covalently attach small ubiquitin proteins moieties the lysine residue of the target protein (6). This is followed by sequential attachment of new ubiquitin molecules to the ubiquitins already linked to specific target protein to build up a polyubiquitin chain. A protein marked with a polyubiquitin chain is consequently transported to cellular organelles responsible for protein degradation, the 26S proteosomes, and through a multi-step process digested into short peptides (6). Several families of ubiquitin ligases exist, and the same target protein can be subjected to ubquitination by several different ubiquitin ligases potentially resulting in different biological outcomes (6, 7).

2. Regulation of Transcription Factor Expression by Targeted Protein Degradation

Protein levels of many transcription factors are regulated by targeted proteosomal degradation. The contribution of mechanisms regulating protein stability to overall protein levels vary greatly between different proteins and cellular conditions. Although biologically intelligible, the simultaneous increase or decrease in mRNA and protein levels may mask the influence of protein stabilization, and thus mislead the interpretations regarding mechanisms that are truly relevant for regulation of steady-state levels of the studied protein. Therefore, should mRNA and protein expression levels, or the trend in their regulation, does not readily correlate, it is relevant to consider if protein stability is affected. On the other hand, regulation of protein stability would

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appropriately be considered in the cases where protein expression is lost without parallel regulation of corresponding mRNA, or when steady-state levels of both endogenous and exogenously expressed forms of the same protein are regulated in a similar fashion. In all these cases, protein stability can be easily assessed by experiments where protein translation is blocked with cycloheximide treatment, and the decline of protein amounts is followed by standard Western blotting or radioactive protein labeling techniques, as described previously (8–10). Here I present an overview of mechanisms regulating stability of p53, c-Myc, and c-Jun transcription factors. Rather than a complete review, the aim here is to introduce p53, c-Myc, and c-Jun as examples how transcription factor expression is regulated by proteosomal degradation. The purpose is to highlight the role of protein phosphorylation in the regulation of transcription factor protein stability. For more details, and for information about other ubiquitin ligases regulating of p53, c-Myc, and c-Jun activities, reader is encouraged to become acquainted with the following articles (1, 3, 6, 7, 9, 11, 12). 2.1. p53

Transcription factor p53 is a short-lived protein in normal quiescent cells. Its half-life is approximately 20 min and it is continuously degraded by the proteosomal system. p53 degradation is mostly controlled by its association with the ubiquitin E3 ligase Mdm2 which binds to the aminoterminal domain of p53 and targets the newly synthesized protein for degradation by tagging it with ubiquitin (1). Interestingly, the Mdmd2 binding domain of p53 is phosphorylated by several kinases regulating p53 transcriptional activity and stability. The kinases known to phosphorylate p53 include Chk1, Chk2, ATM/ATR, the activity of which are rapidly induced in response to genotoxic stress. Phosphorylation of the aminoterminal amino acids of p53 changes structure of the aminoterminal domain inhibiting Mdm2 binding. Interestingly, phosphorylation of Mdm2 also regulates its association with p53, and thereby p53 stability (1). Survival promoting signals, through Akt kinase-mediated phosphorylation, stimulates Mdm2 binding to p53, and consequently enhances p53 degradation. Conversely, some of the stress-activated kinases described above (Chk2, ATM/ ATR) can, in addition to directly phosphorylating p53, inactivate Mdm2 and thereby promote p53 stabilization. Taken together, phosphorylation-dependent regulation of Mdm2 binding to p53, by any of the mechanisms described above, provides a very elegant way to regulate p53 activity, and by these means allows cells to respond rapidly to cellular stress and survival signals.

2.2. c-Myc

Oncogenic transcription factor c-Myc is expressed at low levels in normal cells, but both c-myc mRNA expression and protein

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stability is increased in response to mitogenic stimuli and cellular transformation. Like, p53, regulation of c-Myc stability is closely linked with protein phosphorylation. c-Myc harbors a phosphodegron motif recognized by Fbw7 ubiquitin ligase in its N-terminal domain (5). Within this motif, serine 62 phosphorylation by ERK kinase is required as “priming phosphorylation” for GSK3mediated phosphorylation of an adjacent threonine 58 (3, 5). In vitro, c-Myc peptide that is double phosphorylated on serine 62 and threonine 58 binds to Fbw7 (9), but experiments in cultured cells demonstrate that if serine 62 is phosphorylated, the protein is protected from proteosomal degradation (10, 13). Additionally, serine 62 has been shown to be dephosphorylated by tumor suppressor protein phosphatase 2A (PP2A) and this leads to c-Myc destabilization (10, 13). Therefore, the stability of c-Myc is regulated by a complex interplay between kinases phosphorylating threonine 58 and serine 62, and PP2A phosphatase activity. Interestingly, recent studies have identified new proteins involved in the regulation of c-Myc stability through Fbw7 phosphodegron. Cancerous inhibitor of PP2A (CIP2A) was shown to inhibit c-Myc serine 62 dephosphorylation by PP2A and to stabilize c-Myc (8). Ubiquitin-specific protease 28 in turn was shown to antagonize Fbw7-mediated c-Myc ubiquitination and by these means to prevent c-Myc degradation (14). Both CIP2A and USP28 are upregulated in human cancers illustrating an additional level of regulation of protein stability to promote tumorigenesis. 2.3. c-Jun

c-Jun is an AP-1 family transcription factor implicated in the regulation of cell death and survival as well as in neurological degeneration and cellular transformation (15). Similarly to c-myc, c-jun is an immediate-early gene whose mRNA expression is rapidly induced by both mitogenic and stress signals, and which protein stability is enhanced by the same stimuli that increases gene expression. Activation of stress-activated JNK kinases results in phosphorylation of c-Jun on serines 63/73 and on threonines 91/93. c-Jun N-terminal phosphorylation has been shown to increase the transactivation potential of c-Jun, but it also inhibits ubiquitination and degradation of the protein (15, 16). Whereas JNK-mediated N-terminal phosphorylation stabilizes the c-Jun protein, GSK3-mediated phosphorylation of c-Jun on threonine 239 induces Fbw7 E3-ligase recruitment and protein degradation. Interestingly, the Fbw-7 phospho-degron in c-Jun and c-Myc is highly similar in sequence (9). In both proteins, GSK3mediated phosphorylation of the threonine (239 in c-Jun and 58 in c-Myc) requires a priming phosphorylation at the +4 position. However, whereas in c-Myc the priming phosphorylation is done by ERK kinase and this phosphorylation has to be removed in order for the protein to become ubiquitinylated (10),

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in c-Jun the kinase responsible for serine 243 phosphorylation is not known, and this phosphorylation stimulates protein ubiquitinylation and degradation (9).

3. Conclusions In addition to examples presented above, expression of many other transcription factors, such as b-catenin, HIF-1a, and Smad proteins, is regulated by targeted proteosomal degradation. One common theme among proteins targeted for ubiquitination is that post-translational modification of the E3 ubiquitin ligasebinding site determines the efficiency of protein ubiquitination and degradation. Whereas in the case of most of the other transcription factors this post-translational modification is phosphorylation, in the case of HIF-1a, it is prolyl hydroxylation (17). Moreover, as exemplified with c-Jun above, one protein can be subjected to phosphorylation-dependent regulation of ubiquitination in more than one protein domain, and depending on the site, phosphorylation can either enhance or inhibit ubiquitination and protein stability. On the other hand, in addition to the specific transcription factor to be modified by phosphorylation and ubiquitination, phosphorylation of the ubiquitin ligase regulates its activity and thereby target protein stability. The best established examples of this are phosphorylation of p53 and c-Jun ubiquitin ligases Mdm2 and Itch, respectively (1, 11). Lastly, an additional common theme shared by p53, c-Myc, and c-Jun is that the N-terminal phosphorylation that increases the transactivation capacity of the protein also prevents the protein from proteolytic degradation (1, 16, 18). Even though this complicates conclusions regarding the biological role of such phosphorylations it clearly suggests that natue has evolved such system to enhance activity of these transcription factors in the most economical manner. References 1. Lavin MF, Gueven N (2006) The complexity of p53 stabilization and activation. Cell Death Differ 13:941–950 2. Sears R, Leone G, DeGregori J, Nevins JR (1999) Ras enhances Myc protein stability. Mol Cell 3:169–179 3. Sears RC (2004) The life cycle of C-myc: from synthesis to degradation. Cell Cycle 3:1133–1137 4. Junttila MR, Westermarck J (2008) Mechanisms of MYC stabilization in human malignancies. Cell Cycle 7:592–596

5. Welcker M, Clurman BE (2008) FBW7 ubiquitin ligase: a tumour suppressor at the crossroads of cell division, growth and differentiation. Nat Rev Cancer 8:83–93 6. Welchman RL, Gordon C, Mayer RJ (2005) Ubiquitin and ubiquitin-like proteins as multifunctional signals. Nat Rev Mol Cell Biol 6:599–609 7. Dai MS, Jin Y, Gallegos JR, Lu H (2006) Balance of Yin and Yang: ubiquitylationmediated regulation of p53 and c-Myc. Neoplasia 8:630–644

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8. Junttila MR, Puustinen P, Niemela M, Ahola R, Arnold H, Bottzauw T, Ala-Aho R, Nielsen C, Ivaska J, Taya Y, Lu SL, Lin S, Chan EK, Wang XJ, Grenman R, Kast J, Kallunki T, Sears R, Kähäri VM, Westermarck J (2007) CIP2A inhibits PP2A in human malignancies. Cell 130:51–62 9. Wei W, Jin J, Schlisio S, Harper JW, Kaelin WG Jr (2005) The v-Jun point mutation allows c-Jun to escape GSK3-dependent recognition and destruction by the Fbw7 ubiquitin ligase. Cancer Cell 8:25–33 10. Yeh E, Cunningham M, Arnold H, Chasse D, Monteith T, Ivaldi G, Hahn WC, Stukenberg PT, Shenolikar S, Uchida T, Counter CM, Nevins JR, Means AR, Sears R (2004) A signalling pathway controlling c-Myc degradation that impacts oncogenic transformation of human cells. Nat Cell Biol 6:308–318 11. Gao M, Labuda T, Xia Y, Gallagher E, Fang D, Liu YC, Karin M (2004) Jun turnover is controlled through JNK-dependent phosphorylation of the E3 ligase Itch. Science 306:271–275 12. Wertz IE, O’Rourke KM, Zhang Z, Dornan D, Arnott D, Deshaies RJ, Dixit VM (2004) Human De-etiolated-1 regulates c-Jun by assembling a CUL4A ubiquitin ligase. Science 303:1371–1374

13. Arnold HK, Sears RC (2006) Protein phosphatase 2A regulatory subunit B56alpha associates with c-myc and negatively regulates c-myc accumulation. Mol Cell Biol 26:2832–2844 14. Popov N, Wanzel M, Madiredjo M, Zhang D, Beijersbergen R, Bernards R, Moll R, Elledge SJ, Eilers M (2007) The ubiquitin-specific protease USP28 is required for MYC stability. Nat Cell Biol 9:765–774 15. Shaulian E, Karin M (2002) AP-1 as a regulator of cell life and death. Nat Cell Biol 4:E131–E136 16. Musti AM, Treier M, Bohmann D (1997) Reduced ubiquitin-dependent degradation of c-Jun after phosphorylation by MAP kinases. Science 275:400–402 17. Jaakkola P, Mole DR, Tian YM, Wilson MI, Gielbert J, Gaskell SJ, Kriegsheim A, Hebestreit HF, Mukherji M, Schofield CJ, Maxwell PH, Pugh CW, Ratcliffe PJ (2001) Targeting of HIF-alpha to the von Hippel–Lindau ubiquitylation complex by O2-regulated prolyl hydroxylation. Science 292:468–472 18. Benassi B, Fanciulli M, Fiorentino F, Porrello A, Chiorino G, Loda M, Zupi G, Biroccio A (2006) c-Myc phosphorylation is required for cellular response to oxidative stress. Mol Cell 21:509–519

Chapter 3 Review of Molecular Mechanisms Involved in the Activation of the Nrf2-ARE Signaling Pathway by Chemopreventive Agents Aldo Giudice, Claudio Arra, and Maria C. Turco Abstract Human exposures to environmental toxicants have been associated with etiology of many diseases including inflammation, cancer, and cardiovascular and neurodegenerative disorders. To counteract the detrimental effect of environmental insults, mammalian cells have evolved a hierarchy of sophisticated sensing and signaling mechanisms to turn on or off endogenous antioxidant responses accordingly. One of the major cellular antioxidant responses is the induction of antioxidative and carcinogen-detoxification enzymes through the cytoplasmic oxidative stress system (Nrf2-Keap1) activated by a variety of natural and synthetic chemopreventive agents. Under normal conditions, Keap1 anchors the Nrf2 transcription factor within the cytoplasm targeting it for ubiquitination and proteasomal degradation to maintain low levels of Nrf2 that mediate the constitutive expression of Nrf2 downstream genes. When cells are exposed to chemopreventive agents and oxidative stress, a signal involving phosphorylation and/or redox modification of critical cysteine residues in Keap1 inhibits the enzymatic activity of the Keap1–Cul3–Rbx1 E3 ubiquitin ligase complex, resulting in decreased Nrf2 ubiquitination and degradation. As a consequence, free Nrf2 translocates into the nucleus and in combination with other transcription factors (e.g., sMaf, ATF4, JunD, PMF-1) transactivates the antioxidant response elements (AREs)/electrophile response elements (EpREs) of many cytoprotective genes, as well as Nrf2 itself . Upon recovery of cellular redox homeostasis, Keap1 travels into the nucleus to dissociate Nrf2 from the ARE. Subsequently, the Nrf2– Keap1 complex is exported out of the nucleus by the nuclear export sequence (NES) in Keap1. Once in the cytoplasm, the Nrf2–Keap1 complex associates with the Cul3-Rbx1 core ubiquitin machinery, resulting in degradation of Nrf2 and termination of the Nrf2/ARE signaling pathway. The discovery of multiple nuclear localization signals (NLSs) and nuclear export signals (NESs) in Nrf2 also suggests that the nucleocytoplasm translocation of transcription factors is the consequence of a dynamic equilibrium of multivalent NLSs and NESs. On the other hand, Keap1 may provide an additional regulation of the quantity of Nrf2 both in basal and inducible conditions. This chapter summarizes the current body of knowledge regarding the molecular mechanisms through which ARE inducers (chemopreventive agents) regulate the coordinated transcriptional induction of genes encoding phase II and antioxidant enzymes as well as other defensive proteins, via the nuclear factor-erythroid 2 (NF-E2-p45)-related factor 2(Nrf2)/ (ARE) signaling pathway. Key words: ARE/EpREs, Chemopreventive agents, Cul3, Cysteine residues, Degradation, E3 ubiquitin ligases, Exportins, Importins, Keap1, NES, NLS, Nrf2, Oxidative stress, Ubiquitination

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1. Introduction Human exposures to environmental toxicants have been associated with etiology of many diseases including inflammation, cancer, cardiovascular and neurodegenerative disorders, sepsis, reperfusion damage, and diabetes (1–3). To counteract the detrimental effect of environmental insults, mammalian cells have evolved a hierarchy of sophisticated sensing and signaling mechanisms to turn on or off endogenous antioxidant responses accordingly (2). One of the major cellular antioxidant responses is the induction of antioxidative and carcinogen-detoxification enzymes through the cytoplasmic oxidative stress system (Nrf2-Keap1) activated by a variety of natural and synthetic chemopreventive agents (4). Under normal conditions, Keap1 anchors the Nrf2 transcription factor within the cytoplasm targeting it for ubiquitination and proteasomal degradation to maintain low levels of Nrf2 that mediate the constitutive expression of Nrf2 downstream genes. When cells are exposed to chemopreventive agents and oxidative stress, a signal involving phosphorylation and/or redox modification of critical cysteine residues in Keap1 inhibits the enzymatic activity of the Keap1–Cul3–Rbx1 E3 ubiquitin ligase complex, resulting in decreased Nrf2 ubiquitination and degradation. As a consequence, free Nrf2 translocates into the nucleus and in combination with other transcription factors (e.g., sMaf, ATF4, JunD, PMF-1) transactivates the antioxidant response elements (AREs)/ electrophile response elements (EpREs) of many cytoprotective genes, as well as Nrf2 itself (3–6). The families of enzymes induced by chemopreventive agents have been classified into several categories: (a) phase II xenobiotic-metabolizing enzymes (e.g., glutathione S-transferases (GSTs), UDP-glucuronosyltransferases (UDPGTs), NAD(P)H:quinone oxidoreductase 1 (NQO1), epoxide hydrolase (EH), aflotoxin B1 aldehyde reductase (AFAR), heme oxygenase 1 (HO-1), ferritin); (b) antioxidants and their modulating enzymes (e.g., gamma-glutamyl-cysteine synthetase (g-GCS), superoxide dismutase (SOD), catalase (CAT), glutathione reductase (GR), thioredoxin reductase (TR), peroxiredoxins (Prxs), glutathione S-conjugate efflux pumps, nicotinamide adenine dinucleotide phosphate (NADPH) and cofactors-generating enzymes); (c) molecular chaperones/proteasome systems; (d) DNA repair enzymes; and (e) anti-inflammatory response proteins (e.g., HO-1, ferritin, leucokotriene B4 dehydrogenase) (6–10). The study of carcinogenesis in experimental models has suggested that tumor development consists of at least three distinct stages: initiation (the fixation of mutations in the DNA), promotion (the appearance of benign tumors), and progression (the conversion of benign tumors into malignancies), which gradually transform into highly malignant tumors with strong

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metastatic capabilities (1, 11, 12). Chemopreventive agents (also called collectively as ARE inducers) have been divided into blocking agents and suppressing agents, based on the stage of carcinogenesis during which they act (12, 13). Blocking agents (e.g., coumarins, phenols, flavones, terpenes, indoles, isothiocyanates (ITCs), oltipraz (OPZ)) inhibit initiation or prevent carcinogens from modifying DNA and causing mutations. This is usually achieved by either decreasing the formation of carcinogens from precursor molecules, inhibiting the formation of reactive metabolites from parent carcinogens, or preventing the ultimate electrophilic and carcinogenic species from interacting with critical cellular target molecules like DNA, RNA, and proteins (12–14). Conversely, suppressing (or antiproliferative) agents (e.g., the retinoids, vitamin E, the carotenoids and other antioxidants, selective estrogen receptor modulators (SERMs) like tamoxifen and raloxifene, lipooxygenase (LOX) inhibitors, cyclooxygenase (COX) inhibitors), inhibit the malignant expression of initiated cells in either the promotion or progression stages (12–15). Certain chemopreventive agents (e.g., curcumin, resveratrol, theaflavins, thearubigins, catechins and the dithiolethiones) possess both blocking and suppressive properties, for they not only induce antioxidant and phase II enzymes, but also suppress gene transcription of the cytochrome P450 (CYP) isoenzymes and the genes involved in lipid/cholesterol biosynthesis (6, 12, 14). Two general categories of inducers (or chemopreventive blocking agents) which enhance carcinogen-detoxification enzyme activity have been identified: bifunctional inducers and monofunctional inducers. Bifunctional inducers upregulate both phase I (mainly the CYPs: YP1A1, CYP1A2) and phase II enzymes via the xenobiotic response element (XRE). Monofunctional inducers, on the other hand, primarily elevate phase II enzymes (e.g., GSTs, UDPGTs, NQO-1) via the (ARE)/(EpRE) (electrophile-responsive element) (4, 8, 16–18). It is well known that the majority of dietary or environmental carcinogens to which we are exposed require metabolic activation to unmask their carcinogenic activity (7, 16, 19, 20). The biotransformation of many foreign substances or xenobiotics, though complex, can be considered to comprise two sequential reaction processes: phase I and phase II. Two families of ubiquitous and inducible detoxification enzymes involved in the metabolism of xenobiotics have been identified. Phase I enzymes (which primarily include the CYPs) metabolize compounds (procarcinogens), either by oxidation, reduction, or hydrolysis into inactive and chemically reactive electrophilic metabolites that covalently bind to specific sites on DNA to initiate a carcinogenic response. Phase II enzymes (e.g., GSTs, UDP-GTs, NQO-1, HO-1) primarily inactivate active electrophilic

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metabolites formed by phase I enzymes (16, 19, 20). They also perform conjugation reactions of functionalized compounds (active electrophilic metabolites) with endogenous ligands (e.g., glutathione (GSH), glucuronic acid, amino acids and sulfate) helping convert the biotransformed intermediates from phase I into less toxic, water soluble substances that are easily excreted or eliminated from the body (conjugating metabolism is principally, but not invariably, detoxifying) (7, 16, 17, 19, 20). They may even catalyze reactions independent of phase I activity, acting directly upon a toxin or on endogenous mutagens (free radicals, ROS) not requiring biotransformation (16, 17, 19, 21, 22). This chapter summarizes the primary functions of the inducers of these phase I and phase II xenobiotic detoxification and antioxidant enzymes, in particular examining the molecular mechanisms through which ARE inducers regulate the coordinated transcriptional induction of genes encoding phase II and antioxidant enzymes, as well as other defensive proteins, via the nuclear factor-erythroid 2 (NF-E2-p45)-related factor 2 (Nrf2)/(ARE) signaling pathway.

2. The Importance of the Nrf2 Transcription Factor in the Induction of Genes Encoding Antioxidant and Phase II Detoxification Enzymes by Chemopreventive Agents

The induction of a family of antoxidant/detoxification genes encoding enzymes that protect against electrophilic and reactive oxygen intermediate damage is a potentially major strategy in reducing the risk of cancer and other chronic degenerative diseases linked to oxidative stress (4, 18, 21, 22). There are several important lines of defense: (1) antioxidant and their modulating enzymes (e.g., g-GCS, SOD, CAT, GR, TR, Prxs, glutathione S-conjugate efflux pumps, nicotinamide adenine dinucleotide phosphate (NADPH) and cofactors-generating enzymes), (2) DNA repair enzymes; (3) molecular chaperones/proteasone systems and (4) a large family of phase II enzymes (e.g., GSTs, UDPGTs, NQO1, epoxide hydrolase (EH), aflotoxin B1 aldehyde reductase (AFAR), HO-1, ferritin), capable of converting reactive electrophiles to less toxic and more readily excretable products, thus protecting cells against various chemical stresses and carcinogenesis (6–10). These defensive proteins may collectively facilitate the detoxification of carcinogens, enhance the reducing potential against electrophiles and free radicals, and increase cellular capacity to repair oxidatively damaged DNA and proteins (5). In addition, since antioxidant/detoxification enzyme activities and GSH levels do not normally operate at their maximum capacity, their ability to be transcriptionally induced by a wide variety of natural and

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synthetic chemical agents should promote efficient protection against carcinogenesis (22). Central to this transcriptional response is a recently identified sensor system known as the cytoplasmic oxidative stress system (Nrf2–Keap1), which appears to be the primary molecular target of chemopreventive agents (8, 21, 24). Two proteins participate in the transcriptional activation of ARE-dependent phase II genes: (1) Nrf2, a transcription factor which is a member of the nuclear factor-erythroid 2 (NF-E2) family of nuclear basic leucine zipper (bZIP) transcription factors, and (2) Keap1 (Kelch-like erythroid-cell-derived protein with CNC homology (ECH)-associated protein 1], a cytoplasmic protein homologous to the Drosophila actin-binding protein Kelch. Under basal conditions, Nrf2 molecules are predominantly sequestered in the cytoplasm by a cysteine-rich protein called Keap1 (23, 24). When cells are exposed to chemopreventive agents (e.g., dithiolethiones, flavonoids, ITCs) and oxidative stress, a signal involving phosphorylation and/or redox modification is transmitted to the Nrf2–Keap1 complex, leading the dissociation and the subsequent nuclear translocation of Nrf2. Nrf2, after heterodimerically partnering with other transcription factors (e.g., small musculoaponeurotic fibrosarcoma (sMaf): MafF, MafG, and MafK; JunD; activation transcription factor 4 (ATF4); polyamine-modulated factor-1 protein (PMF-1)], then binds to the ARE/EpREs present in the promoters of phase II genes, increasing their transcription and that of Nrf2 as well (4, 6, 23, 25, 26). Recently, it was shown by Lin et al. (27) that the receptor associated coactivator(RAC3)/steroid receptor coactivator-3 (SRC3) is involved in the functional transactivation of TAD (the transactivation domain) of Nrf2 and that this transactivation activity could be further enhanced by the coregulators such as CBP/p300 (CREB-binding protein), p/CAF (p300/CBP-associated factor), CARM1 (coactivator-associated arginine methyltransferase), and PRMT1 (protein arginine methyl-transferase1). Although significant attention has been directed toward understanding mechanisms of induction, suppression of Nrf2 transactivation is less understood but phenotypically as important. Currently, it is known that the transcriptional state of AREregulated genes is determined by the identity of the dimer recruited. For example, sMaf homodimers, which lack transactivation domains, are not able to drive transcription from this element, while the Bach1 (BTB and CNC homology 1)-containing heterodimers (Bach transcription factors) actively repress transcription (28, 29). In fact, the Bach transcription factors, compete with Nrf2 for both the Maf proteins and the Maf recognition element (MARE)/ARE sequences in the target DNA (28, 29). Importantly, the existence of functionally distinct bZIP dimers allows the cell to control ARE-driven gene transcription, varying

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the quantity of specific cap “n” collar (CNC) and sMaf proteins in the nucleus, and thus the spectrum of dimers expressed (28). In addition, since the small proteins have a wider choice of partner molecules for binding to the MARE (Maf, protein recognition element) depending on the dimeric partner chosen, sMafs are also able to switch transcriptional activity from repression to activation (28). Accordingly, it is expected that many transcriptional repressors (e.g., Nrf3, ATF3, p53) in association with small Maf proteins or by dimerization with other activators of ARE (e.g., Nrf2, c-AMP-responsive element-binding protein (CREB)] bind to the ARE and repress ARE-mediated gene expression (30–33). In agreement with this studies, Wang and Wolf (23) showed that retinoids such as all-trans retinoic acid (ATRA) and other retinoic acid receptor alpha (RARa) agonists, markedly reduce the ability of Nrf2 to mediate induction of ARE-driven genes by cancer chemopreventive agents including the metabolite of butylated hydroxyanisole, tert-butylhydroquinone (tBHQ). It is expected that retinoids antagonize Nrf2 function by stimulating the formation of Nrf2:RARa-containing complexes that do not bind to the ARE. Another possibility is that RARa may cause subnuclear relocalization of Nrf2 because it has been shown that retinoic acid (RA) can affect delocalization of transcriptional intermediary factor 1b into regions of centromeric heterochromatin (34). Recent studies have shown that the Nrf2–MafK heterodimer regulates placental glutathione S-transferase (GST-P) expression, a phase II detoxifying enzyme, which is not expressed in normal liver cells but is highly and specifically induced by the action of the GST-P enhancer 1 (GPE1) during early hepatocarcinogenesis in hepatoma cells (29). Interestingly, several groups have found somatic mutations in the Keap1 gene in human lung cancer cells, which result in increased activity of Nrf2 and higher levels of ARE-regulated genes (35, 36). These findings suggest that lung tumor cells hijack the Nrf2 pathway to increase their survival, likely to combat the high oxygen environment of the lung as well as chemotherapeutic agents (35, 36). Major insight into Nrf2’s contribution to this protective response was provided by the lack of upregulation of phase II genes in mice lacking this factor (20, 37–42). These experiments, which compared Nrf2 knockout and wild-type mice, provided strong evidence that: (a) the protective action of three phase II enzyme inducers (t-BHA, SUL and OPZ) is abolished in the absence of the Nrf2 gene function, (b) the susceptibility to carcinogenesis is markedly increased when the synthesis of phase II proteins is suppressed, and (c) basal levels of phase II proteins exert significant protection against carcinogenesis. However, recently, Marzec et al. (43) identified a number of single nucleotide polymorphisms (SNPs) in the promoter region of Nrf2 present in human subjects across multiple ethnic groups. These also observed that one of the SNPs resulted

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in decreased in vitro binding of Nrf2 to an ARE promoter following exposure to Nrf2-inducing stresses. Importantly, individuals with this SNP were found to be more likely to develop acute lung injury, relative to individuals with a normal Nrf2 sequence, following major trauma. Together, these observations suggest that the Nrf2–Keap1 complex constitutes a cytoplasmic sensor system of great importance in the constitutive and inducible expression of phase II and antioxidant enzymes by chemopreventive agents, capable of dramatically influencing susceptibility to carcinogenesis and other degenerative pathologies.

3. Structure and Functions of the Members of the Cytoplasmic Oxidative Stress System (Nrf2–Keap1)

Nrf2 is a member of the basic leucine zipper (bZIP) transcription factor subfamily featuring a cap ‘n collar motif (44). Structurally, Nrf2 comprises six regions, called Neh (Nrf2-ECH homology) 1–6 domains, which are highly conserved across different species. Neh 1 contains the CNC-bZIP region, which promotes dimerization partners and confers DNA-binding specificity. The Neh4 and Neh5 domains act cooperatively to bind the coactivator CBP (CREB (c-AMP-response element-binding protein)/ATF4] and BRG1 (Brahma-related gene 1) thereby, activating transcription (45). The Neh5 domain is conserved among CNC transcription factors, such as p45 and Nrf1, whereas Neh4 shows more structural similarity to transcription factors, such as p53 and E2F (46). Neh3 is a C-terminal domain and also contributes to Nrf2 transactivation (47). The previously uncharacterized, redox-insensitive Neh6 domain (amino acids 329–379) is essential in the Keap1indipendent degradation of Nrf2 that occurs in the nucleus of oxidatively stressed cells (48). Of particular interest is the N-terminal region of 100 amino acids, called the Neh2 domain, which contains both the DIDLID element (amino acids 17–32) also termed DLG motif and the ETGE tetrapeptide motif (amino acids 79–82), and negatively downregulates Nrf2 function under homeostatic conditions by mechanisms which are not yet fully understood (5, 48). Recently, it was shown that two sites within the Neh2 domain of Nrf2, termed the DGL and ETGE motifs mediate binding to the Keap1 double glycine repeats (DGR) or Kelch repeats region (49, 50). Keap1 protein functions as a bridge between Nrf2 and the Cullin3-based E3-ligase ubiquitination complex, promoting ubiquitination of lysines in the Neh2 domain and subsequent proteasomal degradation of Nrf2, thus preventing nuclear accumulation of Nrf2. These lysines are located between the two Kelch-binding sites on Neh2 (51) and a model has been proposed whereby binding of a Keap1 homodimer to these two sites allows for ubiquitination to occur (49, 52). Recently, two Crm1/exportin-dependent

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nuclear export signal (NESs) sequences within the Nrf2 protein have been identified, as well (53, 54). A canonical redox-insensitive NESZIP has been found within the luecine zipper (ZIP) domain of the Nrf2 protein. With regard to this NESZIP, there has been some confusion in the literature because two redox-insensitive NESs have been found. In fact, the NES reported as 537LKKQLSTLYL546 (53) and the NES reported as 545LKRRLSTLYL554 (55) both refer to the same region of homology in Nrf2 from two different species, human and mouse. Recently, it was shown that Fyn kinase phosphorylation of tyrosine 568 in Nrf2 regulates Crm1-mediated nuclear export and degradation of Nrf2 (56). The mechanism of phosphorylated Nrf2 interaction with Crm1 remains unknown. It is expected that phosphorylation of Nrf2Y568 leads to structural changes that expose the leucine-rich NES region (amino acid 545-554) for interaction with Crm1. Accordingly, mutation of tyrosine 568 to alanine or phenylalanine resulted in the loss of phosphorylation and interaction of Nrf2 with Crm1 and abrogation of nuclear export of Nrf2. An additional redox-sensitive NESTA reported as 175LLSIPELQCLNI186 has been found in the Neh5 transactivation (TA) domain of Nrf2 (54). Under normal physiological conditions, the redox reactivity of the NESTA motif enables Nrf2 to detect oxidative signals and transmit them to the nucleus. Mutation analyses showed that NESTA redox sensitivity may be mediated by the C183 residue. Accordingly, C183A mutation could remarkably slow down translocation kinetics and attenuate Nrf2/ARE-mediated gene expression. It is possible that direct sulfhydryl modification of the Cys-183 residue inhibits the access and binding of nuclear exportin CRM1 to the NESTA motif and consequently results in nuclear accumulation of EGFPNESTA. Alternatively, intramolecular disulfide bond formation may also disable the NES activities (57). Recently, it was shown that Nrf2 protein, in addition to the two NESs, which interact with exportins, also contains multiple nuclear localization signals (NLSs) which likely facilitate Nrf2 nuclear localization upon addition of ARE inducers (58). Both of these monopartite sequences are designated NLS1 and NLS3, respectively, (58) to distinguish them from a previously identified bipartite sequence termed NLS2, which had been implicated in the nuclear translocation of this transcription factor (55). Interestingly, NLS1 occurs within the Neh2 domain of Nrf2 (amino acid residues 42-53) whereas NLS3 is located near the C-terminal region (residues 587-593) (58) (Fig. 1). Because Nrf2 protein is much larger than the diffusion limit of the nuclear pore complex, it is expected that during the nuclear translocation process, Nrf2 molecules are recognized in the cytoplasm, through their nuclear localization signals (NLS1, NLS2, NLS3) by the soluble adaptor proteins

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Fig. 1. Schematic representation of the conserved regions in Nrf2. Structurally, Nrf2 comprises six regions, called Neh (Nrf2-ECH homology) 1–6 domains, which are highly conserved across different species. Neh 1 contains the CNC-bZIP region, which promotes dimerization partners and confers DNA-binding specificity. The Neh4 and Neh5 domains act cooperatively to bind the coactivator CBP [CREB (c-AMP-response element-binding protein)/ATF4] and BRG1 (Brahmarelated gene 1) thereby, activating transcription (45, 46). Neh3 is a C-terminal domain and also contributes to Nrf2 transactivation (47). The previously uncharacterized, redox-insensitive Neh6 domain (amino acids 329–379) is essential in the Keap1-independent degradation of Nrf2 that occurs in the nucleus of oxidatively stressed cells (48). Of particular interest is the N-terminal region of 100 amino acids, called the Neh2 domain, which contains both the DIDLID/DLG element (amino acids 17–32) and the ETGE tetrapeptide motif (amino acids 79–82), and negatively downregulates nrf2 function under homeostatic conditions. The Nrf2 protein also contains nuclear import (NLSs) and export signals (NESs) which regulate Nrf2 shuttling in and out of the nucleus (55, 58–60). A canonical redox-insensitive NES has been found within the leucine zipper (ZIP) domain of the Nrf2 protein (53). With regard this NESZIP , there has been some confusion in the literature because two redox-insensitive NESs have been found. In fact, the NES reported as 537LKKQLSTLYL546 (53) and the NES reported as 545LKRRLSTLYL554 (55) both refer to the same region of homology in Nrf2 from two different species, human and mouse. An additional redox-sensitive NESTA reported as 175LLSIPELQCLNI186 has been found in the Neh5 transactivation (TA) domain of Nrf2 (54). In addition to the two NESs, which interact with exportins, Nrf2 also contains multiple nuclear localization signals (NLSs) which likely facilitate Nrf2 nuclear localization upon addition of ARE inducers (55, 58–60). These monopartite sequences are designated NLS1 and NLS3, respectively (58), to distinguish them from a previously identified bipartite sequence termed NLS2 (494-511 residues), which had been implicated in the nuclear translocation of this transcription factor (55). NLS1 occurs within the Neh2 domain of Nrf2 (amino acid residues 42-53) whereas NLS3 is located near the C-terminal region (residues 587-593) (58). Adapted from McMahon et al. (48); Jain et al. (54); Giudice and Montella (5); Theodore et al. (58); Eggler et al. (51).

termed importins/karyopherins (a and/or b) which upon binding the cargo proteins such as Nrf2, result in a complex that is then ferried through the nuclear pore complex in the nuclear membrane into the nucleoplasm (58–60). Given that there are up to six isoforms of importins a in mammalian cells (62) further studies are required to determine whether other importins participate in binding to, or show selectivity in binding to Nrf2, during its nuclear translocation. Keap1, the other member of the cytoplasmic oxidative stress system, is a cytoplasmic inhibitor of Nrf2, homologous to the Kelch protein that binds actin in Drosophila (23, 24). Biochemical analysis has revealed that the amino acid sequences in mouse, rat, and human Keap1 proteins are highly conserved between these species. Structurally, mouse Keap1 protein consists of 624 amino acids organized into five domains: (1) the N-terminal region (NTR, amino acids 1–60), (2) the BTB/POZ domain[(Broad complex, Tramtrack, and Bric-a-Brac)/(poxivirus and zinc finger)], which is present in actin-binding proteins and mediates Keap1 homodimerization as well as Nrf2 polyubiquitination and the subsequent 26S proteasome-mediated degradation in basal (reducing) conditions (3),

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the intervening region or the linker region (IVR), (amino acids 180–314), an especially cysteine-rich region (eight cysteine residues in 102 amino acids), (4) the double glycine repeats (DGR) or Kelch repeats region (amino acids 315–598), comprising six repeats of the Kelch motif which create multiple protein contact sites (it is the DGR domain of Keap1 that combines with Nrf2’s Neh2 domain [the N-terminal region of 100 amino acids]), and (5) the C-terminal region (CTR, amino acids 599–624) (Fig. 2) (5, 8). Accumulating evidence suggests that three specific cysteines, one in the BTB region (C151) and two in the IVR (C273, C288) are required for Nrf2 regulation. C273 and C288 are required for Keap1-mediated ubiquitination of Nrf2, whereas C151 is required to release Nrf2 from this pathway (63). It was also shown that Keap1 functions as a bridge between Nrf2 and the Cul3-based E3 ubiquitin ligase that targets lysine residues within the Neh2 domain for ubiquitin conjugation (64, 65). Ubiquitin conjugation onto specific N-terminal lysine residues marks Nrf2 for degradation by the 26S proteasome, such that Nrf2 is maintained at low steady-state levels under basal conditions (64). Recently, Keap1 was also reported to have nucleocytoplasmic shuttling capacity. This has opened a new forum for investigation, particularly after Keap1 was found to contain an

Fig. 2. Schematic representation of the conserved regions in Keap1 protein. Biochemical analysis has revealed that the amino acid sequences in mouse, rat, and human Keap1 proteins are highly conserved between these species. Structurally, mouse Keap1 protein consists of 624 amino acids organized into five domains: (1) the N-terminal region (NTR, amino acids 1–60); (2) the BTB/POZ domain[(Broad complex, Tramtrack, and Bric-aBrac)/(poxivirus and zinc finger)], which is present in actin-binding proteins and mediates Keap1 homodimerization as well as Nrf2 polyubiquitination and the subsequent 26S proteasome-mediated degradation in basal (reducing) conditions; (3) the intervening region or the linker region (IVR), amino acids 180–314), an especially cysteine-rich region (eight cysteine residues in 102 amino acids); (4) the double glycine repeats (DGR) or Kelch repeats region (amino acids 315–598), comprising six repeats of the Kelch motif which create multiple protein contact sites (it is the DGR domain of Keap1 that combines with Nrf2’s Neh2 domain [the N-terminal region of 100 amino acids]); and (5) the C-terminal region (CTR, amino acids 599–624). Sources: Talalay et al. (8); Giudice and Montella (5).

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NES sequence within its linker region or IVR (66, 67). Since a nuclear localizing signal (NLS) sequence has not been found in Keap1, it is believed that Keap1 protein is coupled to Nrf2 and enters the nucleus by means of the NLS in Nrf2 (67). In this model, Nrf2 degradation by proteasome system is considered to occur primarily in cytoplasm, in spite of the Keap1-Nrf2 coupled nuclear entry. Conversely, Nguyen et al. (68) proposed that Nrf2 enters the nucleus by a coupled translation and translocation mechanism. In the latter model, Keap1 enters the nucleus transiently, where it mediates ubiquitination and degradation of Nrf2, but the Keap1-dependent Nrf2 degradation in the cytoplasm is excluded. One obvious question for this model is the mechanism by which Keap1 translocates into the nucleus without any NLS sequences. Recently, Watai et al. (69) have shown that Keap1 protein, under normal, unstressed condition is localized primarily in the cytoplasm with minimal amount in the nucleus and endoplasmic reticulum (RE). This subcellular localization profile of Keap1 appears unchanged after treatment of cells, with diethyl maleate, an electrophile, and/or leptomycin B, a nuclear export inhibitor. These results collectively indicate that endogenous Keap1 remains mostly in the cytoplasm, and electrophiles promote nuclear accumulation of Nrf2 without altering the subcellular localization of Keap1. Currently, it is not known how Keap1 travels into the nucleus. It is possible that Keap1 contains a redoxsensitive NLS that is activated upon recovery of intracellular redox homeostasis during the postinduction stage, in addition to its low rate of constitutive trafficking (70). Alternatively, the rate of Keap1 shuttling between the nucleus and the cytoplasm is constant, regardless of the intracellular redox conditions. In this scenario, the activity of Keap1-Cul3-Rbx1 E3 ubiquitin ligase is the only step that is controlled by intracellular redox conditions. Currently there are no data in favor of either hypothesis. Clearly, understanding the nuclear import mechanism of Keap1 will greatly aid our knowledge of how Keap1 regulates the Nrf2dependent antioxidant response. Intriguingly, more recent studies by Lo and Hanning (71) have also demonstrated the existence of a ternary complex containing PGAM5, a member of the phosphoglycerate mutase family, Keap1 and Nrf2 that is localized to mitochondria. It is expected that this ternary complex provides a molecular framework for understanding how nuclear anti-oxidant gene expression is regulated in response to changes in mitochondrial functions. In summary, Keap1 appears to serve as a core component in the regulation of Nrf2, providing several functions; as a scaffold to anchor Nrf2 with the cytoskeleton filaments, as a Cul3 substrate adaptor to bring Nrf2 into the Cul3-dependent E3 complex for ubiquitination of Nrf2, as a sensor to interact with oxidative/electrophilic stimuli for induction of target genes, as a target substrate for Keap1 ubiquitination and degradation by

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a proteasome-independent pathway, as a nucleocytoplasmic shuttling protein and finally as a dimeric protein capable of binding simultaneously both PGAM5 and Nrf2 through their conserved E(S/T)GE motifs . This leads to the formation of a ternary complex (PGAM5-Keap1-Nrf2) that is localized to mitochondria and regulates nuclear anti-oxidant gene expression in response to changes in mitochondrial functions.

4. The Role of the Cytoplasmic Oxidative Stress System (Nrf2–Keap1) Under Basal (Reducing) Conditions

Various studies have shown that Keap1 plays an essential role in the Nrf2–Keap1 stress response system, not only as a sensor of oxidative and electrophilic stresses but also as a regulator of Nrf2 degradation by the ubiquitin (Ub)-proteasome proteolysis system (72). Under basal (reducing) conditions, Keap1 binds very tightly to Nrf2, anchoring this transcription factor within the actin cytoplasm, targeting it for ubiquitination and proteasome degradation, thus repressing its ability to induce phase II genes (73). This repression is especially important in avoiding unnecessary gene activation in the absence of stress stimuli (74). Although the physical restriction of Nrf2 is an important aspect of its repression by Keap1, this cannot fully account for the relatively short-half life of the transcription factor Nrf2 (10–30 min) in the absence of cellular stress (75). Subsequent experimental studies have shown that Keap1, similarly to other Bric-a-brac, BTB family proteins, functions as a substrate adaptor protein for a Cul3-dependent E3 ubiquitin ligase that targets lysine residues within the Neh2 domain for ubiquitin conjugation (64, 65). Ubiquitin conjugation onto specific N-terminal lysine residues marks Nrf2 for degradation by the 26S proteasome, such that Nrf2 is maintained at low steady-state levels under basal conditions (64). The Ub-dependent degradation of regulatory proteins plays important roles in the control of various physiological processes such as cell cycle and signal transduction (76). To target a protein for degradation by the proteasome, eukaryotic cells attach a polyubiquitin chain to the substrate through a three-enzyme cascade involving the ubiquitin-activating enzyme (E1), a ubiquitin-conjugating enzyme (E2), and a ubiquitin-protein ligase (E3). The cullin family proteins are essential components in a group of multisubunit E3 ubiquitin ligases and associate with the RING finger protein Rbx1(also known as Roc1 and Hrt1) to form the integral core (the catalytic component of the enzyme complex) (77). One of the best characterized RING E3 ligases is the SCF complex that targets the ubiquitination of various proteins involved in cell cycle control and signal transduction. The SCF complex is a multisubunit ubiquitin ligase composed of three invariant subunits,

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Cul1, Rbx1 (also known as Roc1 or Hrt1) and Skp1 (77) and a variable F box protein subunit. Cul1 and Roc1 form the catalytic core of the complex, while Skp1 serves as an adaptor, docking different F box protein subunits to the E3 complex (78). Members of the F box protein family all share an N-terminal Skp1-binding F box motif and a C-terminal protein–protein interaction domain, which is able to recruit one or more specific protein substrates. The large number of F box proteins in eukaryotes, with more than 60 members in mammalian, allows many substrates to be specifically ubiquitinated by the E3 catalytic core, recognized and degraded by the 26S proteasome (79). The Cul3 protein (Cullin3) is a core scaffolding protein in the E3 ligase complex that regulates Nrf2. Cul3 can interact with both Keap1 and Rbx1/ Roc1(Ring box1) (65). In addition, Cul3 and Rbx1 form the catalytic component of the enzyme complex and interact with an E2 ubiquitin ligase to transfer ubiquitin to the substrate (e.g., Nrf2). A model proposed by Cope and Deshaies (80) suggests cullin-dependent ubiquitin ligases are very dynamic structures regulated by cycles of assembly and disassembly for efficient degradation of Nrf2. A central feature of this model is that the cullin– Rbx1core complex cycles between active and inactive states. A large body of evidence suggests that cyclical assembly and disassembly of cullin-dependent E3 ubiquitin ligase complex is mediated, in part, by the antagonistic actions of Nedd8 modification of the cullin protein and association of cullin proteins with CAND1, the cullin-RING ligase (CRL) assembly inhibitor (79, 81, 82). In the active complex, the cullin protein is modified by Nedd8 conjugation. In addition, the conjugated Nedd8 polypeptide may also stimulate CRL-catalyzed ubiquitin transfer from E2 to targets and prevent binding of the inhibitor CAND1 (83, 84). The model proposes that the active complex is converted to an inactive complex in two steps. During the first step the ubiquitinlike protein Nedd8/Rub1 is removed from the cullin protein by one or more deneddylases, such as the CSN5 subunit of the COP9 signalosome (85). The available evidence indicates that the COP9 signalosome (CSN) is a multifunctional protein complex comprised of eight subunits, Csn1–Csn8, that can bind, cleave or deneddylate Nedd8-Cul1 conjugates, and modulate the activities of Cul1, Cul3, and Cul4-based ubiquitin ligases (86). However, it was also shown that the cycles of neddylation and deneddylation may be needed to sustain optimal SCF activity (87). The second step is the association of a protein known as CAND1 (also termed TIP120A) with the deneddylated cullin protein CUL1 but not the neddylated one (88). CAND1 binds to several different human cullin proteins, including Cul 3 and blocks binding of the substrate adaptor protein (89). Recently Min et al. (81) have also suggested that enhancement of CSN-mediated deneddylation by CAND1 may contribute to its

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function as a positive regulator of SCFs in vivo. Subsequent conjugation of Nedd8 onto the cullin subunit by Ubc12, a Nedd8specific E2 enzyme (80) is proposed to decrease the affinity of CAND1 for the cullin protein, enabling another substrate adaptor protein to displace CAND1 and initiate another cycle of substrate ubiquitination (80, 82). Taken together, these results are consistent with a model in which the ability of Keap1, to participate in multiple cycles of substrate adaptor exchange is a critical regulatory aspect of Keap1-mediated repression of Nrf2-dependent gene expression. Then, a decreased ability of Keap1 to target Nrf2 for ubiquitin-dependent degradation results in the accumulation of an excess of Nrf2 relative to Keap1 such that free Nrf2 proteins are able to localize to the nucleus and activate Nrf2dependent gene expression. A key feature of this model is that physical release of Nrf2 from Keap1 is not required for activation of Nrf2-dependent transcription. In fact, reactive molecules do not cause the physical release of Nrf2 from Keap1 but interfere with the ability of Keap1 to act in a catalytic manner to efficiently target Nrf2 for ubiquitin-dependent degradation. Accordingly, knockdown of CAND1 markedly increases the level of Keap1associated Nrf2 yet also increases Nrf2-dependent transcription. On the contrary, ectopic expression of CAND1 reduced the level of complex formation between Keap1 and Cul3, while siRNAmediated knockdown of endogenous CAND1 expression increased complex formation between Keap1 and Cul3. Notably, a marked increase in Nrf2-dependent gene expression was observed following siRNA-mediated knockdown of CAND1 expression (90). Previous studies by Wakabayashi et al. (91) suggested that under basal (reducing) conditions, Keap1 appears to occur as a dimer in which two monomers are bound to each other, conceivably through their BTB domains and anchored to the actin cytoskeleton via DGR region. The reactive cysteine thiol groups (C273, C288) located in the IVR are in reduced state. In this conformation, Keap1 retains one Nrf2 molecule between two DGR domains in the cytoplasm, assuring Nrf2’s rapid turnover, targeting it to the proteasome. Upon exposure to inducers such as 3H-1,2dithiole-3-thione (D3T) or sulforaphane (SUL), the reactive C273 and C288 residues form intermolecular disulfide bonds, probably between the C273 of one Keap1 molecule and the C288 of another, resulting in a conformational change in Keap1, rendering it unable to bind to Nrf2, thus translocating Nrf2 to the nucleus where it heterodimerically partners with other transcription factors like sMaf, and then binds to the ARE regulatory region of the phase II genes, enhancing their transcription. Other authors also suggested that Keap1 exists as a dimer in mammalian cells and binds to a single molecule of Nrf2 in this form (49, 50). Recently, Tong et al. (52) proposed a “hinge and

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latch” interaction model, indicating that two distinct Keap1binding sites within the Neh2 domain of Nrf2, the conserved 29 DLG31 and 79ETGE82 motifs, make contact with a single overlapping site, comprising conserved arginine, serine, and asparagines residues, in the double glycine repeat ( DGR) domain of Keap1 (35, 49, 50, 92). The current interaction model indicates that, under basal condition, binding through the high-affinity ETGE motif provides the “hinge,” through which Nrf2 can move in space relatively freely. Concomitant binding via the low-affinity DGL motif provides the “latch,” which tightly restricts Nrf2 to enable optimal positioning of target lysines for conjugation with ubiquitin, thus directing Nrf2 for proteasomal degradation (49, 50). Under conditions of chemical/oxidative stress, binding via the low-affinity DGL latch, as well as Nrf2 ubiquitination are perturbed, perhaps through the phosphorylation of Nrf2 and/or a conformational change in Keap1 brought about through the modification of one or more cysteine residues. Due to the consequent improper spatial positioning of target lysines, Nrf2 is no longer directed for degradation, but remains associated through the high-affinity ETGE hinge. This leads to the saturation of Keap1, such that any new synthesized Nrf2 can evade repression and accumulate within the nucleus, leading to the transactivation of ARE-regulated target cytoprotective genes, which serve to detoxify the triggering cellular stressors. Although the Keap1 anchoring models seem to successfully explain the repression and activation of Nrf2 signaling in response to the changing redox conditions some controversial observations are reported recently. It is suggested that the cytosolic distribution of Keap1 is maintained by active nuclear export rather than cytoskeleton anchoring (67). According to this notion, the same authors have shown that Keap1 sequesters Nrf2 in the cytoplasm, not by docking it to the actin cytoskeleton but instead via an active Crm1/exportindependent nuclear export mechanism. They have also revealed that the IVR domain of Keap1 contains an NES between amino acids 272 and 312 with a conserved leucine-rich sequence (amino acids 301–310) similar to that seen in other proteins exported by Crm1/Exportin (93). Accordingly, deletion of the NES region results in nuclear accumulation of both Keap1 and Nrf2 (63). A similar outcome is seen after inactivation of the Crm1/exportin pathway by leptomycin B (LMB), a specific inhibitor of CRM1, suggesting that nuclear export is the primary mechanism for cytoplasmic sequestration of Nrf2 (67). Recently, Salazar et al. (94) have demonstrated that Nrf2 is a substrate for glycogen synthase kinase-3b (GSK-3b), and it promotes cytoplasmic localization of Nrf2. It is not yet known whether direct phosphorylation of Nrf2 by GSK-3b under basal conditions (in the absence of ARE inducers) promotes nuclear export or inhibit nuclear import, or

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the identity of the site of phosphorylation. However, Li et al. (54) have also proposed that during basal conditions, the combined nuclear exporting forces NESTA and NESzip of Nrf2 counterbalance the nuclear importing force of the bNLS motif and sequester Nrf2 in the cytoplasm.

5. Effects of ARE-InducersDependent Posttranslational Modifications (Modification of Keap1 Cysteines, Phosphorylation) on the Cul3–Keap1–Nrf2 Complex and Ubiquitination of Nrf2 and Keap1

5.1. Modification of Keap1 Cysteines (C257, C273, C258, C297 and in Particular C151) by ARE Inducers and Disruption of the Keap1–Neh2 Complex

It is widely recognized that under normal conditions, Keap1 anchors the Nrf2 transcription factor within the cytoplasm targeting it for ubiquitination and proteasomal degradation to maintain low levels of Nrf2 that mediate the constitutive expression of Nrf2 downstream genes. When cells are exposed to chemopreventive agents (ARE inducers) and oxidative stress, a signal involving phosphorylation and/or redox modification of critical cysteine residues in Keap1 inhibits the enzymatic activity of the Keap1Cul3-Rbx1 E3 ubiquitin ligase complex, resulting in decreased Nrf2 ubiquitination and degradation. As a consequence free Nrf2 translocates into the nucleus and in combination with other transcription factors (e.g., sMaf, ATF4, JunD, PMF-1) transactivates the AREs/electrophile response elements (EpREs) of many cytoprotective genes, as well as Nrf2 itself (5). Increasing attention is being focussed on the molecular mechanisms of the effects of ARE chemopreventive inducers on the activity of the Cul3Keap1-Nrf2 complex. So far, two general mechanisms for Nrf2 nuclear accumulation in response to ARE inducers have been identified. The first is downregulation of Nrf2 ubiquitination, proposed to occur via disruption of the Keap1–Cul3 and Keap1– Nrf2 complexes, and the other is alteration of the nuclear import/ export of Nrf2 (51). Importantly, modification of Keap1 cysteine residues (e.g., oxidation, alkylation) and phosphorylation of Nrf2 have both been suggested to alter the protein–protein interactions within this complex (5, 51). The following sections outline the current understanding of these mechanisms, with a focus on mechanistic studies of ARE induction by various natural chemopreventive agents. Human Keap1 contains 27 cysteine residues, 25 of which are highly conserved across species. Due to its high cysteine content, Keap1 protein has been proposed to act not only as a regulator of Nrf2 degradation by the Ub-proteasome proteolysis system but also as a sensor of oxidative and electrophilic stresses (23). There are probably several types of interactions between these ARE inducers and the reactive thiol groups of Keap1, due to their extraordinary chemical diversity. Many phase II gene inducers belong to a variety of chemical classes, which have apparently few

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similarities except for their ability to modify the sulfhydryl groups either by oxidation, reduction or alkylation (8, 72). Accordingly, Dinkova-Kostova et al. (72) showed that certain highly reactive cysteine thiol groups (C257, C273, C288, and C297) located in the IVR or linker region of Keap1 in its reduced state are probably the primary cellular sensors that recognize and react with the phase II gene inducers. Although these four cysteines may not be the only ones that are most reactive in vivo, their modification could lead to substantial conformational changes in Keap1, resulting in its dissociation from Nrf2. Zhang and Hannink (63) demonstrated that the C273 and C288 cysteine residues are critical in Keap1-dependent ubiquitination and proteasome-mediated degradation, as well as in the Keap1-mediated repression of Nrf2 under basal (reducing) conditions. Because Keap1’s ability to bind to Nrf2 is regulated by critical cysteine residues, perhaps increased levels of GSH, thioredoxin, GR, and TR as part of the phase II response could provide a regeneration system for reduced Keap1 (72). The same group also showed that a third cysteine residue, C151, located in the BTB domain of Keap1, is especially needed by Nrf2 to escape Keap1-mediated repression in response to tBHQ-induced or SUL-induced oxidative stress. Perhaps prior modification of C151 is needed to induce a conformational change that would alter the accessibility of C273 and C288 to the cytoplasmic environment (63). Recently, Wakabayashi et al. (91) demonstrated that C273 and C288 are the critical sensors that are modified by phase II enzyme inducers, leading to the dissociation of the Nrf2–Keap1 complex. Under basal (reducing) conditions, Keap1 appears to occur as a dimer in which two monomers are bound to each other, conceivably through their BTB domains and anchored to the actin cytoskeleton via DGR region. The reactive cysteine thiol groups (C273, C288) located in the IVR are in reduced state. In this conformation, Keap1 retains one Nrf2 molecule between two DGR domains in the cytoplasm, assuring Nrf2’s rapid turnover, targeting it to the proteasome. Upon exposure to inducers such as 3H-1,2dithiole-3-thione (D3T) or sulforaphane (SUL), the reactive C273 and C288 residues form intermolecular disulfide bonds, probably between the C273 of one Keap1 molecule and the C288 of another, resulting in a conformational change in Keap1, rendering it unable to bind to Nrf2, thus translocating Nrf2 to the nucleus where it heterodimerically partners with other transcription factors like sMaf, and then binds to the ARE regulatory region of the phase II genes, enhancing their transcription. The mutation of C273 and C288 to serine, which also renders Keap1 unable to prevent the nuclear translocation of Nrf2, further indicates the importance of these residues in inducer response. Recently, other groups have found that C151 cysteine residue is one of the most reactive in the human Keap1 protein in vitro and the only cysteine highly

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modified in vitro by natural chemopreventive agents such as xanthohumol, isoliquiritigenin, and 10-shogaol (95–98). Taken together, these results suggest that modification of Keap1 cysteines, and in particular C151, by inducers likely impairs the ability of Keap1 to efficiently ubiquitinate Nrf2 and target it for degradation. 5.2. Modification of Keap1 Cysteines by ARE Inducers and Disruption of the Keap1–Cul3 Interaction

There is still little known about the mechanism by which Keap1 cysteine modifications lead to the downregulation of Nrf2 ubiquitination. Previous studies by Dinkova-Kostova et al. (72) proposed a model in which modification of Keap1 cysteines by chemopreventive inducers/agents directly alters the interaction between Keap1 and the Neh2 domain of Nrf2. Based on this attractive model, many investigators have incorrectly interpreted Nrf2 translocation and accumulation as resulting from the disruption of the Keap1–Nrf2 interaction and have reported it as such. Subsequent experimental studies by Eggler et al. (95) have shown that in fact disruption of the Keap1–Nrf2 complex does not occur upon modification of Keap1 cysteines. While modification of Keap1 protein cysteines is insufficient to alter the affinity of Keap1 for Nrf2, recent results suggest that Keap1–Cul3 interaction is disrupted by cysteine modification. Accordingly, Zhang and Hannink (64) have shown by co-immunoprecipitation (coIP) assays that less Cul3 precipitated with Keap1 upon exposure of cells to sulforaphane (SUL) or tBHQ. Interestingly, a mutant Keap1 protein containing a single cysteine-to-serine substitution at residue C151 within the BTB domain of Keap1, largely abrogated this effect, again implying a key role for C151 in ARE induction. In agreement with these results, more recent experimental studies by Gao et al. also demonstrated by co-immunoprecipitation that the oxidative products of n-3 fatty acids such as eicosapentaenoic acid (EPAox), a major component of fish oil, destabilized the association between Keap1 and (99)Cul3. The authors also observed that free radical-mediated oxidation products (e.g., a series of novel cyclopentenone-containing molecules termed J3-isoprostanes) reacted with Keap1 sulfhydryls, altering Keap1 structure. This conformational change was associated with loss of binding to Cul3 and increased ARE-directed gene expression. Then, Keap1 BTB and intervening domains are important for association with the E3 ligase scaffold protein Cullin3. Loss of Keap1–Cullin3 association inhibits Nrf2 ubiquitination, thereby stabilizing (activating) Nrf2 and initiating Nrf2-directed gene expression (65, 74). Taken together, these results collectively indicate that reaction of ARE inducers with Keap1 cysteines leads to a reduced association between Keap1 and Cul3, thereby downregulating Nrf2 ubiquitination. This would in turn lead to Nrf2 accumulation and location to the nucleus and increased expression of ARE-controlled gene products.

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5.3. Role of Ubiquitination of Keap1 in Downregulation of Nrf2 Ubiquitination by Cul3 Keap1–Nrf2 Complex and Increased Nrf2-Directed Gene Expression

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Although the primary focus of the field has been on the regulation of Nrf2 ubiquitination due to its important role, the increased ubiquitination of Keap1 and degradation by a proteasome-independent pathway is beginning to receive attention as it may also play an important role in increased Nrf2-directed gene expression. Recent studies suggest that different ARE inducers may exert different effects on Keap1 ubiquitination and stability/ activation of the transcription factor Nrf2. A recent offering by Zhang et al. (99) for the first time proposed that ubiquitination of Keap1 was markedly increased in glutathione-deficient cells exposed to quinone-induced oxidative stress and resulted in increased degradation of Keap1 by a proteasome-independent pathway. Interestingly, the authors observed that quinone (tBHQ)-induced oxidative stress perturbed the Keap1–Cul3– Rbx1 E3 ubiquitin ligase complex such that Keap1, but not Nrf2, became the target for ubiquitin conjugation. Furthermore, this switch in the ubiquitin ligase activity of the Keap1-dependent E3 ubiquitin ligase complex was specific to quinone-induced oxidative stress and not to sulforaphane. Sulforaphane treatment did not result in Keap1 ubiquitination and degradation, indicating that Keap1 differentially responded to inducers of Nrf2-dependent transcription. They also noted that the increase in Keap1 degradation occurred by a C151-independent pathway, different from the decrease in Nrf2 ubiquitination. Other investigators have also suggested effects of ARE inducers on Keap1 ubiquitination and stability. For example, Hong et al. (100) provided compelling support for the hypothesis that electrophilic adduction to Keap1 cysteines triggered a switching of Cul3-dependent ubiquitination from Nrf2 to Keap1, leading to the degradation of Keap1 and to Nrf2 activation in cells exposed to N-iodoacetyl-Nbiotinylhexylenediamine (BIA). The authors identified that the ubiquitination target site on Keap1 was lysine-298 (Lys-298), which lay adjacent to Cys residues in the central linker domain. Recently, He and coworkers (75, 101), also demonstrated that the ubiquitination of cytoplasmic Keap1 increased in the presence of heavy metals such as arsenic and chromium in mouse Hepa1c1c7 cells, at least upon initial exposure to the metal. Keap1 was shown to be ubiquitinated in the cytoplasm and deubiquitinated in the nucleus in the presence of arsenic without changing the protein level, implicating nuclear-cytoplasmic recyling of Keap1. The same group also revealed novel aspects of Nrf2 activation by Cr(VI). Specifically, they demonstrated that Nrf2 and Keap1 were translocated into the nucleus in association with each other. Both proteins were ubiquitinated in the cytoplasm but were deubiquitinated upon nuclear translocation. By analogy with the findings of p53 ubiquitination/deubiquitination, the authors also postulated that a nuclear deubiquitinase interacts with the ubiquitinated Nrf2/Keap1 complex and deubiquitinates the proteins.

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Then, treatment with Cr(VI) but not phenolic antioxidant tert-butylhydroquinone (tBHQ) disrupts the Nrf2/Keap1 association in the nucleus and recruits Nrf2 to the AREs located in the enhancers of Ho-1 and Nqo1 (75, 101). Therefore, in addition to functions of Keap1 described previously (e.g., anchoring Nrf2 in the cytoplasm, chemical sensing, and serving as a substrate adaptor in the Cul3-dependent E3 complex for Nrf2), this study provided evidence for an active role of Keap1 in Nrf2 nuclear translocation and processing. Another mechanism proposed by Tanigawa et al. (102) suggests that treatment of HepG2 cells with quercetin resulted in decreased endogenous Keap1 levels although in this study no change in Keap1 ubiquitination was detected. Taken together, these results suggest the possibility that a subset of ARE inducers increases ubiquitin transfer to the Keap1 protein, resulting in decreased Keap1 proteins levels, which would then lead to increased Nrf2 activation. Further studies are required to determine if Keap1 cysteine modification or other mechanisms are involved in the increase of Keap1 ubiquitination. 5.4. Nrf2/Keap1 Phosphorylation by Protein Kinases and Activation of the Nrf2–Keap1 Complex

Activation of the Nrf2–Keap1 complex (the dissociation of Nrf2 from Keap1 and Nrf2’s subsequent nuclear migration) upon exposure to chemopreventive agents and oxidative stress may involve the modification of either of these proteins, by indirect or direct mechanisms (5). One hypothesis suggests that a possible Post-translational modification (phosphorylation) of this complex by various protein kinase signaling pathways would affect either the liberation process of Nrf2 from Keap1, the stability of Nrf2, or Nrf2’s translocation into the nucleus (5, 51). Three major signal transduction pathways have been proposed as being involved in transducing oxidative stress signals to gene expression, mediated through the ARE: (1) the protein kinase C (PKC), (2) the mitogen-activated protein kinase (MAPK) cascades, and (3) the casein kinase 2 (CK-2). In fact, it is possible that all three signal transmission pathways play an important role in the transcriptional regulation of AREs due to the cross-reactions that exist between them (4, 103–105). Previous studies by various groups have demonstrated that the activity of PKC enzyme could be stimulated by all three of the ARE inducers tested including tBHQ, 4-hydroxynonenal (4-HNE) and phorone (26, 106). They identified the sole site of phosphorylation by PKC as S40. The importance of S40 phosphorylation in mediating Nrf2directed gene expression was shown by experiments in which the nuclear translocation of a Nrf2 S40A mutant protein in response to 4-HNE was greatly decreased compared to wild-type protein (106). The authors also showed that various atypical PKC isoforms are required (106), while other groups have found a novel isoform, PKCd, in mediating Nrf2/ARE-dependent gene expression (107, 108). A previous offering by Huang et al. (109) has

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also demonstrated by co-immunoprecipitation experiments that in vitro (direct) phosphorylation of Nrf2 by PKC induces its dissociation from Keap1, and that this effect is largely inhibited by the S40A mutation. Other findings by Boom and Jaiswal (110) have revealed that phosphorylation of S40 in response to antioxidants is necessary for Nrf2 release from Keap1, but is not required for Nrf2 stabilization/accumulation in the nucleus and transcriptional activation of ARE-mediated NQO1 gene expression. It is unknown if phosphorylation of Nrf2 by PKC is required only for its release from Keap1/INrf2 or is required also for stabilization/ accumulation of Nrf2 in the nucleus and transcriptional activation of Nrf2/ARE-mediated gene expression. Further studies are required to determine the importance of PKC phosphorylation of S40 in Nrf2-directed gene expression by ARE inducers or on Nrf2 ubiquitination or Keap1 ubiquitination. Another kinase pathway involving p38 mitogen-activated protein kinase (MAPK) isoforms has also been implicated in Nrf2/ARE-directed gene expression, and alteration of the Nrf2-Keap1 affinity has been proposed as the mechanistic explanation. Recently, Keum and Hong (111) demonstrated that p38 MAPK isoforms (p38a p38b, p38g and p38d), were able to phosphorylate purified Nrf2 transcription factor. This in turn promoted its interaction with Keap1 protein that co-immunoprecipitated with Nrf2, thereby contributing to a suppression of Nrf2 nuclear translocation. Importantly, they also showed that sulforaphane not only activated MAP/extracellular signal-regulated kinase (ERK) kinases 1/2 and ERK1/2, but also strongly suppressed anisomycin-induced activation of p38 MAPK isoforms by blocking phosphorylation of upstream kinases, MKK3/6. Collectively, these results indicate that sulforaphane is able to inhibit the phosphorylation of Nrf2 by blockade of p38 MAPK signaling, resulting in a reduced association between Nrf2 and Keap1, and subsequent Nrf2 activation. Accordingly, Cullinan et al. (112) have also shown that Nrf2 could be directly phosphorylated by PKR-like endoplasmic reticulum-resident kinase (PERK), although its targets sites are not yet identified. In the present study, the authors observed that stimulation of p38 MAPK isoforms directly phosphorylated Nrf2 protein and phosphorylation of Nrf2 protein by activated p38 d promoted the association between Nrf2 and Keap1 proteins. Based on these data, it has been speculated that phosphorylation of Nrf2 by p38 MAPK could contribute to inhibition of AREdependent gene expression by increasing the protein–protein interaction between Nrf2 and Keap1. They also identified a unique phosphorylation site by p38d, but not by ERK2 and JNK1, at the COOH terminus of Nrf2. Further studies by other groups have demonstrated that p38 MAPK activation, rather than p38 MAPK inhibition, leads to induction of Nrf2/ARE-directed gene expression (113–115). One possible explanation for different

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results is that while certain ARE inducers (e.g., pyrrolydine dithiocarbamate, cadmium) may activate p38 MAPK, other including the isothiocyanate sulforaphane may inhibit p38 MAPK. Recently, it was also shown by co-IP assays that Keap1 and Nrf2 proteins remain associated after treatments of cells with various ARE inducers such as sulforaphane, tBHQ or quercetin (64, 102, 116). According to this notion, He and coworkers (75, 101) demonstrated by co-IP assays that Nrf2 translocates into the nucleus together with Keap1 and Cul3 and that the heavy metals such as arsenic and chromium dissociate nuclear Nrf2 from Keap1 and Cul3 and activate Nrf2/ARE-mediate gene expression. With regards to the tBHQ, conflicting results were observed in these two studies (75, 101). Taken together, these conflicting results suggest that co-IP is a useful but imprecise assay to determine comparative affinities of protein–protein interactions, and it is difficult to ascertain whether partial disruption of the Nrf2–Keap1 complex occurs in response to ARE inducers in the cellular context. In addition, as the Keap1 protein contains potential phosphorylation sites, it is also possible that Keap1 may be a target for phosphorylation (4). It is important to note that downregulation of Nrf2 ubiquitination is sufficient to induce Nrf2 translocation and subsequent Nrf2/ARE-mediated gene expression, as shown by treatment cells with the proteasome inhibitor MG132 (63). Another mechanism whereby Nrf2 phosphorylation by protein kinases regulates Nrf2 localization and degradation was recently and simultaneously proposed by Pi et al. (104) and Apopa and coworkers (105). They reported that phenolic antioxidant/prooxidant tBHQ induced two forms of the Nrf2 protein in neuroblastoma cells (IMR-32), which migrated as distinctive bands on SDS-PAGE. Unphosphorylated Nrf2 predominated in the cytoplasm, whereas the phosphorylated form preferentially localized in the nucleus. Nuclear Nrf2 could be dephosphorylated by l phosphatase in vitro and be converted to the faster migrating form, involving phosphorylation of Nrf2 in the nuclear translocation and activation of Nrf2 dependent gene expression. They also revealed by deletional analyses the transcription activation (TA) domains Neh4 (Nrf2-ECH homology 4) and Neh5 (Nrf2-ECH homology 5) as a major region necessary for the phosphorylation by casein kinase 2 (CK2). However, treatment with CK-2 inhibitor 2-dimethylamino-4,5,6,7,-tetrabromo-1H-benzimidazole (DMAT) blocked the induction of endogenous target genes of Nrf2 in cells and inhibited the TA activities of both the full length and the TA domains of Nrf2 to a large extent. Collectively, these results, together with those proposed by Pi and coworkers, suggest that sequential phosphorylation of Nrf2 by Ca2+-dependent calmodulin (CaM) regulated protein kinase CK-2 plays an important role not only in the activation and nuclear translocation of Nrf2 but also in the subsequent hyper-phosphorylation of Nrf2.

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This CK-2 function decreases Nrf2 transcriptional activity and helps nuclear Nrf2 translocate back to the cytoplasm for Keap1mediated degradation (104, 105). In summary, downregulation of Nrf2 ubiquitination is an essential factor in mediating Nrf2/ ARE signaling, and changes in Keap1 ubiquitination may play an important role as well. Further researches are required to determine how Keap1 cysteine modification might lead to changes in ubiquitination of Nrf2 and possibly Keap1. Alteration of the Keap1–Cul3 interaction seems to be a likely mechanism, while disruption of the Keap1–Nrf2 complex by Keap1 cysteine modification has been discarded. Alteration of the Keap1–Nrf2 interaction by Nrf2 phosphorylation, leading to decreased Nrf2 ubiquitination, is also an attractive mechanism requiring further research.

6. Alteration of Import and Export of Nrf2 by ARE Inducers 6.1. Nrf2 Phosphorylation Events by Chemopreventive Agents and Nrf2 Localization

A large body of evidence indicates that Nrf2 nuclear localization and accumulation in response to ARE inducers is clearly a result of Nrf2 ubiquitination downregulation, proposed to occur by disruption of the Nrf2–Keap1 and Keap1–Cul3 complexes (51). In addition, recent findings also suggest that nuclear import and export signals of Nrf2 play an key role in the mechanisms governing Nrf2 nuclear localization in response to ARE inducers (51). Virtually little is known about Nrf2 nuclear translocation after it is released from Keap1. In recent years, the discovery of multiple NLSs motifs and NESs in Nrf2 demonstrates that Nrf2 nuclear translocation is not a passive or an automatic process. In fact, it is expected that during the nuclear translocation process, Nrf2 molecules are recognized in the cytoplasm, through their NLS1, NLS2, NLS3 by the soluble adaptor proteins termed importins/ karyopherins (a and/or b) which upon binding the cargo proteins, such as Nrf2, result in a complex that is then ferried through the nuclear pore complex in the nuclear membrane into the nucleoplasm (58–60). Given that there are up to six isoforms of importins a in mammalian cells (61, 62), further studies are required to determine whether other importins participate in binding to, or show selectivity in binding to Nrf2, during its nuclear translocation. These observations collectively suggest that the nucleocytoplasm translocation of transcription factors is the consequence of a dynamic equilibrium of multivalent NLS and NES (58–60). So far, both Nrf2 phosphorylation events and Nrf2 cysteine modification have been proposed to affect Nrf2 nuclear localization. With regards to the importance of Nrf2 phosphorylation, two Nrf2 phosphorylation events have been recognized to modulate Nrf2 nuclear import/export in response to various

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chemopreventive agents (51). First, Salazar and Cuadrado (94) have proposed a mechanism involving direct phosphorylation of the transcription factor Nrf2 by GSK-3b, downstream of PI3K/ Ser/Thr kinase Akt pathway. They observed that induction of Nrf2/ARE-mediated gene expression and long-term antioxidant effect of carnosol, a diterpene derived from the herb rosmary, that induces the expression of phase II enzymes such as HO-1, GST, were dependent on PI3K and the Akt1 kinase, downstream from PI3K (116). Interestingly, the same group has also demonstrated that GSK-3b was a negative regulator of Nrf2 activity which directly phosphorylated the transcription factor Nrf2, downstream of PI3K/Ser/Thr kinase Akt pathway. Other groups have noted that the PI3K inhibitor treatment did not appear to affect Nrf2 degradation, but it did decrease nuclear accumulation and localization of Nrf2, indicating that PI3K activation by various ARE inducers results in alteration of the nuclear import/ export of Nrf2 rather than the ubiquitination status of Nrf2 (117, 118). Therefore, it appears that ARE inducers may activate the PI3K/Akt pathway, leading ultimately to Nrf2 nuclear accumulation and localization by downregulating Nrf2 phosphorylation by GSK-3b. It is not yet known whether phosphorylation of Nrf2 by GSK-3b under basal conditions (in the absence of ARE inducers) promotes nuclear export or inhibits nuclear import, or the identity of the site of phosphorylation. A second mechanism whereby Nrf2 phosphorylation regulates Nrf2 localization when cells are exposed to chemopreventive agents and oxidative stress was proposed by Jain and Jaiswal (56). They have described that phosphorylation of tyrosine 568 in Nrf2, by the tyrosine kinase Fyn, is essential for Crm1-mediated nuclear export and degradation of the transcription factor. The mechanism of phosphorylated Nrf2 interaction with Crm1 exportin remains unknown. It is expected that phosphorylation of Nrf2 Y568 leads to structural changes that expose the leucine-rich NES region (amino acid 545-554) for interaction with Crm1. Accordingly, mutation of tyrosine 568 to alanine or phenylalanine resulted in the loss of phosphorylation and interaction of Nrf2 with Crm1 and abrogation of nuclear export of Nrf2. The wild-type Nrf2 and mutant Nrf2Y568A both interacted with Keap1/INrf2 and were released/imported in the nucleus in response to endogenous cellular stressors. In addition, the mutant Nrf2Y568A lacking the tyrosine phosphorylation accumulated in the nucleus due to the loss of nuclear export of mutant protein (56). This was clearly evident from the observations that accumulation of mutant protein inside nucleus was insensitive to nuclear export inhibitor leptomycin B (LMB) and was similar to nuclear accumulation of wild-type Nrf2 protein in response to leptomycin B. The authors have also noted that hydrogen peroxide (H2O2) initially led to nuclear

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accumulation of Nrf2, presumably to activate cytoprotective genes, and later, because persistent increase in chemoprotective genes expression threatens cell survival, induced phosphorylation of tyrosine 568 for enhanced nuclear export of Nrf2. Taken together, these results suggest that phosphorylation of tyrosine 568 in Nrf2 is essential for Crm1-mediated nuclear export and degradation but not for binding of Nrf2 with Keap1, because both wild-type Nrf2 and mutant Nrf2Y568A interacted with Keap1/INrf2 (56). Previous studies by Grimes and Jope (119) have shown that phosphorylation status of GSK-3b regulates its activity. Recently, Jain and Jaiswal (120) demonstrated that GSK-3b acts upstream of Fyn kinase in control of nuclear export and degradation of Nrf2. They also observed that activation of GSK-3b is mediated by phosphorylation at tyrosine 216 residue(s) and/or dephosphorylation of serine 9 via hydrogen peroxide (119). On the contrary, GSK-3b phosphorylated at a serine 9 residue via PKC, Akt or other similar kinases was inactive. The activated GSK-3b phosphorylated Fyn kinase at threonine residue, leading to nuclear localization of Fyn. Based on previous results (56) and the new ones, Jain and Jaiswal (120) proposed an interesting model depicting the role of GSK-3b in regulating nuclear export of Nrf2 via Fyn phosphorylation. This suggests that exposure of cells to chemical stress leads to the release of Nrf2 from its cytosolic inhibitor Keap1 as an early cellular response to chemical, xenobiotic, drugs, UV, and radiation stress. The release of Nrf2 from Keap1 is mediated by PKC phosphorylation and/or cysteine modification of Keap1 (121) Nrf2 contains well-defined signals that control its nuclear import and export (55). Among them, a bipartite NLS directs Nrf2 to the nucleus (55). After nuclear translocation, Nrf2, in combination with other transcription factors (e.g. sMaf, ATF4, JunD, PMF-1) transactivates the AREs/ EpREs of many cytoprotective genes, as well as Nrf2 itself (5). The increase in expression of chemopreventive genes expression neutralizes the chemical stress. Because persistent increase in chemoprotective genes expression threatens cell survival, Nrf2 is exported out of the nucleus and degraded. The nuclear export of Nrf2 is a delayed/late response of cells to oxidative/electrophilic stress, presumably mediated via a leucine-rich NES at the C terminus of Nrf2 (55). However, the NES in Nrf2 is activated only after Fyn kinase is accumulated inside the nucleus that phosphorylates tyrosine 568 of Nrf2 (56). The phosphorylated Nrf2Y568 binds to Crm1 and is exported out of the nucleus (56). The authors in the present report demonstrate that GSK-3b acts upstream of Fyn kinase in control of nuclear export and degradation of Nrf2 (120). They also observed that activation of GSK-3b is mediated by phosphorylation at tyrosine 216 residue(s) and/or dephosphorylation of serine 9 via hydrogen peroxide (119).

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Once GSK-3b is activated, it phosphorylates Fyn kinase at threonine residue, leading to nuclear localization of Fyn. Inside the nucleus, phosphorylated Fyn kinase phosphorylates tyrosine 568 of Nrf2 (56). Phosphorylated Nrf2Y568 is exported out of the nucleus, ubiquitinated, and degraded (Fig. 3) (120). Recently, GSK-3b was also shown to phosphorylate Nrf2 at unknown residues with implications in nuclear export of Nrf2 (94). In summary, the early response leads to nuclear import of Nrf2 resulting in coordinated activation of chemoprotective genes. Conversely, the delayed response to stress is because of the Fyn-mediated phosphorylation of Nrf2Y568 inside nucleus (56). Tyrosine 568 phosphorylation leads to Crm1-mediated nuclear export of Nrf2 (56).

Fig. 3. Model depicting the role of glycogen synthase kinase-3 b (GSK-3b) in regulating nuclear export of Nrf2 via Fyn phosphorylation. This model suggests that exposure of cells to ARE inducers (e.g., low dose of antioxidants/xenobiotics) or oxidative stress leads to release of Nrf2 from its cytosolic inhibitor Keap1/INrf2 as an early cellular response to chemical stress. The release of Nrf2 from Keap1 is mediated by cysteine modification of Keap1 and or by protein kinase C (PKC) (121). Moreover, a bipartite nuclear localization signal (NLS2) localized at amino acid residues 494-511 of Nrf2 protein promotes nuclear translocation of Nrf2 (55). After translocation in the nucleus, Nrf2 in combination with other transcription factors (e.g., sMaf, ATF4, JunD, PMF-1) transactivates the AREs of many cytoprotective genes, as well as nrf2 itself (5). Because persistent increase in chemopreventive genes expression threatens cell survival, Nrf2 is exported out of the nucleus and degraded. The nuclear export of Nrf2 is a delayed/late response mediated by Fyn kinase signaling pathway (56). GSK-3b is upstream to Fyn kinase in regulation of nuclear export of Nrf2 (120). Besides, phosphorylation status of GSK-3b regulates its activity (119). GSK-3b phosphorylated at a serine 9 residue by PKC or other protein kinases is inactive. Conversely, activation of GSK-3b is mediated by phosphorylation at tyrosine 216 residue and/or dephosphorylation of serine 9 via hydrogen peroxide produced in response to chemical stress (119). The activated GSK-3b phosphorylates Fyn kinase at threonine residue(s) leading to nuclear localization of Fyn kinase. In the nucleus, Fyn kinase phosphorylates Nrf2 at tyrosine residue 568 (56). The phosphorylated Nrf2Y568 binds Crm1 exportin and is exported out the nucleus, ubiquitinated and degraded (56). Source: Jain and Jaiswal (120).

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Interestingly, the aforementioned study by Jain and Jaiswal (56) suggests several interesting questions that remain to be answered. For example, is the observed ARE activation by genistein due mainly to its ability to inhibit phosphorylation of Nrf2Y568 by Src kinases, or is it also acting at the level of downregulating Nrf2 ubiquitination? Moreover, might other natural chemopreventive agents similar in structure to genistein act primarily by inhibiting phosphorylation of Nrf2Y568 or downregulating Nrf2 ubiquitination? 6.2. Modification of Nrf2 Cysteines (C183, C506), Nrf2 Nuclear Accumulation, and Transactivation of Target Genes

In addition to Nrf2 phosphorylation, modification of Nrf2 cysteines (C183,C506) may also regulate Nrf2 localization and transactivation of target genes, despite its importance compared to Keap1 cysteine modification in sensing ARE inducers has been disputed (52). Recently, Li et al. (54) identified and characterized a new functional redox-sensitive nuclear export signal (NESTA) sequence (i.e., 175LLSIPELQCLNI186) located in the Neh5 transactivation (TA) domain of Nrf2. Unlike the redox-insensitive NES in the ZIP domain of Nrf2 (NESzip) (53), this NESTA contains a reactive cysteine residue (Cys-183). They also demonstrated that tBHQ and H2O2 were unable to stimulate ARE induction in HeLa cells transfected with the Nrf2 C183 mutant, as compared to wild-type Nrf2. Based on these results, Li et al. (54) proposed that modification of Nrf2 C183 by oxidative stressors and electrophiles abrogated the function of the redox-sensitive NESTA, resulting in nuclear accumulation of Nrf2 and transactivation of target genes. Accordingly, the same authors suggested a hypothetic model of Keap1-independent Nrf2 signaling. This sustains that during basal conditions, the combined nuclear exporting forces NESTA and NESZIP counterbalance the nuclear importing force of the bNLS motif and sequester Nrf2 in the cytoplasm. When cells are exposed to chemopreventive agents and oxidative stress, the reactive cysteine (Cys-183) in the NESTA, can detect the presence of reactive oxygen species (ROS) or reactive nitrogen species (RNS) and inactivate the NESTA. As a consequence, the nuclear importing force mediated by the bNLS prevails and triggers Nrf2 nuclear translocation (54). A large body of evidence suggests that Nrf2 activation can consequently increase the expression and enzymatic activities of g-glutamyl-cysteine synthetase/GST as well as the GSH level in cells (122, 123). Considering the reversible nature of sulfhydryl modification by sulforaphane, the elevated GSH levels may favor the restoration of NESTA activity and trigger Nrf2 nuclear export. Further studies are required to examine this possibility. Then, Keap1 may not be only arresting force to sequester Nrf2 in the nucleus. Previous studies by Bloom and coworkers (124) instead have implicated the cysteine at position 506 in Nrf2 redox regulation

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of ARE-mediated gene expression. Accordingly, the C506S mutation in Nrf2 lowered its affinity for the ARE, leading to decreased expression, and antioxidant induction of NQO1. With regards to the relative importance of Nrf2 cysteine modification in the overall signaling mechanism, Tong et al. (52) assert that the Nrf2 self-redox induction model is not well-validated by other scientific data in the literature. In the experiments by Li and coworkers (54) examining Nrf2 C183 dependence, only the transcription factor Nrf2, and not cytosolic inhibitor Keap1, is overexpressed. However, since Keap1 protein exerts a strong inhibitory effect on the transcription factor Nrf2 nuclear accumulation (125), it is unlikely that Nrf2/ ARE-mediated gene expression would be significantly dependent upon Nrf2 C183 if overexpressed Keap1 proteins were present in those experiments. In support of this idea, an Nrf2 mutant lacking the ETGE motif and the ability to bind Keap1 was shown to no longer be responsive to oxidative stressors and electrophiles in the presence of overexpressed Keap1, even though the rest of Nrf2, including the Neh5 domain, had an intact NES including C183 (48). Finally, there is a general belief that cysteine residues with two flanking basic amino acids exhibit higher reactivity for disulfide exchange than does a cysteine residue with either one or no adjacent basic amino acids. Since Cys183 residue of human Nrf2 does not have any positively charged neighbors, the proposed redox sensitivity of this cysteine in Nrf2 is unlikely to be as dominant as those observed in Keap1, in which many of the 25 cysteine residues are located adjacent to basic amino acids. The cysteines of Keap1 protein, and C151 in particular, have been shown by several groups to play an essential role in signaling, including in the presence of overexpressed Nrf2 (48, 72, 95).

7. Conclusions Epidemiological studies suggest that cancer susceptibility is influenced significantly by diet. A large number of naturally occurring chemicals, as well as synthetic food additives, have been shown to protect against carcinogenesis. The underlying molecular mechanisms by which these compounds influence the development of cancer are poorly understood. It appears that Keap1 retention in the cytosol and subsequent release of Nrf2 is an essential step in the antioxidant response. In this chapter, we have highlighted recent advances in understanding how cancer chemopreventive agents transcriptionally activate the expression of genes encoding phase II and phase III detoxification enzymes as well as antioxidant enzymes and other defensive proteins. So far, two general

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mechanisms for Nrf2 nuclear accumulation in response to ARE inducers have been identified. The first is downregulation of Nrf2 ubiquitination, proposed to occur via disruption of the Keap1– Cul3 and Keap1–Nrf2 complexes, and the other is alteration of the nuclear import/export of Nrf2. Importantly, modification of Keap1 cysteine residues (e.g., oxidation, alkylation) and phosphorylation of Nrf2 have both been suggested to alter the Keap1– Nrf2 and Keap1–Cul3 interactions within the Cul3–Keap1–Nrf2 complex. Previous studies by Dinkova-Kostova et al. (72) proposed a model in which modification of Keap1 cysteines by chemopreventive inducers/agents directly alters the interaction between Keap1 and the Neh2 domain of Nrf2. Based on this attractive model, many investigators have incorrectly interpreted Nrf2 nuclear translocation and accumulation as a resulting from the disruption of the Keap1–Nrf2 interaction and have reported it as such. Subsequent experimental studies by Eggler et al. (95) have shown that in fact disruption of the Keap1–Nrf2 complex does not occur upon modification of Keap1 cysteines. While modification Keap1 protein cysteines is insufficient to alter the affinity of Keap1 for Nrf2, recent results suggest that Keap1–Cul3 interaction is disrupted by cysteine modification. Taken together, these results collectively indicate that reaction of ARE inducers with Keap1 cysteines leads to a reduced association between Keap1 and Cul3, thereby downregulating Nrf2 ubiquitination. This would in turn lead to Nrf2 accumulation and location to the nucleus and increased expression of ARE-controlled gene products. Convincing evidence also suggests that increased ubiquitination of Keap1 at lysine-63 residue and proteasome-independent degradation of Keap1 may play an important role in increased Nrf2-directed gene expression. However, alteration of the Keap1–Nrf2 complex by Nrf2 phosphorylation, leading to decreased Nrf2 ubiquitination, is also an attractive mechanism requiring further investigation. In summary, downregulation of Nrf2 ubiquitination is a key factor in mediating Nrf2/ARE signaling, and changes in Keap1 ubiquitination may play a role as well. Further researches are required to determine how Keap1 cysteine modification might lead to changes in ubiquitination of Nrf2 and possibly Keap1. Alteration of the Keap1–Cul3 interaction seems to be a likely mechanism, while disruption of the Keap1–Nrf2 complex by Keap1 cysteine modification has been discarded. Alteration of the Keap1–Nrf2 interaction by Nrf2 phosphorylation, leading to decreased Nrf2 ubiquitination, is also an attractive mechanism requiring further studies. In addition to Nrf2 ubiquitination downregulation, proposed to occur via disruption of the Keap1– Cul3 and Keap1–Nrf2 complexes, recent findings also suggest that nuclear import and export signals of Nrf2 play an key role in the mechanisms governing Nrf2 nuclear localization in response to ARE inducers. Both Nrf2 phosphorylation events and Nrf2

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cysteine modification have been proposed to affect Nrf2 nuclear localization. So far, the mechanistic studies of Nrf2 activation were mainly focused on Keap1. Virtually little is known about Nrf2 nuclear translocation after it is released from Keap1. In recent years, the discovery of multiple NLSs motifs and NESs in Nrf2 suggests that Nrf2 nuclear translocation is not a passive or an automatic process (58). In fact, it is expected that during the nuclear translocation process, Nrf2 molecules are recognized in the cytoplasm, through their NLS1, NLS2, NLS3 by the soluble adaptor proteins termed importins/karyopherins (a and/or b) which upon binding the cargo proteins such as Nrf2, result in a complex that is then ferried through the nuclear pore complex in the nuclear membrane into the nucleoplasm (58–60). These observations collectively suggest that the nucleo-cytoplasm translocation of transcription factors is the consequence of a dynamic equilibrium of multivalent NLS and NES (53–56, 58–60). With regards to the importance of Nrf2 phosphorylation, two Nrf2 phosphorylation events have been recognized to modulate Nrf2 nuclear import/export in response to various chemopreventive agents (51). First, Salazar and Cuadrado (94) have proposed a mechanism involving direct phosphorylation of the transcription factor Nrf2 by glycogen synthase kinase-3b (GSK-3b), downstream of the PI3K/Ser/Thr kinase Akt pathway. It appears that ARE inducers may activate the PI3K/Akt pathway, leading ultimately to Nrf2 nuclear accumulation and localization by downregulating Nrf2 phosphorylation by GSK-3b. It is not yet known whether phosphorylation of Nrf2 by GSK-3b under basal conditions (in the absence of ARE inducers) promotes nuclear export or inhibits nuclear import, or the identity of the site of phosphorylation. A second mechanism whereby Nrf2 phosphorylation regulates Nrf2 localization when cells are exposed to chemopreventive agents and oxidative stress was proposed by Jain and Jaiswal (56). They have described that phosphorylation of tyrosine 568 in Nrf2, by the tyrosine kinase Fyn, is essential for Crm1mediated nuclear export and degradation of the transcription factor. The same authors have also noted that H2O2 initially led to nuclear accumulation of Nrf2, presumably to activate cytoprotective genes, and later, because persistent increase in chemoprotective genes expression threatens cell survival, induced phosphorylation of tyrosine 568 for enhanced nuclear export of Nrf2. However, it was also shown that GSK-3b acts upstream of Fyn kinase in regulation of nuclear export and degradation of Nrf2. In summary, the early response leads to nuclear import of Nrf2 resulting in coordinated activation of chemoprotective genes. Conversely, the delayed response to stress is Fyn-mediated phosphorylation of Nrf2Y568 inside to nucleus. Tyrosine 568 phosphorylation leads to Crm1 exportin-mediated nuclear export of Nrf2. Recently, Sun et al. (70) suggested that Keap1 acts as a key

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postinduction repressor of the Nrf2-mediated antioxidant response by escorting nuclear export of Nrf2. However, Pi et al. (104) have also shown that sequential phosphorylation of the transcription factor Nrf2 by Ca2+-calmodulin (CaM)-dependent protein kinase CK2 plays an important role not only in Nrf2 activation but also in the subsequent Keap1-mediated degradation within the cytoplasm. It is clear that Keap1 and Nrf2 play a pivotal role in the transcriptional activation of cytoprotective genes. In fact, these genes may collectively facilitate the detoxification of carcinogens, enhance the reducing potential against electrophiles and free radicals and increase cellular capacity to repair oxidatively damaged DNA and proteins. However, further work needs to be undertaken to clarify certain functions: (a) which sensing mechanism within cells causes oxidative stress to activate the ARE pathway; (b) how Nrf2 dimerization partners (sMaf, JunD, ATF4, PMF-1), as well as other additional factors (e.g., CREB-binding protein/p300 factors; coactivator RAC3/SRC3; CARM1 and PRMT1) influence the transcriptional response; (c) how the transcriptional repressors Bach1, Bach2, small Maf, Nrf3, ATF3, p53, and retinoic acid receptor alpha (RAR-a) antagonize the Nrf2 function; (d) the number of genes that can be included in the ARE gene battery; (e) how individual cancer chemopreventive agents activate Nrf2; (f) how multiple NLSs and NESs in Nrf2 control the nucleocytoplasmic translocation; (g) how importins and exportins proteins regulate the shuttling in and out of Nrf2 and Keap1; (h) how phosphorylation status of GSK-3b leads to nuclear export/import of Nrf2 resulting in coordinated activation of chemoprotective genes; (i) how Fyn kinase-mediated phosphorylation of tyrosine 568 in Nrf2 (Nrf2Y568) inside the nucleus leads to Crm1exportin-dependent nuclear export of Nrf2; (l) how ubiquitination at lysine-63 or 298 residue of Keap1 and subsequent proteasome-independent degradation of Keap1 regulate Nrf2 ubiquitination downregulation in the transcriptional response; and finally (m) how NES of Keap1 shuttling protein acts in the nucleocytoplasmic translocation. Much also remains to be discovered with regard to the regulation of Nrf2 via phosphorylation by the protein kinase signaling pathways (MAPK, PKC, PERK, or CK2), the cross reactions that exist between these kinases, and the role of Keap1 either as a regulator of Nrf2 degradation by the ubiquitin–proteasome system and postinduction repressor of the Nrf2-mediated antioxidant response by escorting nuclear export of Nrf2. Finally, further studies are also required to determine how the ternary complex containing PGAM5, Keap1, and Nrf2 that is localized to mitochondria regulates nuclear anti-oxidant gene expression in response to changes in mitochondrial functions. Clarification of these issues, along with advances in genomic, proteomics, microarray technology, and informatics, should provide further information as to the importance

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of the Keap1–Nrf2–ARE gene regulatory pathway within cells and, hopefully, to the design of safe therapeutic agents that may prevent the progression of cancer and other inflammatory and neurodegenerative diseases (126).

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adaptors can facilitate cullin-mediated ubiquitylation of proteins by a “tethering” mechanism: a two-site interaction model for the Nrf2-Keap1 complex. J Biol Chem 281(34):24756–24768 Tong KI, Katoh Y, Kusunoki H, Itoh K, Tanaka T, Yamamoto M (2006) Keap1 recruits Neh2 through binding to ETGE and DLG motifs: characterization of the two-site molecular recognition model. Mol Cell Biol 26(8):2887–2900 Eggler AL, Gay KA, Mesecar AD (2008) Molecular mechanisms of natural products in chemoprevention: induction of cytoprotective enzymes by Nrf2. Mol Nutr Food Res 52(Suppl 1):S84–S94 Tong KI, Kobayashi A, Katsuoka F, Yamamoto M (2006) Two-site substrate recognition model for the Keap1-Nrf2 system: a hinge and latch mechanism. Biol Chem 387:1311–1320 Li W, Jain MR, Chen C, Yue X, Hebbar V, Zhou R, Kong AN (2005) Nrf2 Possesses a redox-insensitive nuclear export signal overlapping with the leucine zipper motif. J Biol Chem 280(31):28430–28438 Li W, Yu SW, Kong AN (2006) Nrf2 possesses a redox-sensitive nuclear exporting signal in the Neh5 transactivation domain. J Biol Chem 281(37):27251–27263 Jain AK, Bloom DA, Jaiswal AK (2005) Nuclear import and export signals in control of Nrf2. J Biol Chem 280(32): 29158–29168 Jain AK, Jaiswal AK (2006) Phosphorylation of tyrosine 568 controls nuclear export of Nrf2. J Biol Chem 281(17):12132–12142 Kuge S, Arita M, Murayama A, Maeta K, Izawa S, Inoue Y, Nomoto A (2001) Regulation of the yeast Yap1p nuclear export signal is mediated by redox signal-induced reversible disulfide bond formation. Mol Cell Biol 21(18):6139–6150 Theodore M, Kawai Y, Yang J, Kleshchenko Y, Reddy SP, Villalta F, Arinze IJ (2008) Multiple nuclear localization signals function in the nuclear import of the transcription factor Nrf2. J Biol Chem 283(14):8984–8994 Macara IG (2001) Transport into and out of the nucleus. Microbiol Mol Biol Rev 65(4):570–594 Pemberton LF, Paschal BM (2005) Mechanisms of receptor-mediated nuclear import and nuclear export. Traffic 6(3):187–198 Köhler M, Ansieau S, Prehn S, Leutz A, Haller H, Hartmann E (1997) Cloning of two novel human importin-alpha subunits

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Kotsuji T, Otsuka F, Roop DR, Harada T, Engel JD, Yamamoto M (2003) Keap1-null mutation leads to postnatal lethality due to constitutive Nrf2 activation. Nat Genet 35(3):238–245

Chapter 4 Subnuclear Localization and Intranuclear Trafficking of Transcription Factors Sayyed K. Zaidi, Ricardo F. Medina, Shirwin M. Pockwinse, Rachit Bakshi, Krishna P. Kota, Syed A. Ali, Daniel W. Young, Jeffrey A. Nickerson, Amjad Javed, Martin Montecino, Andre J. van Wijnen, Jane B. Lian, Janet L. Stein, and Gary S. Stein Abstract Nuclear microenvironments are architecturally organized subnuclear sites where the regulatory machinery for gene expression, replication, and repair resides. This compartmentalization is necessary to attain required stoichiometry for organization and assembly of regulatory complexes for combinatorial control. Combined and methodical application of molecular, cellular, biochemical, and in vivo genetic approaches is required to fully understand complexities of biological control. Here we provide methodologies to characterize nuclear organization of regulatory machinery by in situ immunofluorescence microscopy. Key words: Nuclear organization, Runx, Confocal microscopy, Immunofluorescence microscopy, FRAP, Live cell microscopy, Nuclear matrix

1. Introduction The focal distribution of regulatory macromolecules within the nucleus can effectively support the integration of regulatory networks and establish threshold levels of factors for positive and negative control in a broad spectrum of biological contexts that include development and tissue remodeling. Equally important, changes in the composition and organization of regulatory machinery in nuclear microenvironments provides insight into perturbed mechanisms that relate to human disease which is strikingly illustrated by, but not restricted to, skeletal disorders and tumorigenesis (1–5). Examples are modifications in the size,

Paul J. Higgins (ed.), Transcription Factors: Methods and Protocols, Methods in Molecular Biology, vol. 647, DOI 10.1007/978-1-60761-738-9_4, © Springer Science+Business Media, LLC 2010

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number, and composition of intranuclear sites that support transcription, replication, repair, and altered regulatory domains that are causally associated with cleidocranial dysplasia and competency for metastatic breast cancer cells to form osteolytic lesions in bone. Our understanding of the location of regulatory machinery for gene expression, replication, and repair and its role in functional outcome of various biological processes is increasingly evident. However, it is important to define and develop techniques that provide both specific and quantitative insight into various parameters of nuclear architecture. Development and deployment of such approaches is essential for establishing the biological relevance of subnuclear organization as well as necessary for diagnosing disease or providing a platform for development of targeted therapies. Traditionally, compartmentalization of regulatory machinery has been identified and characterized by subnuclear fractionation followed by biochemical and molecular analyses. These are informative approaches, but with limitations. During the past several years, advances in microscopy, together with the development of highly specific antibodies and epitope tags have allowed to examine the assembly and activities of regulatory machinery at single cell level in both the fixed as well as live cell preparations. Thus, the combined use of high-resolution cellular, biochemical, and molecular approaches maximizes the extent to which regulatory mechanisms can be defined. We will focus on visualization of nuclear microenvironment using Runx transcription factors as an example for compartmentalization of regulatory machinery within nuclei of osteoblastic cells. We will present approaches for imaging of focally localized regulatory complexes in interphase nuclei as well as throughout mitosis. Specificity and quantitation of regulatory complexes that are visualized by microscopy are required to informatively relate cell morphology with regulatory mechanisms. In addition, we will describe a recently developed approach in our laboratory, designated “Intranuclear Informatics” that quantitatively assimilates multiple parameters of regulatory protein localization within the nucleus into contributions toward skeletal gene expression from a temporal/spatial perspective (6).

2. Materials 2.1. Preparation of Metaphase Chromosome Spreads from Suspension and Adherent Cell Cultures

1. Karyomax Colcemid (10 µg/ml). 2. 0.075 M Potassium chloride (KCl) solution. Prewarmed to 37°C in a water bath. 3. Fixative: Methanol/Glacial acetic acid at 3:1, made fresh each time. The fixative should be ice-cold prior to use. 4. Microscopy glass slides (prechilled at 4°C).

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1. Sterile Glass coverslips, 22 mm round, coated with 0.5% gelatin. 2. Cytoskeleton (CSK) Buffer: 10× Stock Solution: 1 M NaCl, 100 mM PIPES pH 6.8, 30 mM MgCl2, 10 mM EGTA. 1× Working Solution: Freshly prepare 100 ml of 1× CSK buffer by dissolving 10.27 g sucrose in 77.6 ml of double-distilled water. Add 10 ml of 10× stock CSK buffer, 0.5 ml of Triton X-100, 0.8 ml of ribonucleoside–vanadyl complex (RVC, New England Biolabs, Ipswich, MA), and 0.8 ml of 150 mM AEBSF [4-(2-aminoethyl) benzenesulfonyl fluoride] (Sigma). 3. Digestion Buffer (DB): 10× Stock Solution: 0.5 M NaCl, 100 mM PIPES pH 6.8, 30 mM MgCl2, 10 mM EGTA. Freshly prepare 1× DB as described above for 1× CSK buffer except use 10× DB instead of 10× CSK buffer. 4. Phosphate-buffered saline (PBS): 9.1 mM dibasic sodium phosphate (Na2HPO4), 1.7 mM monobasic sodium phosphate (NaH2PO4) and 150 mM NaCl. Adjust pH to 7.4 with NaOH. 5. Fixatives: 3.7% formaldehyde in PBS (WC fixative), or in 1× CSK buffer (CSK fixative), or in 1× DB (NMIF fixative). All fixatives should be freshly prepared. 6. Stop solution: 250 mM ammonium sulfate in 1× DB. (Add 1 volume of 2 M ammonium sulfate to 7 volumes of 1× DB). 7. Permeabilizing solution: 0.25% Triton X-100 in PBS. 8. RNase-free DNase. 9. PBSA: 0.5% bovine serum albumin (BSA) in PBS. Filter sterilize. 10. Prolong Gold (Invitrogen, Carlsbad, CA).

2.3. Microscopy

1. 40 mm glass coverslips (Bioptechs, Butler, PA). 2. 60 mm Corning culture dishes. 3. Microwave oven. 4. McCoy’s 5A complete media: McCoy’s 5A media with L-glutamine, supplemented with 10% FBS, 1% PenicillinStreptomycin and 1% L-glutamine (200 mM). 5. Live Cell Stage-Closed System Chamber (Bioptechs). 6. Aqueduct slide, gaskets (Bioptechs). 7. Perfusion fluid: McCoy’s 5A complete medium with 20 mM Hepes Buffer Solution (1 M), heated to 37°C. 8. Bioptechs objective heater and slide heater.

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3. Methods 3.1. Preparation of Metaphase Chromosome Spreads from Suspension and Adherent Cell Cultures

Metaphase chromosome spreads are traditionally used for identification of chromosomal abnormalities (translocation, deletions, and insertions) in patients. It has been recently observed that some lineage-specific proteins, such as Runx transcription factors, retain association with chromosomes during mitosis (Fig. 1a). In such cases, metaphase chromosome spreads provide powerful means for the identification of chromosomes where Runx transcription factors reside during mitosis. For example, Runx proteins associate with nucleolar organizing regions (NORs) on metaphase chromosomes. NORs can be visualized in situ by immunolabeling metaphase chromosome spread preparations for Upstream Binding Factor (UBF), a known regulator of ribosomal RNA transcription (Fig. 1b). Below is a protocol used in our laboratory for preparation of metaphase chromosome spreads (7–9).

Fig. 1. Metaphase chromosome spreads. (a) Runx2 is stable throughout mitosis. Synchronously growing Saos-2 cells were fixed and stained for DNA by using DAPI and for Runx2 by using a rabbit polyclonal antibody. Mitotic cells were identified by chromosome morphology. High-resolution images obtained by three-dimensional deconvolution algorithms reveal that Runx2 (green) is localized in mitotic chromosomes. A subset of Runx2 colocalizes with the microtubules, labeled by a-tubulin staining (red). (b) A metaphase chromosome spread of mouse premyoblast C2C12 cells, immunolabeled for Upstream Binding Factor (UBF; red) to identify nucleolar organizing regions (NORs).

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1. Passage 1 × 106 cells in regular growth medium, 1–2 days prior to performing the chromosome spreads. 2. Feed cells with fresh media 12–14 h prior to harvesting and add Colcemid to a final concentration of 0.05 µg/ml; incubate at 37°C for 3–4 h. 3. Transfer the cells into a centrifuge tube and pellet at 750 × g for 5 min. (see Note 1) 4. Aspirate the supernatant completely. 5. Add 10 ml of 0.075 M KCl solution drop by drop, i.e., hypotonic treatment. Resuspend the pellet by pipetting up and down gently. Incubate at 37°C for 30 min. (see Note 2) 6. Add 1 ml of ice-cold fixative (Methanol/Glacial acetic acid 3:1) to the cell suspension and keep at room temperature for 15 min. 7. Spin the cells at 400 × g for 5 min. 8. Aspirate the supernatant completely. Add 2 ml of fresh fixative (methanol/Glacial acetic acid in a 3:1 ratio) to the cells and keep at 4°C for slide preparation. (At this stage the cell suspension can be stored at 4°C for few days or at −20°C long term.)

3.1.2. Adherent Cells

1. Follow the initial steps 1 and 2 as described above in Subheading 3.2.1 for suspension cells. 2. During the mitotic block, some adherent cells become rounded and detach from the plate. In this case, media should not be discarded but should be transferred to a centrifuge tube. 3. Rinse the plate with PBS and detach the cells with 0.5 ml of trypsin. Mix the detached cells with the medium which contains the cells from mitotic shake off (Step 2). 4. Centrifuge the cells at 750 × g for 5 min. Discard the supernatant and resuspend pellet in ice-cold PBS and centrifuge again at 750 × g for 5 min. Repeat the PBS wash. 5. Continue as in step 5 above for suspension cells and follow each step exactly as described.

3.1.3. Slide Preparation

1. Cool slides to 4°C before using them for metaphase spreads. Adjust hot plate for medium heat (ideally 50–60°C). (see Note 3) 2. Take 100–200 µl of the cell suspension and add drop by drop to the slide from a height of about 20 cm. Drain the excess solution by tilting the slide. 3. Immediately put the slide on hot plate (heat shock) for 1 min.

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4. Air dry the slide, check for chromosome spreading in the phase contrast microscope. 5. Keep the slides in a box at 4°C (up to 8 weeks) or at −80°C for a longer period of time. 3.2. Nuclear Matrix Intermediate Filament Preparation

3.2.1. Whole Cell (WC) Preparation

It is becoming increasingly evident that regulatory proteins are organized in highly specialized compartments within the mammalian nucleus. The biological activity of proteins often correlates with their presence or absence in these nuclear microenvironments (1). The subnuclear organization of regulatory proteins can be assessed by the sequential removal of soluble proteins and chromatin from the mammalian cell (Fig. 2) followed by either in situ immunofluorescence or western blot analysis. Below is an optimized protocol that we routinely use for in situ assessment of parameters of gene expression. 1. Plate cells at a density of 0.5 × 106 cells per six-well plate and incubate in humidified incubator with 5% CO2 at 37°C for 24 h. 2. After 24 h, wash cells twice with ice-cold PBS, fix the WC preparation on ice for 10 min (in an experiment, typically two wells of a six-well plate are allocated to each of the WC, CSK, and NMIF preparations) by adding 2 ml of WC fixative per well. 3. Wash cells once with PBS. 4. To facilitate antibody staining of WC preparations, permeabilize WC preparations with 1 ml of permeabilizing solution on ice for 20 min. 5. Aspirate permeabilizing solution and wash twice with PBS. 6. Add 1 ml of PBSA to the wells.

3.2.2. Cytoskeleton (CSK) Preparation

7. Wash cells twice with ice-cold PBS. 8. Add 1 ml of 1× CSK buffer and incubate plates on ice for 5 min while swirling plates once or twice. 9. Wash wells allotted for CSK preparation (see Subheading 3.3.1) once with ice-cold PBS and fix cells by adding 2 ml of CSK fixative per well. 10. Aspirate CSK fixative after 10 min and wash twice with PBS. 11. Add 1 ml of PBSA to the wells.

3.2.3. Nuclear Matrix Intermediate Filament (NMIF) Preparation

12. Wash cells twice with ice-cold PBS. 13. Add 1 ml of 1× CSK buffer and incubate plates on ice for 5 min while swirling plates once or twice.

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Fig. 2. In situ assessment of nuclear microenvironments. Regulatory proteins can be visualized by indirect immunofluorescence in situ. Proteins involved in distinct nuclear processes are localized in specialized nuclear microenvironments. These microenvironments can be further visualized by removing soluble cytosolic and nuclear proteins as well as chromatin. The procedure is schematically outlined. The upper panel shows a cell after sequential extractions that remove soluble cytosolic as well as soluble nuclear proteins. Finally, the chromatin is digested with DNaseI to reveal a network of ribonuclear proteins, designated the nuclear matrix. The middle panel shows corresponding in situ immunofluorescence of an osteoblast costained with tubulin (red) and Runx2 (green). DNA is depicted as a blue colored circle. Each fraction can be also resolved by SDS-PAGE (sodium dodecyl sulfate-polyacrylamide gel electrophoresis) to identify proteins of interest. The bottom panel shows a schematic of western blot analysis of biochemical fractionation for Runx2, which is an architecturally associated protein primarily present in the NMIF fraction.

14. Prepare 1 ml of DB by adding 400 units of RNase-free DNase I to 1× DB. 15. Flatten parafilm on the covers of plates, label and dispense 20 ml drop of DB-containing RNase-free DNase I on the covers of respective plates. (This step is to conserve the amount of DNase I, otherwise add 1 ml of DB-containing RNase-free DNase I to each well.) 16. To digest the chromatin with DNase I, carefully invert the coverslip, so that cells will face DB-containing DNase I. 17. Incubate cells for 50 min at 30°C. Place coverslips back in their respective labeled wells. Add 1 ml stop solution to the wells and incubate plates on ice for 10 min to stop the activity of DNase I.

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18. Wash once with ice-cold PBS and fix NMIF preparations in 2 ml of NMIF fixative on ice for 10 min. 19. Aspirate fixative and wash twice with PBS. 20. Add 1 ml of PBSA. 3.2.4. Immunostaining of the Samples

21. Dilute antibody in PBSA to an appropriate dilution. As quality and specificity of antibodies vary among suppliers and lots, we recommend testing several dilutions to optimize antibody concentration. When immunolabeling cells with two proteins, caution must be practiced to assure that the antibodies used are raised in different species (e.g., mouse versus rabbit). If raised in the same species, they must be of different isotypes (e.g., IgG versus IgM). 22. On parafilm already flattened on the lids of plates, dispense a 20 ml drop of diluted antibody per well. Carefully invert a coverslip on the drop so that the cells are in direct contact with the antibody. Avoid creating air bubbles by gently placing the coverslip on one edge on the antibody droplet and slowly lowering it. Incubate for 1 h at 37°C. 23. Place coverslips back in respective wells on ice with cells facing upward and wash four times with ice-cold PBSA. 24. Stain cells with appropriate secondary antibodies conjugated with fluorochromes (e.g., Texas Red or FITC) for 1 h at 37°C. 25. Place coverslips back in their respective wells and wash four times with ice-cold PBSA. 26. Stain cells with DAPI (0.5 mg/ml DAPI in 0.1% Triton X-100-PBSA) for 5 min on ice. 27. Wash once with 0.1% Triton X-100-PBSA followed by two washes with PBS. Leave cells in last wash on ice. 28. Immediately mount coverslips in an antifade mounting medium (e.g., Prolong Gold) and aspirate excess of mounting medium. After 10–15 min, seal coverslips with nail polish and store at −20°C in dark.

3.3. Microscopy 3.3.1. Fluorescence Microscopy

Fluorescence microscopy provides a powerful tool to assess subcellular and subnuclear localization of regulatory proteins as well as nucleic acids. A variety of microscopes are available; each microscope has its own set of unique features. Below are the instructions specific for the Zeiss Axioplan 2 Microscope. 1. Turn on the mercury lamp, microscope, charged-coupled device (CCD) camera, and computer. Clean all lenses with microscopic lens paper.

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2. Wipe your slide with tissue paper and lens cleaner. Place a small drop of lens oil on your slide. It is important that the coverslips are sealed to avoid any mixing of the mounting medium with lens oil. 3. To find your cells, start at 10× magnification and then proceed to 40× magnification to start your analysis. Once you have identified a cell or field to document, you can then increase the magnification, depending upon your specific requirements. 4. Once you have selected a cell or field for documentation, you are ready to acquire an image using MetaMorph Software (Molecular Devices, Downingtown, PA). 5. Before acquiring an image, make sure that the arrow on the knob, which diverts light either toward 35 mm camera or toward charged-coupled device (commonly called as CCD camera), is pointing toward the CCD camera. 6. In the main menu, go to ACQUIRE to access the Acquire Dialog Box. 7. Enter an exposure time. Set the region of interest by selecting Entire Chip or Central Quadrant option on the Acquire dialog box. 8. Acquire the image on all filters (DAPI, Fitc, Texas Red and Phase) if analyzing a dual labeled slide. 9. The default image depth of the CCD camera is 12 bits. However, these images cannot be opened by Adobe Photoshop or Microsoft PowerPoint Software. Copy images to 8 bit by selecting “Copy to 8 Bits” command on the main task bar. Always keep your original image (i.e., raw data) as it contains the most information. 10. Once acquired, images can be presented (PowerPoint) or published (Adobe Photoshop, Illustrator) directly or can be further analyzed quantitatively by using MetaMorph Imaging Software or the Intranuclear Informatics mathematical algorithm (see below). 3.3.2. Viewing Live Cells Using the Confocal Microscope

Our lab has characterized the Runx family of transcription factors, describing spatial distribution, subnuclear architectural scaffolding and relationships with coregulatory factors. Much of this work was done with fixed cells on an epifluorescence microscope with verification using a confocal microscope. This naturally led to an interest in documenting the Runx protein dynamics using live cell imaging; looking at mobility, mitotic labeling, and protein– protein interactions. The laser scanning confocal microscope coupled with a Bioptechs micro observation system offers us higher image resolution of live cells with the ability to capture images that are

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sharply defined optical sections produced by the elimination of out of focus light and background information from which threedimensional renderings can be recreated. This coupled with the live cell stage allow us to verify the location of Runx foci throughout dynamic processes, for example, mitosis and to assess the mobility of Runx foci in interphase and during mitosis (Fig. 3b) using Fluorescence Recovery After Photobleaching (FRAP) techniques. A common problem occurs while live cell imaging GFPtransfected cells. Cells become extremely sensitive when exposed to blue filtered UV light (green fluorescence) and die while viewing over long periods of time. For example, in a dynamic process like mitosis, cells permanently stall in Metaphase. Using the scanning laser confocal microscope relieves this situation. Red fluorescence proteins (RFP) are not as sensitive to UV light, thus we transfected a RFP mitotic stage marker, found the stage of mitosis we were interested in, then used the lasers to image the spatial localization of GFP Runx labeled protein through mitosis.

Fig. 3. Live cell microscopy by confocal laser scanning. (a) The Bioptics Live Cell Stage-Closed System Chamber allows the viewing of living cells by maintaining 37°C temperature; pH and nutrient supply by perfusing media across a cellladen coverslip. The slide heater (1) warms an Aqueduct Slide (4) to 37°C. The Perfusion Ring Chamber (2) allows media to enter, cross, and exit the chamber, and Gaskets (3, 5) sandwich the Aqueduct Slide and cell-laden coverslip (6) to prevent leaks. This assembly sits in the self-locking base (7) that is placed on the microscope stage. (b) Live image of U2-OS cell in Anaphase. Runx2-EGFP foci (green) localize to mitotic chromosomes in Anaphase. Histone 2B-RFP and Differential Interference Contrast (DIC) images are used to identify mitotic stage. (c) Initially, using high-intensity laser power, a Spot or Region of Interest inside the fluorescent cell is bleached. After bleaching, a series of images are taken at predetermined time intervals to measure the recovery of fluorescence in the bleached spot (Left Panel). Y-axis represents the relative fluorescence after photo bleaching in the bleached spot and X-axis represents time in seconds. Postnormalization, the relative fluorescence in the bleached spot is zero. This time point is represented as zero time. The relative fluorescence increases with time until it reaches asymptote (Right Panel ).

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Another clear advantage to laser scanning confocal microscopy is the elimination of possible bleed through from double labeling by turning off one of the lasers to confirm specific localization. 1. Sterilize 40 mm coverslips set in 60 mm dishes in a microwave for 20 min. 2. Plate cells in McCoy’s 5A complete medium at a density of ~0.3 × 106. Allow cells to grow for 40–48 h to 50–80% confluency. 3. Transfection. Ascertain and document cell growth. Warm complete and incomplete McCoy’s 5A media. Using Roche FuGENE 6 Transfection Reagent, follow the standard protocol for a 60 mm dish using 5 ml total volume of media, 200– 600 ng of DNA and 200 ml total volume of complex per coverslip. 3.3.3. Preparation of Live Cell Stage, a Closed System Chamber

The Focht’s Closed System Chamber (FCS) allows the microscopic observation of living cells by duplicating conditions of an incubator on the microscope. Temperature is controlled by a slide heater and an objective heater. The slide heater works in conjunction with a microaqueduct slide that has a thermally conductive coating which the slide heater arms rest on. The temperature is set by a controller unit. The objective heater’s temperature is also set by a controller unit and has an adjustable loop which surrounds the objective lens. These heaters are designed to eliminate heat-sink loss. Over time, cells under microscopic observation must be fed, pH level maintained and waste eliminated. A micro peristaltic pump working in conjunction with a microaqueduct slide allows medium to perfuse across the coverslip at a precise, very slow rate, feeding the cells, maintaining pH, and eliminating waste. 1. Prewarm the following items to 37°C. One hour prior to using the confocal microscope, warm water bath to 37°C, placing a flask with water and a thermometer in the bath to verify bath temperature. 2. After reaching temperature, place near confocal microscope for perfusion media. 3. Warm perfusion medium, which is McCoy’s 5A medium with 20 mM Hepes-Buffered solution (Hepes maintains the media pH throughout the imaging session), to 37°C. 4. Warm the Focht’s Chamber System (FCS) and parts to 37°C as well as an insulated transporter box with 2 × 50 ml centrifuge tubes of 37°C water (on each side of the chamber) that will keep the chamber warm en route from the warm room to the scope.

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5. One half hour prior to viewing, confirm that the objective lens and slide heater controller systems are calibrated to 37°C, then attach the objective heater to objective lens and turn on to warm. 3.3.4. Microscope Preparation

1. Confirm DIC is aligned, correct condenser (wide field) and rotating condenser prism are in place, and stage height is correct. 2. Assemble chamber with cells in warm room. All chamber pieces, gaskets and tubing have been cleaned with 70% ethanol and distilled water after the last use and again before current use. 3. The order of assembly of the Focht’s Chamber System is as follows (Fig. 3a): Place upper gasket (part 3) on perfusion ring chamber (part 2) matching two holes over two pegs, then align the aqueduct slide (part 4) on the pegs. The aqueduct slide allows the perfusion fluid to flow across the coverslip, keeping the cells fed and warm. Place the lower gasket (part 5) on top of the aqueduct slide. Pipet 0.25 ml of medium onto the aqueduct slide from the 60 mm dish containing the coverslip with cells, filling the channels and mid space area. Finally, invert the coverslip (part 6) with cells onto the layered assembly. It is very important to touch one edge of the coverslip to one side of the aqueduct slide and slowly lower the coverslip onto the aqueduct slide in order to avoid air bubbles. Invert this sandwich assembly and secure in the self-locking base (part 7). Wipe the medium off the underside of the apparatus. 4. Place chamber in the transporter box with the 2 × 50 ml, 37°C warming tubes to keep it warm while walking from the warm room to the microscope room. Place chamber on stage, attach inlet tubing, and turn on dialysis pump to a very slow rate (ex. 0 and 2) and confirm perfusion fluid is entering, crossing, and exiting the chamber. Then attach exit tubing and drain to receiver vessel. Insert slide heater (part 1, Fig. 3a) in the slide warmer receptacle and oil the objective lens.

3.4. Fluorescence Recovery after Photobleaching (FRAP)

A powerful approach for measuring the dynamics of nuclear microenvironments is to track fluorescently tagged molecules in living nuclei. There are always at least two pools of molecules in nuclear microenvironments: free and bound. Molecules will move at diffusion rates when free and at the same rate as the structure when bound. Taking advantage of this difference, photobleachingrecovery techniques can help characterize the binding equilibriums for molecules docking on a simple, stable structure. The analysis becomes complicated if the protein has multiple and heterogeneous interactions. Several studies have examined the

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dynamics of cellular and nuclear GFP-fusion proteins by Fluorescence Recovery after Photobleaching (FRAP) (10–13). In FRAP, fluorescence in a Spot of Interest (SOI) or Region of Interest (ROI) inside the cell is irreversibly bleached with a high-intensity focused laser (Fig. 3c, left panel). This results in nonfluorescent molecules in the bleached areas surrounded by fluorescent molecules outside the bleached region. Since the binding of these molecules is dynamic, bleached molecules will unbind and diffuse away (14, 15). Molecules that are still fluorescent and are bound in the unbleached region will unbind and diffuse into the bleached zone where they can bind. Photobleach recovery rates are determined by unbinding, diffusion, and binding rates (10, 11). After bleaching, a series of images of the bleached cell are taken to measure the recovery of fluorescence in the bleached spot (Fig. 3c, right panel). Most papers in the biological literature report FRAP recovery rates in terms of half time of recovery or T1/2. Some others report “apparent diffusion coefficients” even though FRAP recovery rates are dependent on binding but not on diffusion (11). Therefore, measurement of binding and unbinding constants is very important in understanding FRAP recovery rates, especially for nuclear proteins (10, 11). After FRAP, data from the confocal system is exported into a spreadsheet software package, corrected for the loss of fluorescence in the whole cell and normalized for pre- and immediate postbleach fluorescence in the bleached zone. Postnormalization, full recovery of the fluorescence in the bleached zone might be expected. If there is no full recovery, then a fraction of the protein is immobile, which means the protein is tightly bound to the subcellular structure and exchanges too slowly to be measured in the postbleach session. FRAP has been valuable in many applications including, but not limited to, measurement of the binding of Histone proteins to DNA (11, 16, 17), the binding of the estrogen receptor to promoters (18), the binding of the glucocorticoid receptor to promoters (19), and the dynamic binding of Exon Junction Complex proteins to RNA splicing speckled domains (12, 13). 1. Once the live cell chamber is set up, fluorescent cells are identified and images of them are collected at low laser power. Optimum gain and offset values for images are determined and the settings are saved under a user profile. 2. Initially, 5–10 images are recorded before bleaching a Region of Interest (ROI) or a Spot of Interest (SOI). 3. In spot photo bleaching, one or more than one spot inside the fluorescent cell is selected. Bleaching is usually done with maximum laser power from 1 to 3 s until about 70% of the fluorescence in the spot is bleached. If more than one spot is

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selected for bleaching, the confocal system performs bleaching sequentially. Bleaching can be done in larger regions of the cell, for example, in half of the nucleus, by zooming up to a ROI within the cell and scanning it 30–50 times at full laser intensity, resulting in the photobleaching of this enlarged region of the cell. 4. After photobleaching, routinely 30–50 images are taken at intervals of 1.7–20 s (it can be minutes if desired) depending on the dynamics of the fluorescent protein. (If large area bleaching was performed, images are enlarged to the size equal to the prebleach image.) 5. With the aid of Leica Confocal Software version 2.0, the fluorescence within the bleached spot, the fluorescence in the whole cell or nucleus, and the fluorescence in a region outside the bleach zone are measured for the entire stack of images. 6. For data analysis, fluorescence intensity values from the Leica software are exported to a spreadsheet software package, for example Microsoft Excel. Normalization is done at two levels. At the first level, the initial postbleach intensity is subtracted for the fluorescence in the ROI so that any fluorescence in the bleached area after bleaching is normalized to zero. At the second level, the prebleach fluorescence intensity, corrected for the fluorescence loss in the whole cell that is caused by the bleach, is normalized to 1. 7. The relative fluorescence intensity (I rel) in the bleached spot is measured as described by Phair and Misteli (20): Irel = T0 × If/Tt × I0, with T0 being the total cellular intensity before bleach, Tt the total cellular intensity at time t, I0 the intensity in the bleached area before bleach, and If, the intensity in the previously bleached area at time t. 8. Recovery curves are obtained using Microsoft Excel or Kaleidograph.3.5 (Synergy Development). 9. The immobile protein fraction is calculated by subtracting the relative intensity at the asymptote of the recovery curve from a relative recovery 1. For example, an asymptote at 0.7 reflects an immobile fraction of 30%. 3.5. Intranuclear Informatics

Intranuclear informatics is a mathematical algorithm that is designed to identify and assign unique quantitative signatures that define regulatory protein localization within the nucleus ((6); Fig. 4). Quantitative parameters that can be assessed include nuclear size and variability in domain number, size, spatial randomness, and radial positioning (Fig. 4, top panel). The significance and implication of Intranuclear Informatics can be shown by two distinct biological examples (Fig. 4).

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Fig. 4. Intranuclear informatics. This figure shows how intranuclear informatics can be used to examine nuclear alterations in cancer cells compared with normal cells. The top panel shows the conceptual framework for the quantitation of subnuclear organization by intranuclear informatics. Four main groups of parameters, selected based on inherent biological variability, can be examined. Example 1. Regulatory proteins with different activities can be subjected to Intranuclear Informatics analysis, which assigns each protein a unique architectural signature. The overlap between the architectural signatures of different proteins is often correlated to their functional overlap. Shown here are Runx transcription factor, SC35 splicing protein, and RNA Pol II. Example 2. The subnuclear organization of Runx domains is linked with subnuclear targeting, biological function, and disease. Biologically active Runx2 and inactive subnuclear targeting defective mutant (mSTD) show distinct architectural signatures, indicating that the biological activity of a protein can be defined and quantified as subnuclear organization.

Regulatory proteins with different activities can be subjected to Intranuclear Informatics analysis that assigns each protein a unique architectural signature. The overlap between the architectural signatures of different proteins is often correlated to their functional overlap. Alternatively, the subnuclear organization of a protein domain can be linked with subnuclear targeting, biological function, and disease. For example, biologically active Runx2 and its inactive subnuclear targeting defective mutant (mSTD) show

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distinct architectural signatures, indicating that the biological activity of a protein can be defined and quantified as subnuclear organization. These architectural signatures have the potential to discriminate between intranuclear localization of proteins in normal and cancer cells. Intranuclear informatics can be combined with proteomics (changes in protein–DNA and protein–protein interactions) and genomics (altered gene expression profiles) to develop a novel platform for identification and targeting of perturbed regulatory pathways in cancer cells.

4. Notes 1. If the metaphase chromosomes are highly condensed, use a lower concentration of Colcemid and decreased time of Colcemid treatment. 2. Appropriate hypotonic treatment is vital to the quality of metaphase spreads. The concentration of KCl can be changed according to the quality of chromosome spread. 3. Drop the cell suspension on cold, slightly moist slides. Chromosomes will spread poorly on dry slides. If necessary, breathe on the slide before dropping the suspension.

Acknowledgments Studies reported in this article were in part supported by grants from NIH (5PO1CA82834-05, 5PO1AR048818-05, 2R01GM32010, 5R01AR049069). Core resources supported by the Diabetes Endocrinology Research Center grant DK32520 were also used. References 1. Stein GS, Zaidi SK, Braastad CD, Montecino M, van Wijnen AJ, Choi J-Y et al (2003) Functional architecture of the nucleus: organizing the regulatory machinery for gene expression, replication and repair. Trends Cell Biol 13:584–592 2. Zaidi SK, Young DW, Choi JY, Pratap J, Javed A, Montecino M et al (2004) Intranuclear trafficking: organization and assembly of regulatory machinery for combinatorial biological control. J Biol Chem 279:43363–43366

3. Zaidi SK, Young DW, Choi JY, Pratap J, Javed A, Montecino M et al (2005) The dynamic organization of gene-regulatory machinery in nuclear microenvironments. EMBO Rep 6:128–133 4. Zink D, Fischer AH, Nickerson JA (2004) Nuclear structure in cancer cells. Nat Rev Cancer 4:677–687 5. Taatjes DJ, Marr MT, Tjian R (2004) Regulatory diversity among metazoan coactivator complexes. Nat Rev Mol Cell Biol 5:403–410

Subnuclear Localization and Intranuclear Trafficking of Transcription Factors 6. Young DW, Zaidi SK, Furcinitti PS, Javed A, van Wijnen AJ, Stein JL et al (2004) Quantitative signature for architectural organization of regulatory factors using intranuclear informatics. J Cell Sci 117:4889–4896 7. Henegariu O, Heerema NA, Lowe WL, BrayWard P, Ward DC, Vance GH (2001) Improvements in cytogenetic slide preparation: controlled chromosome spreading, chemical aging and gradual denaturing. Cytometry 43:101–109 8. Claussen U, Michel S, Muhlig P, Westermann M, Grummt UW, Kromeyer-Hauschild K et al (2002) Demystifying chromosome preparation and the implications for the concept of chromosome condensation during mitosis. Cytogenet Genome Res 98:136–146 9. Deng W, Tsao SW, Lucas JN, Leung CS, Cheung AL (2003) A new method for improving metaphase chromosome spreading. Cytometry A 51:46–51 10. Lele T, Oh P, Nickerson JA, Ingber DE (2004) An improved mathematical approach for determination of molecular kinetics in living cells with FRAP. Mech Chem Biosyst 1:181–190 11. Lele T, Wagner SR, Nickerson JA, Ingber DE (2006) Methods for measuring rates of protein binding to insoluble scaffolds in living cells: histone H1-chromatin interactions. J Cell Biochem 99:1334–1342 12. Wagner S, Chiosea S, Ivshina M, Nickerson JA (2004) In vitro FRAP reveals the ATPdependent nuclear mobilization of the exon junction complex protein SRm160. J Cell Biol 164:843–850

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13. Wagner S, Chiosea S, Nickerson JA (2003) The spatial targeting and nuclear matrix binding domains of SRm160. Proc Natl Acad Sci U S A 100:3269–3274 14. Berezney R, Basler J, Kaplan SC, Hughes BB (1979) The nuclear matrix of slowly and rapidly proliferating liver cells. Eur J Cell Biol 20:139–142 15. Kruhlak MJ, Lever MA, Fischle W, Verdin E, Bazett-Jones DP, Hendzel MJ (2000) Reduced mobility of the alternate splicing factor (ASF) through the nucleoplasm and steady state speckle compartments. J Cell Biol 150:41–51 16. Lever MA, Th’ng JP, Sun X, Hendzel MJ (2000) Rapid exchange of histone H1.1 on chromatin in living human cells. Nature 408:873–876 17. Misteli T, Gunjan A, Hock R, Bustin M, Brown DT (2000) Dynamic binding of histone H1 to chromatin in living cells. Nature 408:877–881 18. Stenoien DL, Patel K, Mancini MG, Dutertre M, Smith CL, O’Malley BW et al (2001) FRAP reveals that mobility of oestrogen receptor-a is ligand- and proteasome-dependent. Nat Cell Biol 3:15–23 19. Stavreva DA, Muller WG, Hager GL, Smith CL, McNally JG (2004) Rapid glucocorticoid receptor exchange at a promoter is coupled to transcription and regulated by chaperones and proteasomes. Mol Cell Biol 24:2682–2697 20. Phair RD, Misteli T (2000) High mobility of proteins in the mammalian cell nucleus. Nature 404:604–609

Chapter 5 Analysis of Ligand-Dependent Nuclear Accumulation of Smads in TGF-b Signaling Douglas A. Chapnick and Xuedong Liu Abstract The growth inhibition of dividing cells and most of the transcriptional responses upon TGF-b treatment depend on the Smad2, Smad3, and Smad4 transcription factors. These proteins shuttle continuously between the cytoplasm and the nucleus, transmitting the ligand status of the TGF-b receptors to the nuclear transcription machinery. In the absence of TGF-b ligand, Smads 2/3/4 reside predominantly in the cytoplasm. Following ligand binding to the TGF-b receptors, the dynamic equilibrium of shuttling Smads 2/3/4 shifts toward a predominantly nuclear state, where a high concentration of these transcription factors drives transcriptional activation and repression of genes required for proper cellular response. Here, we describe live cell imaging and immunofluorescence microscopy methods for tracking Smads subcellular localization in response to TGF-b and leptomycin B treatment. In addition, a method of fractionating nuclear and cytoplasmic proteins used to confirm the imaging results was presented. Our results support the notion that the R-Smad shuttling mechanism is distinct from Co-Smad. Key words: TGF-b, Smad4, Smad2, Nuclear accumulation, Cellular fractionation, Immunofluorescence, Leptomycin B

1. Introduction Transforming Growth Factor Beta (TGF-b) is a cytokine that can cause several distinct cellular responses, including growth inhibition, apoptosis, and differentiation (for review (1)). The active form of the TGF-b ligand binds as a dimer to the TGF-b receptors on the plasma membrane of target cells, thus carrying a signal from one cell to another (For Review on ligand–receptor interactions (2)). Type I and Type II TGF-b receptors (TBRI and TBRII, respectively) are serine/threonine kinases that transmit the signal across the plasma membrane (1). TGF-b binding to TBRII induces hetero-oligomerization of TBRII with TBRI (1). Paul J. Higgins (ed.), Transcription Factors: Methods and Protocols, Methods in Molecular Biology, vol. 647, DOI 10.1007/978-1-60761-738-9_5, © Springer Science+Business Media, LLC 2010

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The receptor oligomerization initiates a protein phosphorylation cascade, eventually propagating the signal into the nucleus. This cascade begins when TBRII phosphorylates TBRI, resulting in activation of TBRI’s kinase activity (1). Active TBRI then phosphorylates receptor regulated Smad proteins also known as R-Smads (Smad2 and Smad3) (3). These phosphorylated Smad2/3 proteins homo-oligomerize into protein complexes, as well as hetero-oligomerize with Smad4 or Co-Smad (4, 5). Oligomerization of Smads 2/3/4 correlates with nuclear accumulation of these proteins (6). Smads 2/3/4 are transcription factors that activate the transcription of p21 and p15, which cause cell cycle arrest in the G1 phase of the cell cycle, as well as repress transcription of growth promoting genes such as c-myc and CDC25A (1). It is through transcriptional regulation of these genes, and possibly others yet to be identified, that Smad nuclear accumulation leads to cytostatic responses. Nucleocytoplasmic shuttling of signaling proteins is a common theme shared by many cellular signaling pathways. Smads are constantly cycling in and out of nucleus even in the absence of ligand stimulation (7, 8). The Smad nuclear export rates exceed their import rates in the basal state; consequently, both Smad2/ Smad3 and Smad4 are predominantly localized to the cytosol (9, 10). Ligand stimulation decreases the export rates of Smad4 without significantly affecting the import rates resulting in nuclear accumulation of Smad4 (10). Similar mechanism may also account for R-Smad nuclear accumulation although nuclear import and export mechanism of R-Smad and Smad4 appears to be distinct. Earlier studies have shown that Smad3 but not Smad2 or Smad4 can directly interact with importin b1 and interaction may be important for nuclear translocation of Smad3 (11–13). Subsequent studies suggest that nuclear import of Smad2 and Smad3 can also occur through direct binding of Smad2/3 to nucleoporins Nup214 and Nup153 (8). Thus, both importin b-dependent and importin b-independent pathways are involved in trafficking R-Smad into nucleus. For shuttling R-Smad out of the nucleus, exportin 4 has been implicated as the export factor for Smad3 and most likely for Smad2 as well (14). Unlike R-Smad, nuclear import of Smad4 likely relies on importin 7/8 or importin alpha (13, 15) while Smad4 nuclear export occurs by binding to the nuclear export factor CRM1 as treatment of the specific small molecule inhibitor Leptomycin B (LMB), which targets CRM1, is sufficient to drive nuclear accumulation of Smad4 but not Smad2 or Smad3 (16–18). Two nuclear export signals have been identified in Smad4 and mutation of these signals causes Smad4 to exclusively localize to nucleus (12, 16, 17). Despite all these advances, a number of outstanding questions still remain unanswered. For example, how is the phosphorylated R-Smad induced by TGF-b treatment translocated to the nucleus? Is the

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import rate for the phosphorylated Smad2/3 higher than the unphosphorylated Smad2/3? How Smad homo- or heterooligomerization regulates Smad nuclear accumulation? Can the rate of Smad import or export be readjusted intrinsically or in response to signaling cross-talk? Here we described some of the key methods that can be used to determine the trafficking mechanisms of Smad in the mammalian system.

2. Materials 2.1. Cell Culture

1. Dulbecco’s Modified Eagle Medium (DMEM) (GIBCO, Invitrogen). 2. DMEM lacking phenol red (GIBCO, Invitrogen). 3. GlutaMAX L-Glutamine Supplement (GIBCO, Invitrogen). 4. Dulbecco’s Phosphate Buffered Saline (D-PBS) (GIBCO, Invitrogen). 5. 100 X Penicillin G Solution (Solid Penicillin G from Sigma in distilled water to 10,000 U/ml). 6. Streptomycin Sulfate solution (Solid streptomycin sulfate from Sigma in distilled water to 10,000 U/ml).

2.2. Live Cell Treatment

1. Leptomycin B, 500 µg in absolute ethanol (LC Laboratories).

2.3. Cellular Fractionation

1. Hypotonic Lysis Buffer: 10 mM Tris Base HCl, 10 mM KCl, 1.5 mM MgCl2, 1 mM sodium orthovanadate, 0.2 mM phenylmethanesulphonylfluoride, 1 mM DTT.

2. Transforming growth factor-b1 (TGF-b1) (R and D Systems).

2. RIPA buffer: 150 mM NaCl, 1% v/v Triton X 100, 1% w/v sodium deoxycholic acid, 0.1% w/v sodium dodecyl sulfate, 25 mM Tris–HCl, 1 mM ethylenediaminetetraacetic acid, 0.2 mM phenylmethanesulphonylfluoride, 1 mM sodium orthovanadate, 1 mM DTT, 25 mM b-glycerophosphate, 25 mM NaF. 3. Salt Balancing Solution: 10% v/v Triton X 100, 1 M NaCl, 100 mM b-glycerophosphate, 100 mM NaF. 4. Cell Lifter (Costar, Corning 3008). 5. Dounce Homogenizer with 1 ml volume (Wheaton). 6. Ponceau S Staining Solution: 0.5 g Ponceau S Dye, 5% glacial acetic acid to 100 ml with deionized water. 7. Protogel 30% acrylamide/bisacrylamide solution (National Diagnostics). 8. Protogel 4× Resolving Gel Buffer (National Diagnostics).

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9. Protogel Stacking Gel Buffer (National Diagnostics). 10. 10% sodium dodecyl sulfate solution: 10% w/v SDS in distilled water. 11. Mouse anti-Smad1/2/3 antibody (Santa Cruz Biotechnology). 12. Mouse anti-Lamin A/C antibody (Santa Cruz Biotechnology). 13. Mouse anti-betaActin antibody (AbCam). 14. Horse radish peroxidase-conjugated sheep anti-mouse antibody (GE Healthcare). 15. SuperSignal West Dura Extended Duration Substrate chemiluminesence kit (Pierce Biotechnology). 16. Protran nitrocellulose membrane (Whatman). 17. Whatman chromatography paper (Whatman). 18. Semi-dry transfer apparatus (Hoefer TE70). 19. BCA protein assay kit (Thermo 23225). 20. Spectra broad ranged multicolor protein ladder (Fermentas SM184). 21. Powerwave X Scanning Spectrophotometer Plate Reader (Bio-Tek). 22. 4× SDS-Gel loading buffer (to Make 10 ml): 8 mg Bromophenol Blue, 1 ml 0.5 M EDTA, 40 mM DTT, 4 ml 100% glycerol, 0.8 g SDS, 2 mL 1 M Tris–HCl pH 6.8, to 10 ml with deionized water. 23. Transfer buffer: 5.8 g Glycine, 11.6 g Tris–HCl, 0.72 g sodium dodecyl sulfate, 400 ml methanol, to 2 L with deionized water. 24. Tris-buffered saline supplemented with Tween-20 detergent (TBS-t): 8.8 g NaCl, 0.2 g KCl, 3 g Tris–HCl, 1 ml Tween 20, pH 7.4, to 1 L with deionized water. 25. Western blot film (ISC BioExpress). 2.4. Immunofluorescence

1. Poly-D-lysine hydrobromide solution: 1 mg/ml poly-D-lysine hydrobromide (Sigma), 23.5 mM sodium tetraborate, 50 mM boric acid, pH 8.5). 2. Round glass coverslips (Fisherbrand). 3. Glass slides (VWR). 4. Rabbit anti-Smad2 antibody (Zymed Laboratories). 5. Mouse anti-Smad4 antibody (Santa Cruz Biotechnology). 6. Goat anti-mouse Alexafluor488-conjugated (Molecular Probes, Invitrogen).

antibody

7. Goat anti-rabbit AlexaFluor555-conjugated (Molecular Probes, Invitrogen).

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8. Normal goat serum (Invitrogen). 9. 3.7% paraformaldehyde solution: dilution of 16% solution in deionized water (Electron Microscopy Sciences). 10. 10 mg/ml Hoescht 33258 (Invitrogen) solution in deionized water. 11. Clear nail polish (Sally Hansen “Hard as Nails”). 12. Nikon ECLIPSE TE2000 inverted fluorescence microscope equipped with the following; excitation filters: 360/40 Hoescht 55258, 490/20 AlexaFluor488, 555/28 AlexaFluor555, 470/30 GFP, 492/18 YFP. Emission Filters: 457/50 Hoescht 33258, 528/38 AlexaFluor488, 617/73 AlexaFluor555, 510/30 GFP, 535/30 YFP. Camera: COOLSNAP ES. Software: Metamorph Premier Imaging System. 2.5. Live Cell Imaging

1. GFP-Smad4 HaCaT cells were created by retroviral-mediated gene transfer. Briefly, pMX-GFP-Smad4, a retroviral expression vector described previously (18), was transfected into the amphotrophic packaging cell line jNX 293 T cells using Mirus (Mirus Bio, Madison, WI). Infection and selection of GFP-positive stable cell lines using FACS sorting were performed as described (19). Similar procedure was used to create YFP-Smad2 HaCaT Cells except the expression vector used was pREX-YFP-Smad2-IRES-Hygromycin. 2. Glass bottom 35 mm petridishes (Mat Tek Corporation).

3. Methods 3.1. Cellular Fractionation of Cytoplasm and Nuclear Proteins

1. Using the methods of cellular fractionation and western blot analysis, we find that both TGF-b and Leptomycin B treatment of cells is sufficient to increase the fraction of total cellular Smad4 protein located in the nucleus. This is not observed to be the case for either Smad2 or Smad3, where only TGF-b treatment was sufficient to increase the fraction of total cellular Smad2/3 protein in the nucleus (Fig. 1). 2. 4 × 10 cm tissue culture Petri dishes were seeded with 10 ml and 2 × 106 adherent HaCaT cells and allowed to grow for 24 h at 37°C in a 5% CO2 atmosphere (see Note 1) Two plates were treated for 30 min with 20 ng/ml LMB prior to addition of 100 pM TGF-b1 to one of these plates and to one of two plates not treated with LMB. Cells were treated with TGF-b1 for 1 h. Total volume of media in all plates is 10 ml.

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Fig. 1. Determining the effect of LMB on the relative amounts of Smad2/3/4 in the cytoplasm and the nucleus. HaCaT cells were treated with and without 100 pM TGF-b1 for 1 h. This experiment was repeated with a treatment of 20 ng/ml LMB, where LMB was applied 30 min prior to addition of TGF-b. Cells were fractionated into cytoplasm and nuclear fractions and western blotted to determine the relative amounts of Smad2/3/4. Lamin A/C and b-actin were used as both loading controls and indicators of the purity of fractions.

3. Each 10 cm plate containing 2 × 106 adherent HaCaT cells are rinsed (see Note 2) one time with 10 ml 4°C Dulbecco’s Phosphate Buffered Saline (D-PBS). D-PBS is removed by tilting the plate vertically at an 80° angle in a bucket full of ice for 1 min and aspirating all liquid from the bottom corner of the plate using a vacuum trap and a glass Pasteur pipette. 10 ml of 4°C hypotonic lysis buffer is then added to the plate, which is then incubated horizontally on ice for 15 min. During this incubation, cells will swell, providing an easier lysis in the following steps (see Note 3). All hypotonic lysis buffer is removed in the same manner as stated above. 70 µl of 4°C hypotonic lysis buffer supplemented with 0.4% TX-100 is added to each plate. With the plate tilted at an 80 degree angle in a bucket of ice, cells are scraped using a cell lifter, and all cells are pushed toward the pooling liquid in the bottom end of the tilted plate. 4. Scraped cells in lysis buffer are briefly homogenized by pipetting up and down the pooled liquid in the plate, being careful not to introduce air bubbles (see Note 4). The mixture of buffer and cells is then transferred to a 1.7 ml Eppendorf tube on ice, and incubated at 4°C for 15 min. 5. Using a 200-µl pipette, cells are further homogenized by pipetting up and down without introducing air bubbles. Cells and buffer are transferred to a clean 4°C 1 ml Dounce homogenizer, and the “loose fit” plunger is raised and lowered 60 times. The plunger is removed and the liquid is allowed to collect in the bottom of the homogenizer for 30 s.

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1–2 µl of liquid homogenate is removed, and placed on a microscope slide and a cover-slip is applied to prevent evaporation. The homogenate is confirmed to be composed of nuclei, which look wrinkled oval and dark, and debris, which is the result of plasma membrane breakage. There should be no presence of in-tact cells at this point. However, if there are still 5–10% unbroken cells one may apply the plunger again and homogenize the cells 40–60 more times before proceeding to step 5 (see Notes 5 and 6). 6. Homogenate is transferred to a fresh 4°C 1.7 ml Eppendorf tube and spun for 5 min at 800 × g and 4°C. A small pellet forms at the end of the tube, and the supernatant contains plasma membrane components and cytoplasm contents. Most of the supernatant is removed by pipetting and labeled as cytoplasmic fraction, and care is taken not to disturb the pellet (see Note 7). Cytoplasmic fractions are balanced for salt, detergent, and phosphatase inhibitors by addition of 20 µl salt balancing buffer. 7. The last bit of supernatant is removed and discarded. The nuclear pellet is rinsed one time by addition of 100 µl of 4°C hypotonic buffer lacking 0.4% TX-100. The tube containing hypotonic wash buffer and nuclei is then spun for 5 min at 800 × g and 4°C (see Note 8). All liquid contents of the tube are removed by aspiration, being careful not to disturb the pellet or to scrape the inner walls of the tube. The nuclei are then lysed completely by addition of 70 µl of RIPA buffer followed by gentle flicking and inverting of the tube. These tubes are then labeled as nuclear fractions. 8. Cytoplasmic and nuclear fractions are rotated for 45 min at 4°C. 9. Fractions are then spun for 10 min at 13,200 × g and 4°C. Supernatants are transferred to new labeled tubes and stored on ice (see Note 9). 3.2. Determining Protein Concentration and Performing Western Blot Analysis

1. Protein concentration is determined using a BCA assay kit, according to the manufacturer’s instructions. Briefly, serial dilutions of 2.0 mg/ml BSA stock solution in a 1:10 dilution of RIPA solution: distilled water are made to produce 1.0 mg/ml, 0.5 mg/ml, 0.25 mg/ml standards, while a blank is made from 1:10 dilution of RIPA solution:distilled water alone. A 2 µl aliquot of each fraction is removed to make a 1:10 dilution of each unknown sample in distilled water, yielding a total of 20 µl of diluted unknown sample. 2. 5 µl of each unknown and each standard are mixed with 100 µl of 1× BCA solution (50:1 mixture of solutions A and B, from the BCA kit), each in 1 well of a clear 96-well

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polyethylene plate. Plates are completely sealed to prevent evaporation using parafilm tape, and incubated at 37°C for 30 min. 3. The 96-well plate is read at 562 nm for absorbance using a 96-well plate scanning spectrophotometer. Using Excel, a spreadsheet is constructed to determine the mathematical relationship of BSA concentration to absorbance for the standards, and this relationship is used to determine the concentration of the proteins in each unknown sample. The total yield in the cytoplasmic fraction is 150 µg in 100 µl, while the total yield in the nuclear fraction is 100 µg in 75 µl. 4. For each cytoplasmic fraction, 50 µg of total protein is prepared for loading into a single well, while 33.3 µg of total protein is prepared for loading into a single well for nuclear fractions (see Note 10). Each sample for loading into an SDS-PAGE gel is mixed with 7 µl of 4× SDS-loading buffer and incubated at 95°C for 5 min. Tubes are inverted and liquid contents are briefly spun at 5,000 × g for 30 s. In this experiment, two identical sample sets are used to make two identical gels. 5. A 1.5 mm, 12% polyacrylamide mini-gel polymerized with a 10-well comb (manufactured using SDS-PAGE Protogel reagents from National Diagnostics according to the manufacturer’s instruction) is loaded with samples and 5 µl Spectra protein ladder (Fermentas) and 10 µl of SDS-loading buffer is added to any spare/empty wells prior to application of current. Each gel is run for 1 h at 35 mA (190 V), at which time the bromophenol blue dye in the SDS-loading buffer runs just out of the bottom of the gel. 6. Each gel is transferred in a semi-dry western blot horizontal transfer unit, using a sandwich from cathode (bottom piece) to anode (top piece) with the following scheme: three pieces of chromatography paper, one piece of nitrocellulose paper, SDS-PAGE gel containing samples, three pieces of chromatography paper. This sandwich is assembled under 50 ml of transfer buffer, and removed as a sandwich and placed in the transfer apparatus such that the gel is above the membrane, as proteins will be deposited on the nitrocellulose membrane as they move down toward the cathode. The sandwich is then lightly ironed with a 10 ml glass test tube to ensure no air bubbles are trapped between the membrane and the gel. For each sandwich containing one gel, for 1.5 h at 45 mA or 7 V is applied to the apparatus, which is assembled as indicated by the manufacturer. 7. Nitrocellulose is removed from the apparatus following transfer of proteins, and stained with 10 ml of Ponceau S Staining solution at room temperature for 1 min. Staining solution is removed and the membrane is rinsed five times with 20 ml of

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distilled water to remove nonspecific Ponceau S stain. Each cytoplasmic lane should have even and bright red staining. In contrast, far less staining should be present for each nuclear sample. At this point, one can make a reasonable assessment of the purity of the nuclear fractions by seeing whether abundant protein bands in the cytoplasmic fraction lanes are shared in the nuclear fraction lanes. 8. Each membrane is blocked with 10 ml of 3% (w/v) non-fat dry milk in TBS-t at room temperature for 45 min. 9. Blocking buffer is completely removed and discarded prior to the addition of 5 ml of 1:500 mouse anti-Smad1/2/3 in 3% (w/v) non-fat dry milk in TBS-t to one membrane, while another solution of 1:1,000 mouse anti-Smad4 in 3% (w/v) non-fat dry milk in TBS-t is added to the other membrane. 10. Each blot is incubated with primary antibody solution for 3 h at room temperature, on a table top rocker. 11. Membranes are washed two times for 2 min with 10 ml of TBS-t. 12. Each membrane is then incubated with 3 ml of 1:2,000 antimouse HRP-conjugated secondary antibody solution in 3% (w/v) non-fat dry milk in TBS-t for 50 min at room temperature on a table top rocker. 13. Each membrane is rinsed with 10 ml of TBS-t, and subsequently washed three times with 15 ml of TBS-t for 8 min each wash. 14. Membranes are removed from wash buffer and allowed to drip for 10 s before being laid protein side up on a piece of clear plastic(SARAN wrap is sufficient, or a cut three-ringed binder sheet protector). To each membrane is added 200 µl of West Dura solution (a mixture of 100 µl solution A and 100 µl solution B) and lightly tilted in several directions, by hand, to ensure that this 200 µl of solution covers the entire membrane. Another clear plastic sheet is laid over the protein side up membrane, creating a sandwich of two plastic pieces around the membrane, which is allowed to sit for 30 s prior to ironing out excess liquid from inside the sandwich with a paper towel. 15. Plastic/membrane sandwiches are then taped to the inside of an imaging cassette, and exposed with X-ray developing film in a dark room for 15 s, 30 s, 1 min, and 5 min to produce varied exposures of the protein bands. Film is then developed using an automatic film developer. 16. This process from steps 9 to 15 is repeated with a 1:1,000 dilution of mouse anti-Lamin A/C for one membrane, and a 1:1,000 dilution of mouse anti-b-actin for the other membrane. Both antibody dilutions are made in 5 ml of 3% (w/v) nonfat dry milk in TBS-t.

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3.3. Immunofluorescence Detection of Smad2 and Smad4

1. Using immunofluorescent image analysis, we find that the ratio of cytoplasmic Smad4 to nuclear Smad4 (C:N) is 1.22 ± 0.15 in the basal state, 0.57 ± 0.08 when treated with TGF-b, 0.53 ± 0.16 when treated with LMB, and 0.46 ± 0.08 when treated with both TGF-b and LMB. Thus, either LMB or TGF-b treatment is sufficient to drive the Smad4 from mostly cytoplasmic (C:N>1) to mostly nuclear (C:N90% of the cytoplasm was bleached using 100% laser power, and fluorescence recovery was monitored over a period of up to 350 s. (b) The mean recovery curve for b-catenin export was plotted against time. The fluorescence intensity was calculated as the cytoplasmic to nuclear (C:N) ratio which was preset to 100% based on prebleach values. The recovery was measured at 0.5 s for the first 32 s and then at longer time intervals. Cd-tomato (~60 kDa) protein was used as a negative control. (c) The initial export rate is plotted in the bar graph. The slope was calculated using linear regression analysis as outlined in Subheading 3.5.

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stage – 30/40 frames at 10 s interval. The third recovery stage needs to be optimized for each construct and assay. (Remember to append all images after acquisition.) Acquire fluorescence intensities for the cytoplasm (bleached region), nucleus (nonbleached part of the cell), and background (where there is no cell in view) using Olympus Fluoview software and export to a Microsoft Excel file. Repeat the experiment twice with at least ten cells (see Fig. 3a).

4. Data Analysis 1. Background subtraction: All images contain background signal. Subtract background values from cytoplasmic and nuclear values. 2. Normalization: To compare the rates of nuclear export between different samples, express the fluorescent data as a cytoplasmic/nuclear ratio. For each cell data set, set the prebleach ratio to 100%. Set the time for first prebleach image as 0 s. Set the recovery curve to 25% recovery at time zero (this was the closest value). Average the data for at least ten cells. Plot the recovery curve versus time (Fig. 3b). 3. Initial export rate: import the average data for the first 32 s from Microsoft Excel into Graph Pad Prism 5.0 (see Note 12). Analyse the curve using linear regression and obtain the best slope fit value. This is the initial export rate. Compare this value for each construct (Fig. 3c).

5. Notes 1. Prepare in a glass container. Stir on a warm plate for complete mixing. 2. LipofectAMINE is a lipid-based agent which associates with DNA through a charge interaction. The formation of the lipid– DNA complex promotes cellular internalization of the DNA. Mixing with the vortex can reduce transfection efficiency. 3. Changing the media after 5 h reduces the toxicity to cells. However, some cell lines are more resistant and may not require media change. 4. Staining using two primary antibodies: Two primary antibodies can be mixed together if they are from different origin (e.g., mouse monoclonal b-catenin antibody can be mixed with rabbit IQGAP1 polyclonal antibody in step 5, Section 3.2). Make sure to add secondary and tertiary antibodies accordingly.

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5. Dry the coverslip thoroughly using the aspirator and establish a good meniscus before applying the antibody evenly onto the coverslip. 6. Coverslips can be coated with poly-L-lysine prior to seeding so that cells remain attached during the protocol. 7. The concentration needs to be optimized for each preparation of digitonin, and the digitonin should be made fresh within 2 weeks and stored at 4°C. 8. Cells are very fragile at this stage. Make sure to use the aspirator at a minimum level. 9. Two- or four-well chamber cover glass (described in Subheading 2.4) are ideal for live cell imaging as the cover glass is 0.17 mm thick with minimal autofluorescence. 10. Nuclear import assay can be done using similar protocol except choosing the nuclear ROI for bleaching and expressing data as nuclear/cytoplasmic ratio during normalization. 11. Avoid using autofocus parameter on the microscope as it will take a longer time to image at every time point. 12. We compared the actual rate of export during the first 30 s, when transport is least influenced by retention and reequilibration and was measured most accurately at 0.5 s intervals. References 1. Lustig B, Behrens J (2003) The Wnt signaling pathway and its role in tumor development. J Cancer Res Clin Oncol 129:199–221 2. Polakis P (2007) The many ways of Wnt in cancer. Curr Opin Genet Dev 17:45–51 3. Gumbiner BM (2000) Regulation of cadherin adhesive activity. J Cell Biol 148:399–404 4. Nelson WJ (2008) Regulation of cell-cell adhesion by the cadherin-catenin complex. Biochem Soc Trans 36:149–155 5. Brabletz T, Jung A, Reu S et al (2001) Variable beta-catenin expression in colorectal cancers indicates tumor progression driven by the tumor environment. Proc Natl Acad Sci U S A 98:10356–10361 6. Thiery JP (2002) Epithelial-mesenchymal transitions in tumour progression. Nat Rev Cancer 2:442–454 7. Nelson WJ, Nusse R (2004) Convergence of Wnt, beta-catenin, and cadherin pathways. Science 303:1483–1487 8. Gavert N, Ben-Ze’ev A (2007) Beta-catenin signaling in biological control and cancer. J Cell Biochem 102:820–828 9. Henderson BR, Fagotto F (2002) The ins and outs of APC and beta-catenin nuclear transport. EMBO Rep 3:834–839

10. Henderson BR (2000) Nuclear-cytoplasmic shuttling of APC regulates beta-catenin subcellular localization and turnover. Nat Cell Biol 2:653–660 11. Eleftheriou A, Yoshida M, Henderson BR (2001) Nuclear export of human beta-catenin can occur independent of CRM1 and the adenomatous polyposis coli tumor suppressor. J Biol Chem 276:25883–25888 12. Wiechens N, Fagotto F (2001) CRM1- and Ran-independent nuclear export of betacatenin. Curr Biol 11:18–27 13. Koike M, Kose S, Furuta M et al (2004) betaCatenin shows an overlapping sequence requirement but distinct molecular interactions for its bidirectional passage through nuclear pores. J Biol Chem 279:34038–34047 14. Henderson BR, Eleftheriou A (2000) A comparison of the activity, sequence specificity, and CRM1-dependence of different nuclear export signals. Exp Cell Res 256: 213–224 15. Sharma M, Henderson BR (2007) IQ-domain GTPase-activating protein 1 regulates betacatenin at membrane ruffles and its role in macropinocytosis of N-cadherin and adenomatous polyposis coli. J Biol Chem 282: 8545–8556

Chapter 12 Imaging of Transcription Factor Trafficking in Living Cells: Lessons from Corticosteroid Receptor Dynamics Mayumi Nishi Abstract Adrenal corticosteroids (cortisol in humans/corticosterone in rodents) readily enter the brain and exert markedly diverse effects, such as the stress response of target neural cells. These effects are regulated via two receptor systems, the mineralocorticoid receptor (MR) and the glucocorticoid receptor (GR), both are ligand-inducible transcription factors. GR and MR predominantly reside in the cytoplasm in the absence of corticosterone (CORT), but are quickly translocated into the nucleus upon binding CORT. Then these receptors form dimers to bind hormone responsive elements and regulate the expression of target genes. Given the different actions of MR and GR in the central nervous system, it is important to elucidate how the trafficking of these receptors between the cytoplasm and nucleus and their interaction are regulated by ligands or other molecules to exert transcriptional activity. However, these processes have still not been completely clarified. To address these issues, we have tried to observe more dynamic subcellular trafficking processes in living cells by employing a green fluorescent protein (GFP). In this chapter, we describe our recent studies of corticosteroid receptor dynamics in living cells focusing on three points: (1) time-lapse imaging of GFP-labeled corticosteroid receptors; (2) intranuclear dynamics of GFP-labeled corticosteroid receptors using the fluorescence recovery after photobleaching (FRAP) technique; and (3) the possibility of heterodimers formation using the fluorescence resonance energy transfer (FRET) technique. These studies demonstrate that GR and MR were quickly translocated from the cytoplasm to nucleus after CORT treatment. The time course of the nuclear translocation of GR and MR differed depending on the concentration of CORT. The FRAP study showed that liganded GR and MR in the nucleus were highly mobile, and not trapped by specific organelles. We detected GR-MR heterodimers, which were affected by changes in CORT concentrations in response to various hormonal milieu such as circadian rhythm and stress. Our findings may provide new insights into the dynamic status of corticosteroid receptors in living cells and the molecular basis of the regulation of stress by these receptors. Key words: GR, MR, Importin, GFP, FRAP, FRET, Living cell imaging, Nuclear localization

Paul J. Higgins (ed.), Transcription Factors: Methods and Protocols, Methods in Molecular Biology, vol. 647, DOI 10.1007/978-1-60761-738-9_12, © Springer Science+Business Media, LLC 2010

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1. Introduction Brain corticosteroid receptors were discovered by McEwen and colleagues (1) in the limbic system, particularly the hippocampus where corticosterone (CORT) was retained in large amounts. The glucocorticoid receptor (GR) and mineralocorticoid receptor (MR) show a high degree of colocalization in the hippocampus (2, 3). Since MR has about tenfold higher affinity for CORT than does GR, hippocampal MR responds strongly to CORT (4). Thus, in the hippocampus, one compound, CORT, serves to regulate two signaling pathways via MR and GR (5). The progressive activation of MR at low CORT concentrations and additional activation of GR when CORT levels increase might cause extreme changes in neuronal integrity for responding to stress and in neuronal excitability associated with changes in neuroendocrine regulation and behavior (6). These corticosteroid receptors predominantly reside in the cytoplasm in the absence of ligands associated with various chaperone proteins such as heat shock protein 90 (hsp90) (7). Upon binding with a hormone, their conformation changes dramatically, the nuclear localization signal (NLS) masked by hsp90 is unmasked, and the receptors translocate into the nucleus. For inducing transactivation, the hormone–receptor complex binds to hormone responsive elements (HREs) in the promoter region in a homodimer or a heterodimer (8). Thus, the elucidation of mechanisms for the subcellular and subnuclear trafficking of these receptors is remarkably important for understanding the biological activities of the receptors. In this review, we describe how to analyze the dynamics of GR and MR in neural cells and nonneural cells using living cell imaging techniques (9, 10).

2. Materials 2.1. Plasmid Construct

1. Vectors are obtained commercially that have multiple cloning sites which allow a cDNA or genomic sequence of interest to be placed in-frame at the carboxyl or amino terminus of the GFP-coding region (e.g., pAcGFP1-N1, 2,3; pAcGFP1C1, 2,3; pAmCyan1-N1; pAmCyan1-C1; pZsYellow1-N1; pZsYellow1-C1, Clontech). The choice of vector depends on the location of critical regions of interaction or folding in the protein under investigation. 2. QuickChange Site-Directed Mutagenesis Kit (Stratagene, La Jolla, CA). Quick Gel Extraction Kit (QIAGEN). DNA ligation Kit Ver.2 (TAKARA Bio).

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3. OligoDNA primers designed for subcloning cDNA of the protein under investigation into vectors are supplied by various companies. 2.2. Transfection

1. Although optimal transfection procedures (e.g., calcium phosphate, electroporation, lipofection) vary depending on cell type, the simple and easy lipofection method is used here. Various kinds of lipofection reagents are commercially available; Lipofectamine Plus (BD Biosciences); FuGENE (Roche Applied Science) etc. 2. OPTI MEM (Invitrogen, Carlsbad, CA) for transfection medium.

2.3. Cell Culture

1. Dulbecco’s Modified Eagle’s Medium (DMEM) (Gibco/ BRL, Bethesda, MD) supplemented with bovine serum (FBS, HycClone, Ogden, UT). 2. Dissecting solution: 0.8% NaCl, 0.04% KCl, 0.006% Na2HPO4⋅12H2O, 0.003% KH2PO4, 0.5% glucose, 0.00012% phenol red, 0.0125% penicillin G, and 0.02% streptomycin for primary culture. 3. Neurobasal Medium without phenol red (Invitrogen, Carlsbad, CA) supplemented with 2 mM glutamine and B-27 (1:50, Invitrogen) for neuronal primary cultures. 4. Culture dish: coverslip bottom dishes (Glass Bottom No.0: coverslip bottom diameter of 10 mm2 or Glass Bottom No.1.5: coverslip bottom diameter of 35 mm2; MatTeK Corporation, Ashland, MA). 5. Polyethylenimine (Sigma, St. Louis, MO), poly-D-lysine, or collagen for plate coating.

3. Live Cell Imaging 1. Quantix high-resolution cooled CCD camera (Photometrics, Tucson, AZ) attached to a microscope (IXL70, Olympus, Tokyo) equipped with an epifluorescence attachment. 2. Filter sets: GFP fluorescence: a 480-nm excitation filter, a 515-nm emission filter, and a 505-nm dichroic mirror (Olympus); YFP fluorescence: a 500AF25 excitation filter, a 545AF35 emission filter, and a 525DRLP dichroic mirror (Omega Optical, Inc., Brattleboro, VT); CFP fluorescence: a 440AF21-nm excitation filter, a 480AF30-nm emission filter, and a 455DRLP dichroic mirror (Omega Optical, Inc), etc. FRET filter set (XF88, Omega), which consisted of a 440AF21

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excitation filter for the donor, a 455DRPL dichroic mirror and a 535AF26 emission filter for the acceptor. 3. LSM 510 META confocal microscope (Carl Zeiss, Jena, Germany) equipped with an argon ion laser and a heliumneon laser. 4. CO2 incubator attached on the microscope stage (e.g., MODEL CZI-3, Carl Zeiss). 5. Image analysis software: Meta Morph and Meta Flour (Universal Imaging Corporation, PA).

4. Methods 4.1. Plasmid Construction

1. A cDNA fragment containing the desired gene is obtained by introducing a suitable restriction enzyme site just upstream of the first ATG in the gene cloned into an appropriate vector with a QuickChange Site-Directed Mutagenesis Kit according to the Manufacturer’s instructions using oligonucleotide primer sets (11). 2. Plasmid DNA was extracted in a small-scale preparation to confirm the insertion of the restriction enzyme site, and then extracted in a large-scale preparation. 3. The plasmid obtained and GFP vector mentioned in the Methods are digested with the same restriction enzymes and subjected to agarose gel electrophoresis for isolating the desired bands. Then the DNA fragments are extracted with the QIAquick Gel Extraction Kit. Commercially available vectors are usually supplied with cytomegalovirus (CMV) or simian virus-40 (SV40) promoters, both of which are strong and drive high-level expression. In particular cases when low expression levels are needed, it may be necessary to replace such promoters with weaker promoters such as SV2 (12) or endogenous promoter sequences. 4. The DNA fragment of interest and GFP vector are ligated with DNA Ligation Kit Ver.2. 5. Finally, plasmids are extracted and purified in a large-scale preparation using NucleoSpin Tissue (Macherey-nagel, Easton, PA) according to the manufacturer’s instructions. The sequence of expression plasmids is recommended (see Note 1).

4.2. Cell Culture and Transfection

1. Dissociated hippocampal primary neuronal cultures are prepared from 18-day-old Sprague-Dawley rat fetuses according to the previous methods (13). The fetuses are removed from the placenta in a laminar flow hood and transferred to ice–cold

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dissecting solution. The isolated hippocampus is mechanically dissociated by triturating through a fire-polished glass pipette. The dissociated cells are plated on 35-mm glass bottom dishes precoated with 0.1 mg/ml of polyethylenimine at an initial plating density of 5 × 105 cells/cm2 by adding 200 ml of the cell suspension to each well (glass bottom area of 0.78 cm2; Glass Bottom No.0; MatTeK Corporation, Ashland, MA) (see Note 2). The cultures are maintained in Neurobasal/B27 medium in a CO2 incubator at 37°C with 5% CO2/95% air. 2. COS-1 cells are maintained in DMEM without phenol red, supplemented with 10% FCS overnight in 35-mm glass bottom dishes well (Glass Bottom No.0; MatTeK Corporation) at an initial plating density of 1 × 104 cells/cm2 in 200 ml of medium. 3. Plasmid DNA is transiently transfected into cells by a liposomemediated method using LipofectAMINE PLUS/LTX, according to the manufacturer’s instructions. The amount of plasmid DNA and transfection conditions depend on the cell type. 4. Western blotting and immunocytochemistry by using GFP antibody or antibodies specific for the original proteins should be conducted to confirm whether the fusion proteins have the desired molecular weight and proper subcellular localization. 4.3. Examination of Transcriptional Activity

1. In order to elucidate the functional properties of the fusion proteins, the construct is co-transfected with a reporter MMTV-Luc into COS-1 cells. As an internal standard, the b-galactosidase gene was also co-transfected. COS-1 cells plated on 35-mm dishes were co-transfected with 1 mg of mouse mammary tumor virus promoter (MMTV)-Luc reporter and 1 mg of GFP-MR, YFP-MR, or CFP-GR by lipofection. One microgram of pCH110, a mammalian positive control vector for the expression of b-galactosidase, is also co-transfected as an internal standard. 2. Cell lysate is centrifuged at 12,000 rpm for 2 min at 4°C, and the luciferase activity of the resulting supernatant is assayed at 25°C using the luciferase assay system Pica Gene (Toyo Inki, Tokyo) according to the manufacturer’s protocol, and normalized to b-galactosidase activity. The maximum level obtained with 10-7M CORT for 18 h was taken as 100 after the normalization, and relative values for reporter luciferase activity are plotted.

4.4. Live Cell Imaging 4.4.1. Time-Lapse Imaging

1. For the living cell imaging experiments, the culture medium is replaced with CO2-independent OPTI MEM and images acquisition are acquired in a temperature-controlled room at

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37°C (see Notes 3 and 5). When using a CO2 incubator attached to the stage of the microscope, the culture medium does not need to be replaced (see Note 6). 2. Images are acquired using a high-resolution, cooled CCD camera attached to a microscope equipped with an epifluorescence attachment or a LSM510 META confocal laser microscope (see Note 4). GFP fluorescence is observed using a filter set with 480 nm excitation and 515 nm emission, and a 505 nm dichroic mirror. YFP fluorescence is observed using a filter set with 500 nm excitation and 545 nm emission, and a 525 nm dichroic mirror and CFP fluorescence using a filter set with 440 nm excitation and 480 nm emission, and a 455 nm dichroic mirror. In the time-lapse analysis, images are captured using the time-lapse program of IPLab Spectrum or Meta Morph. For the high-resolution analysis, an image deconvolution procedure (Meta Morph) is applied to a series of images. The “nearest neighbor estimate” is calculated from the raw data. An example of results produced by the timelapse analysis of CFP-GR and YFP-MR is shown in Fig. 1.

Fig. 1. Dual-color time-lapse imaging of GR and MR with GFP spectral variants in single COS-1 cells. COS-1 cells co-transfected with CFP-GR and YFP-MR were cultured in the absence of serum and steroids for 24 h before observation, and then the culture medium was replaced with SFM buffered with HEPES for the time-lapse study. (a, c) Representative fluorescence images of CFP-GR. (b, d) Representative fluorescence images of YFP-MR. Cells shown in (a) and (b) were treated with 10-6M CORT, while cells shown in (c) and (d) were exposed to 10-9M CORT. Note that in the presence of 10-6M CORT, CFP-GR and YFP-MR showed essentially the same nuclear accumulation rates, whereas YFP-MR was accumulated in the nuclear region faster than CFP-GR in the presence of 10-9M CORT. These results suggest that the difference in trafficking kinetics detected in the presence of 10-9M CORT reflects the difference in affinity for CORT between MR and GR: more specifically, MR has about tenfold higher affinity for CORT than that of GR. The findings also suggest that both MR and GR are saturated in cells treated with 10-6M CORT, causing the lack of difference in trafficking kinetics.

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3. Data are evaluated with the image analysis software program, IPLab Spectrum or Meta Morph. In order to measure nuclear/cytoplasmic ratios of fluorescence intensity, data are collected and quantified using a line intensity profile across the cell. For each set of conditions, the average intensities of the pixels are collected within the individual nuclei and cytoplasm of at least five cells from three independent experiments. Nuclear/cytoplasmic fluorescence ratios are calculated for each time point. The results are normalized relative to the value at 0 min taken as 1. 4.4.2. FRAP Analysis

In a typical FRAP analysis, a small region of the specimen (region of interest; ROI) is exposed to photobleaching by an intense laser, usually at maximum intensity. The recovery of fluorescence from surrounding unbleached fluorophores into the area being photobleached is then measured using imaging intensity, with the laser at a lower power. The interval between image scans varies depending on the duration of recovery in an initial pilot experiment. The fraction of labeled proteins that participate in the recovery, called the mobile fraction, can be measured. 1. In qualitative FRAP experiments, the entire cell with the structure of interest is imaged before the bleaching and during recovery. Images are collected at comparatively long intervals (i.e., ~10 s to 1 min). This approach allows the recovery in areas within and outside the bleached zone to be monitored with good morphological resolution and provides an assessment of the overall effect of photobleaching on the cell. 2. In quantitative analyses, fluorescence recovery is only imaged within the bleached area, ROI, typically a 2 to 4 mm wide strip across the cell. This allows fluorescence intensities to be acquired very rapidly (i.e., every 0.5 s), which is crucial for an accurate determination of half-recovery time. The mean intensity in the ROI is plotted versus time, where the time (half-time) indicates the speed of this mobility, i.e., diffusion time, and the level of fully recovered intensity gives information on mobile/immobile species of the fluorescent molecule (14). An example of FRAP experiments using GFP-GR and GFP-MR in cultured hippocampal neurons is shown in Fig. 2.

4.4.3. FRET Analysis

FRET is the radiationless transfer of excited-state energy from an initially excited donor (in this case, CFP) to an acceptor (in this case, YFP) (15, 16). It depends on the proper spectral overlap of the donor and acceptor, their distance from each other (>10 nm), and the relative orientation of the chromophore’s transition dipoles. Here we will introduce an analysis of protein–protein

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Fig. 2. FRAP analyses of GFP-GR and GFP-MR in cultured hippocampal neurons. Defined regions marked with a circle having cultured hippocampal neurons with a diameter of 2 mm transfected with GFP-GR (a) and GFP-MR (b) were exposed to 100% laser intensity for 100 iterations. After the photobleaching, images were detected every 1 s using the laser at a lower power. The initial fluorescence immediately after bleaching was normalized to 0, and the final fluorescence at equilibrium was designated as 1. Then the mean value of the half-maximal recovery time (t1/2) was calculated from ten cells in three independent experiments.

interactions between CFP-GR and YFP-MR using GFP-based FRET microscopy in COS-1 cells and cultured hippocampal neurons. FRET is evaluated by using three different methods; (i) ratio imaging, (ii) emission spectra by emission finger printing method using a LSM 510 META (Zeiss), and (iii) acceptor photobleaching (10). In all FRET experiments, cells showing nearly the same fluorescence intensity in the donor and acceptor were selected for analysis. A summary of the three methods is shown in Fig. 3. 4.4.4. Ratio Imaging

1. For ratio imaging with FRET microscopy using a fluorescence microscope, images are taken with the donor filter set for CFP described above and with a FRET filter set (XF88, Omega), which consisted of a 440AF21 excitation filter for the donor, a 455DRPL dichroic mirror and a 535AF26 emission filter for the acceptor. Images are captured with both filter sets under identical conditions. 2. Ratio images are calculated by dividing FRET (acceptor filter image) by CFP (donor image) using MetaMorph software according to the manufacturer’s instructions after appropriate background subtraction. Background fluorescence is measured in a space in which no cell is present, and total fluorescence is then subtracted from background fluorescence. Ratio images are constructed with a numerator image (FRET image) and a denominator image (donor image), whereby the ratio of the intensity of a pixel from the two images is obtained. Then the ratio images are pseudocolored with red

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Fig. 3. The scheme of principle of intermolecular FRET and methods of evaluating FRET. FRET is the radiationless transfer of excited-state energy from an initially excited donor (in this case, CFP) to an acceptor (YFP). It depends on the proper spectral overlap of the donor and acceptor, their distance from each other (>10 nm), and the relative orientation of the chromophore’s transition dipoles. The bottom scheme show three evaluation methods; (i) ratio imaging, (ii) emission spectra by Emission Fingerprinting using LSM 510 META (Zeiss), and (iii) acceptor photobleaching.

indicating a high ratio and blue indicating a low ratio. To prevent the detection of false-positive FRET images, the imaging conditions are adjusted to favor donor emission over acceptor emission. We should confirm that the level of bleedthrough of CFP and YFP in our filter sets is very low (17). The FRET value is calculated by various measures: Ff/Df; Fc/Df (see Note 11) (18). Fc represents a calculated FRET value termed “correct FRET.” Fc = Ff – Df(Fd/Dd) – Af(Fa/Aa) Dd: Signal from a donor-only specimen using the donor filter cube. Fd: Signal from a donor-only specimen using the FRET filter set. Ad: Signal from a donor-only specimen using the acceptor filter set. Da: Signal from an acceptor-only specimen using the donor filter set. Fa: Signal from an acceptor-only specimen using the FRET filter set.

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Aa: Signal from an acceptor-only specimen using the acceptor filter set. Df: Signal from a FRET specimen using the donor filter set. Ff: Signal from a FRET specimen using the FRET filter set. Af: Signal from a FRET specimen using the acceptor filter set. 1. Using the laser microscope LSM510, images are taken by stimulating with a 458 nm laser employing the dichroic mirror and filter; HFT 458/514, HFT515, band pass filter 475–525, and FRET images are taken by stimulating with a 458 nm laser employing the combination of dichroic mirror and filter; HFT 458/514 and band pass filter 520–560. To make ratio images, FRET images are divided by donor images. 4.4.5. Emission Spectra

1. For detecting emission spectral changes in FRET imaging, an emission finger printing method using the confocal laserscanning microscope LSM 510 META is employed. First, spectral signatures of the fluorescence within the specimen are captured by means of lambda stack acquisition; excitation at 458 nm and detection at 10 nm-intervals from 460 to 596 nm using an HFT 458/543 dichroic mirror (9). 2. Several regions of interest (ROIs) with a diameter of 1–2 mm are randomly selected for drawing emission spectral patterns, and the mean ratio of fluorescence intensity at 527 and 474 nm is calculated from selected ROIs at each time point after ligand addition (20 ROIs per cell, in ten cells from three independent experiments). Since the level of protein expression in each cell is not exactly the same, especially between donor and acceptor molecules, the fluorescence intensity should be normalized in each cell by dividing the mean ratio of fluorescence intensity after ligand treatment by that before ligand treatment.

4.4.6. Acceptor Photobleaching

1. Photobleaching will cause the acceptor to lose its capacity to absorb energy from the donor, causing the donor to surge to the maximum as if there is no FRET. This will confirm that the emission detected by the FRET channel comes from the true FRET, and is not due to channel cross talk or cross excitation of the acceptor by the donor excitation light. The acceptor is photobleached by using a 514-nm laser at 100% power after 60 and 90 min of 10-6M and 10-9M CORT treatment, respectively. Then the cells are subjected to an emission spectral analysis as described above for detecting the change in fluorescence intensity of the donor molecule. The increase in donor fluorescence intensity is shown as a percentage.

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Fig. 4. GFP-based FRET analysis of protein–protein interaction between CFP-GR and YFP-MR. Representative results of ratio imaging analyses. COS-1 cells were co-transfected with CFP-GR and YFP-MR. Images of donor, FRET and ratio (FRET/donor) were captured at the indicated time after treatment with 10-6M CORT. Areas marked by rectangles in the nucleus of 30 and 60 min were enlarged as insets. Note a red hue showing a positive FRET sign in the nucleus indicates a heterodimer, while very little red hue in the cytoplasm indicates a very low incidence of heterodimerization. The area and intensity of red hue at 60 min after CORT treatment were more than those at 30 min. Bar = 10 mm.

An example of FRET analyses investigating the possible heterodimerization of GR and MR is shown in Fig. 4.

5. Notes 1. If the fusion proteins are not fluorescent or nonfunctional, there are several reasons; the fluorescent protein is not fused in the correct frame; the expression level of the fusion proteins is too low; the fusion protein is unstable. We must confirm the primer construction, and sequence of expression plasmids. It may be also desirable to generate two fusions: one at the amino terminus and one at the carboxyl terminus. 2. For successful live cell imaging, one must plate cells in dishes with coverslip bottoms. They come uncoated or coated with poly-D-lysine or collagen. One should examine which coating is best for the cells. Although many well-slides claim to be good for imaging, most are still too thick for use with high magnification high numerical aperture objectives. A thickness of 0.17–0.18 mm is recommended. 3. The pH indicator phenol red can interfere with the collection and interpretation of weak fluorescent signals. For best results, the cells should be grown in a phenol red-free medium. 4. Although live cell imaging can be done with a number of different systems, an inverted microscope is much more suitable than an upright microscope. The choice of microscope depends on your needs. If you need to observe very rapid events, images should be acquired with exposure times as short as possible, around 30 ms, per plane, which makes it

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possible to image the entire volume of a cell with z-steps in the submicron range in as little as 1–2 s per channel. If a very high speed is not required, the Zeiss LSM510META system is excellent for confocal imaging of living cells. 5. When performing live cell imaging experiments, one critical point is to maintain the cells in a healthy state with normal function while they are on the microscope stage. Control of the cells’ environment is vital to the success of live cell experiments. Cells that appear even slightly unhealthy should not be used for imaging and data collection. 6. Cells are typically cultured in a cell incubator at 37°C with 5% CO2. The pH value of NaHCO3-buffered media depends on the CO2 content of the incubator’s atmosphere. When the CO2 supply to an incubator fails, media become alkaline and cells are adversely affected and may die. We can image cells in their normal medium if we supply 5% CO2 to the dish on the microscope stage. However, it is easiest to use a HEPESbuffered medium, for instance, OPTI MEM. 7. Cellular function is highly sensitive to temperature. In addition to a stage warmer, objective warmers are required to collect better images. Heating of both the dish and the objective prevents temperature gradients across the dish. We sometimes suffer from instability of z-positioning over time during live cell imaging. This focus drift is mostly due to thermal expansion that occurs due to a temperature gradient. When using high NA objectives, the dish is thermally coupled to the objective by the immersion medium. This is why it is necessary to use not only a stage warmer but also an objective heater if z-stability is needed for an experiment. 8. The most critical aspects of image acquisition are maintaining the same setting and order of exposure for each cell. Particularly in the case of FRET experiments, acquiring images with all three filter sets records all possible information and permits many different levels of analyses. If the software allows, it is convenient to employ macro for acquiring three exposures (FRET, CFP, and YFP) of cells with optimal settings determined in exposure settings calibration. 9. Photobleaching occurs when a fluorophore undergoes irreversible covalent modification and loses its ability to fluoresce. Different fluorophores suffer different numbers of excitation emission cycles before photobleaching. Phototoxicity largely results from the formation of oxygen radicals due to nonradiative energy transfer, and these oxygen radicals can be toxic to cells. To minimize both phototoxicity and photobleaching, minimize the energy level of the excitation light and the duration of excitation. Use as little light as possible, particularly when acquiring an extended time series.

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10. If fusion proteins do not show desired protein–protein interaction, it may be necessary to generate two sites of fusions: one at the amino terminus and one at the carboxyl terminus. Because the FRET signal is highly dependent on the interfluorophore distance, the presence of a fluorophore in one domain of a protein may not induce a FRET signal if another domain is responsible for the protein–protein interaction in question. Performing the experiment with different sites of fusions increases the chance of detecting FRET. 11. There are no rules for deciding which method of calculation (Ff/Df, Fc/Df) should be adopted, and the most reasonable choice often depends on the specific experiment. The influence of many aspects of an experiment on the different methods of calculations has been discussed. Ideally, the results do not depend on the calculation, as shown explicitly in some cases. In general, the simpler calculations involving fewer measurements are preferable.

Acknowledgments The author would like to thank Professor Kawata for his advice and encouragement. This work was supported by Grant-in Aid from Scientific Research from MEXT 19300120. References 1. McEwen BS, de Kloet ER, Rostene W (1986) Adrenal steroid receptors and actions in the nervous system. Physiol Rev 66:1121–1187 2. Arriza JL, Simerly RB, Swanson LW, Evans RM (1988) The neuronal mineralocorticoid receptor as a mediator of glucocorticoid response. Neuron 1:887–900 3. de Kloet ER, Vreugdenhil E, Oitzl MS, Joels M (1998) Brain corticosteroid receptor balance in health and disease. Endocr Rev 19:269–301 4. Rupprecht R, Reul JMHM, van Steensel B, Spengler D, Soder M, Berning B, Holsboer F, Damm K (1993) Pharmacological and functional characterization of human mineralocorticoid and glucocorticoid receptor ligands. Eur J Pharmacol 247:145–154 5. Kawata M (1995) Roles of steroid hormones and their receptors in structural organization in the nervous system. Neurosci Res 24:1–46 6. Magarinos AM, McEwen BS, Flugge G, Fuchs E (1996) Chronic psychosocial stress causes apical dendritic atrophy of hippocampal CA3

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pyramidal neurons in subordinate tree shrews. J Neurosci 16:3534–3540 Yang J, DeFranco DB (1996) Assessment of glucocorticoid receptor-heat shock protein 90 interactions in vivo during nucleocytoplasmic trafficking. Mol Endocrinol 10:3–13 Umesono K, Evans RM (1989) Determination of target gene specificity for steroid/thyroid hormone receptors. Cell 57:1139–1146 Nishi M, Ogawa H, Ito T, Matsuda KI, Kawata M (2001) Dynamic changes in subcellular localization of mineralocorticoid receptor in living cells: in comparison with glucocorticoid receptor using dual-color labeling with green fluorescent protein spectral variants. Mol Endocrinol 15: 1077–1092 Nishi M, Tanaka M, Matsuda K, Sunaguchi M, Kawata M (2004) Visualization of glucocorticoid receptor and mineralocorticoid receptor interactions in living cells with GFPbased fluorescent resonance energy transfer (FRET). J Neurosci 24:4918–4927

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11. Matsuda K, Ochiai I, Nishi M, Kawata M (2002) Colocalization and ligand-dependent discrete distribution of the estrogen receptor (ER)alpha and ERbeta. Mol Endocrinol 16:2215–2230 12. Janicki SM, Tsukamoto T, Salghetti SE, Tansey WP, Sachidanandam R, Prasanth KV, Ried T, Shav-Tal Y, Bertrand E, Singer RH, Spector DL (2004) From silencing to gene expression: real-time analysis in single cells. Cell 116:683–698 13. Nishi M, Takenaka N, Morita N, Ito T, Ozawa H, Kawata M (1999) Real-time imaging of glucocorticoid receptor dynamics in living neurons and glial cells in comparison with nonneural cells. Eur J Neurosci 11:1927–1936 14. Lippincot-Schwartz J, Snapp E, Kenworthy A (2001) Studying protein dynamics in living cells. Nat Rev Mol Cell Biol 2:444–456

15. Periasamy A, Day RN (1999) Visualizing protein interactions in living cells using digitized GFP imaging and FRET microscopy. Methods Cell Biol 58:293–314 16. Miyawaki A (2003) Visualization of the spatial and temporal dynamics of intracellular signaling. Dev Cell 4:295–305 17. Tanaka M, Nishi M, Morimoto M, Kawata M (2003) Nuclear import of glucocorticoid receptor in association with importin a and importin b: analysis with real-time fluorescence imaging and fluorescence resonance energy transfer in living cells. Endocrinology 144:4070–4079 18. Gordon GW, Berry G, Liang XH, Levine B, Herman B (1998) Quantitative fluorescence resonance energy transfer measurements using fluorescence microscopy. Biophys J 74: 2702–2713

Chapter 13 Hypoxia-Inducible Factors: Post-translational Crosstalk of Signaling Pathways Elitsa Y. Dimova and Thomas Kietzmann Abstract Hypoxia-inducible factor-1 (HIF-1) has a central role in the mammalian program by which cells respond to hypoxia in both physiological and pathological situations. HIF-1 transcriptional activity, protein stabilization, protein–protein interaction, and cellular localization are mainly modulated by Post-translational modifications such as hydroxylation, acetylation, phosphorylation, S-nitrosylation, and SUMOylation. Here, we summarize current knowledge about Post-translational HIF-1 regulation and give additional information about useful methods to determine some of these various modifications. Key words: Hypoxia, Hypoxia-inducible factors, Hydroxylation, PHD, Acetylation, Phosphorylation, MAPK, PI3K

1. Introduction 1.1. HIFs Family: Basic Biology

One of the most important proteins involved in the mammalian response to oxygen deficiency is hypoxia-inducible factor-1 (HIF-1), which belongs to the basic helix-loop-helix/Per-ARNT-Sim (bHLH/PAS) family of transcription factors (for a review on the PAS protein family, see ref. (1)). HIF-1 is a heterodimer of an oxygen-sensitive HIF-1alpha and a constitutively expressed HIF-1beta (also known as arylhydrocarbon receptor-nuclear translocator, ARNT) subunit and binds hypoxia-responsive elements (HREs) within promoters or enhancers of its target genes. The HREs are represented by the consensus sequence 5¢-BACGTSSK-3¢ (B = G/C/T; S = G/C; K = G/T) (2), which contains the core sequence 5¢-RCGTG-3¢ (3). In addition to the HIF-1alpha and HIF-1beta subunits, two other HIF-alpha subunits (HIF-2alpha

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and HIF-3alpha) and two other HIF-1beta subunits (ARNT2 and ARTN3) have been identified and may give rise to different HIF heterodimers (for review see ref. (4)). The human HIF-1alpha subunit consists of 826 amino acids (aa) with an approximate molecular weight of 120 kDa (5). It contains two nuclear localization sequences responsible for translocation of HIF-1alpha to the nucleus under hypoxia; the -N-terminal (aa 17–33) and C-terminal (aa 718–721) (6), respectively. HIF1alpha transactivity is determined by two transactivation domains (TAD); -N-terminal (TADN, aa 531–575) and C-terminal (TADC, aa 786–826) (7–9), respectively. The amino acid residues 576–785 comprise an inhibitory domain, as its deletion was shown to increase TAD function under normoxia (7). An oxygen-dependent degradation domain ODDD (aa 401–603) is subject of Posttranslational modifications and has an impact on hypoxia-dependent stabilization of HIFs (10). Several HIF-1alpha splice variants have been reported (for review see ref. (11)). Of particular interest are HIF-1alpha516, HIF-1alpha557, and HIF-1alpha735 that terminate respectively at codon 516, 557, and 735, resulting in the absence of both TADN and TADC, or of only TADC (12, 13). However, the biological significance of these isoforms is yet unclear. The HIF-2alpha subunit, also known as endothelial PerARNT-Sim (PAS) protein (EPAS) (14), HIF-like factor (HLF) (15), HIF-related factor (HRF) (16), and member of the PAS superfamily 2 (MOP2) (17) is an 874 amino acids containing protein, which shares a high degree of sequence homology and virtually all of the functional domains with HIF-1alpha. Although HIF-1alpha and HIF-2alpha are quite similarly regulated in human cells in vitro (18), they have distinct functions that overlap only partially in vivo. In tissues expressing both HIF-1 and HIF-2, HIF-2 is considered to be the main regulator of EPO expression (19–21) and VEGF expression (22–24). Recently, experiments with HIF-1alpha/HIF-2alpha chimeric proteins, where TADN and/or TADC domains of each paralogue were “swapped,” showed that the respective TADN appears to be involved in HIF-1 and HIF-2 target gene specificity (25). Interestingly, the replacement of the TADC between HIF-1alpha and HIF-2alpha had no effect on the target transcripts measured, whereas substitution of both the TADN and the TADC effectively caused HIF-1alpha to behave as HIF-2alpha, and vice versa (for review see ref. (26)). In contrast to the ubiquitously expressed HIF-1alpha, HIF2alpha is predominantly expressed in the lung, the endothelium, and the carotid body (15, 27). Indeed, the HIF-2alpha expression is restricted to particular cell types such as glial cells, vascular endothelial cells, cardiomyocytes, lung type II pneumocytes, hepatocytes, and interstitial cells of the kidney (28). The importance of HIF-2alpha in catecholamine homeostasis, vascular remodeling, and lung maturation during neonatal development

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in vivo has been demonstrated by the independent generation of three knockout HIF-2alpha mice (27, 29, 30). The third HIFalpha protein, HIF-3alpha was cloned from mouse (31), rat (32), and human (33). In contrast to the other proteins from the HIF family, it does not contain the TADC and its function remains unclear. Several alternatively spliced variants of HIF-3 have been identified in both mouse and human (34–37). The best characterized is the so called mouse inhibitory PAS protein (IPAS), which lacks both TADN and TADC (34–37) and exerts an inhibitory effect on HIF-1 by binding either to HIF-1alpha, thereby preventing the interaction between HIF-1 and HREs of target genes, or by binding to HIF-1beta thus serving as HIF-1alpha antagonist (34, 35, 37). While HIF-3alpha is found predominantly expressed in the adult thymus, lung, brain, heart, and kidney (31), IPAS is detected mainly in the Purkinje cells of the cerebellum and the corneal epithelium of the eye in adult mice (34, 35). By contrast, in murine heart and lung tissues, IPAS mRNA is hypoxia-regulated, indicating a negative feedback mechanism that controls HIF-1alpha activity (34–36). 1.2. HIF Regulation

The predominant mode of HIF-alpha regulation appears to be Post-translational although regulation at the transcriptional and translational level was shown (32, 38–42). Post-translational modifications such as hydroxylation, acetylation, phosphorylation, S-nitrosylation, and SUMOylation have been proved to influence not only protein stability but also the transcriptional activity of HIFs (1, 2).

1.2.1. Hydroxylation

Hydroxylation is characterized by the introduction of hydroxyl group(s) into a protein. The principal residue to be hydroxylated in proteins is proline, hence forming hydroxyproline. Although proline hydroxylation is best known from collagenes, it also became a crucial component of the hypoxia response, and the reaction is catalyzed by oxygen-dependent HIF-prolyl hydroxylases. In order to hydroxylate prolines, these oxidases also require iron, alpha-ketoglutarate, and ascorbate as cofactors (43–45). Four different HIF prolyl hydroxylases have been identified so far: PHD1 (prolyl hydroxylase domain 1; EglN2), PHD2 (EglN1), PHD3 (EglN3), and PHD4 (C-P4H-I) (46– 51). Under normoxia, specific prolines within the ODDD of HIFs (Pro 402 and Pro 564 in human HIF-1alpha (52, 53), Pro 405 and Pro531 in human HIF-2alpha, and Pro 490 in human HIF-3alpha (37)) are targets of PHDs. Hydroxylation at these prolines allows binding of the von Hippel–Lindau tumor suppressor protein (pVHL) as part of an E3 ubiquitin ligase complex leading to ubiquitination and proteasome-dependent degradation of HIF alpha subunits (54–59).

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1.2.2. Ubiquitination

Ubiquitin is a small 76 aa molecule acting as a tag that signals the protein-transport machinery to carry the protein to the proteasome for degradation. In addition to polyubiquitination by pVHL, several other proteins have been reported to affect HIF1alpha ubiquitination and stability such as the oncogenic E3 ubiquitin ligase murine double minute 2 (MDM2) (60) and Jab1, a transcriptional coactivator of c-Jun and Jun D (61). In addition to prolines, HIF-1alpha and HIF-2alpha can also be hydroxylated at asparaginyl residues in the TADC (62) by another hydroxylase named factor inhibiting HIF (FIH) (63–65). The asparagine hydroxylation prevents binding of p300/CBP (CREB binding protein) and suppresses HIF transactivity (62).

1.2.3. Acetylation

Acetylation, a reaction introducing an acetyl group into a compound, is a widespread Post-translational modification in eukaryotes and usually occurs at the N-terminal end of a protein. Lysine 532 (K532) located in the ODDD of HIF-1alpha has been reported to be acetylated by an acetyltransferase named arrestdefective-1 (ARD1) (66). This process favors interaction of HIF1alpha with pVHL and thus leads to HIF-1 destabilization (66). The lysine residue acetylated by ARD1 is conserved in HIF1alpha and HIF-2alpha, but not in HIF-3alpha (66). The mRNA and protein levels of ARD1 were shown to be decreased under hypoxia whereas acetyltransferase activity was not influenced by oxygen levels (66). These findings were challenged by another study showing that human ARD1 can bind to HIF-1alpha but does not acetylate and destabilize it (67). Further, it was reported that ARD1 had no impact on the stability of HIF-1alpha or -2alpha and that ARD1 mRNA and protein levels were not regulated by hypoxia in several human tumor cell lines such as HeLa, HT1080, HEK293, and MCF-7 (68, 69). Thus, the role of ARD1 in HIF1alpha modification may involve cell-type specific factors.

1.2.4. Phosphorylation

Phosphorylation is another crucial Post-translational modification for HIF alpha-subunits affecting their transcriptional activity and stability. Under normoxia, HIF-1alpha expression and activity were shown to be regulated by mitogen-activated protein kinase (MAPK/ERK) and phosphatidylinositol 3-kinase/Akt signaling (for review see ref. (70)). The MAPK signaling pathway predominantly induces HIF-1alpha transcriptional activity via direct phosphorylation (71–74). Additionally, the MEK1 inhibitor PD98059, the p42/p44 inhibitor U0126, and the p38MAPK inhibitor SB203580 decreased HIF-1-dependent gene expression (75–77). While the role of ERK in HIF-1alpha phosphorylation appears to be clear, the involvement of p38 MAPK or c-Jun N-terminal kinase seems to vary since one study showed that HIF-1alpha was not phosphorylated by p38 MAPK or c-Jun N-terminal kinase (71) whereas other studies reported that JNK

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(78) and p38 kinase (72) contribute to the activation of HIF1alpha/HIF-2alpha. The phosphorylation events exerted by ERKs seem to be independent from the HIF-1alpha degradation pathway since p42/p44 are mainly located in the nucleus whereas degradation takes place in the cytoplasm (42). The ERK phosphorylation sites were initially mapped to the TADC (aa 531– 826) of HIF-1 (72–74) and later on shown to specifically target Ser-641 and Ser-643 within HIF-1alpha (79). Intriguingly, inhibition of these phosphorylation sites impaired not only HIF1alpha activity but also the nuclear localization (79, 80). Actually, a MAPK-targeted, atypical but CRM1-dependent NES in human HIF-1alpha (aa 616–658) has been identified (81). Thus phosphorylation of HIF-1alpha by ERK provides an additional means of HIF-1a regulation. The phosphatidylinositol 3-kinase (PI3K)/Akt cascade has been suggested to control both HIF-1alpha transcriptional activity and protein synthesis. Constitutively active PI3K and Akt, as well as loss of PTEN, appear to enhance HIF-1 activity both under hypoxia and normoxia (82–84). Actually, PI3K/Akt do not directly phosphorylate HIF-1alpha, but downstream components of Akt such as mammalian target of rapamycin (mTOR) (85), glycogen synthase kinase 3 (GSK3) (86), and mouse double minute homologue (HDM2) (87) were described to either activate or inactivate HIF-1alpha. The mTOR kinase has been shown in vitro to be a positive regulator of HIF-1-dependent gene transcription under hypoxia whereas the mTOR inhibitor rapamycin decreased hypoxia-induced HIF-1alpha protein levels involving the ODD domain of HIF-1alpha (85). In addition, mTOR has been reported to enhance HIF-1 transcriptional activity directly via the regulatory associated protein of mTOR (Raptor) interacting with an mTOR signaling (TOS) motif located in the N-terminal of HIF1alpha; this regulation occurred independent from the VHL-degradation pathway (88). Further, GSK3 is negatively regulated by Akt and can directly phosphorylate S551, T555, and S589 within the ODD domain of HIF-1alpha and thus leading to HIF-1alpha destabilization by promoting proteasomal degradation independent of prolyl hydroxylation and VHL binding (86, 89). In addition, HDM2 directly interacts with HIF-1alpha preventing destabilization of HIF-1alpha independently of pVHL (87). Thus, the PI3K/Akt cascade may interfere with HIF-1alpha regulation at different levels due to the involvement of different Akt targets. 1.2.5. S-Nitrosylation

S-nitrosylation is another fundamental mechanism for Posttranslational control of protein activity and represents redoxrelated modification of protein Cys residues by nitric oxide (NO). NO is a messenger with the ability to stabilize HIF-1alpha and to transactivate HIF-1alpha under normoxia (90, 91). One mechanism

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contributing to HIF-1 stabilization is the impairment of the PHDs activity (92, 93). S-nitrosylation of Cys800 has been reported to stimulate HIF-1 transcriptional activity by activating HIF-1alpha interaction with p300 (94). By contrast, recently it has been shown that Cys800 S-nitrosylation of HIF-1alpha decreases p300 binding (95). An alternative pathway, independent of the PHDs, where Cys533 is subjected to S-nitrosylation and thereby interfering with pVHL binding to the ODDD has been also proposed (96). These two mechanisms might coordinately contribute to HIF-1alpha stability in the presence of NO. 1.2.6. SUMOylation

Another Post-translational modification is SUMOylation. Small Ubiquitin-like Modifier or SUMO proteins are a family of small proteins covalently attached to and detached from other proteins in the cell in order to modify their function. SUMOylation of HIF-1alpha has also been described by several groups with conflicting results. On one side, SUMOylation of HIF-1alpha has been suggested to increase both HIF-1alpha stability and transcriptional activity (97, 98). On the other side, it has been suggested that SUMOylation of HIF-1alpha leads to decreased activity and enhanced VHL-mediated ubiquitination (99, 100). Thus, SUMOylation of HIF-1alpha appears to affect HIF-1 activity, which may vary from cell type to cell type.

1.3. Conclusion

Post-translational modifications such as hydroxylation, acetylation, phosphorylation, S-nitrosylation, and SUMOylation influence the transcriptional activity, the protein stabilization, the protein–protein interaction, and the cellular localization of HIF alpha subunits. These responses may act either alone or in concert to influence HIF alpha-subunits. Thereby the functional consequences of each modification may differ from cell type to cell type, which adds an even more complex picture to the regulation of HIF alpha subunit activity. A number of modifications described here are found with HIF-1alpha, HIF-2alpha, or both, and it remains so far open whether the same modification in each subunit will have the same or similar functional consequences, and although our understanding for all these signaling pathways is becoming clearer, the Post-translational crosstalk of these cascades still needs to be investigated in more details.

2. Materials 2.1. Cell Culture Techniques

1. Earle`s minimum essential medium (MEM, PAA) supplemented with 10% (w/v) fetal bovine serum (Biochrom), 1% nonessential amino acids (PAA), and 0.5% (w/v) antibiotic. 2. 1 mM trypsin/EDTA (Sigma).

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3. Normoxia: 16% O2, 5% CO2, and 97% humidity at 37°C in a cell culture incubator. 4. Hypoxia: 8% O2, 87% N2, 5% CO2 (by vol.), and 97% humidity at 37°C in a cell culture incubator or hypoxia workstation (Ruskin). 5. Transfection mixture/plate: 5–10 µg plasmid DNA, 10% CaCl2, 50% HEPES, H2O. 6. Complete protease inhibitors cocktail tablet (Roche) – 1 tablet/10 ml buffer. 7. Transfection reagents: Lipofectene (Invitrogen), Metafectene (Biontex). 2.2. Recombinant Proteins 2.2.1. GST-Recombinant Protein Expression and Purification in Bacteria

1. LB medium: 1% (w/v) tryptone, 0.5% (w/v) yeast extract, 1% (w/v) NaCl; pH 7.0 (with NaOH), autoclave, store at 4°C. 2. Ampicillin (Applichem): 25 mg/ml in H2O, stored in single use aliquots at −20°C, working concentration 60 µg/ml. 3. IPTG (Sigma): 0.1 M in H2O, stored at −20°C, working concentration 0.1 mM. 4. PMSF (Sigma): 200 mM in isopropanol, stable at RT for ca. 9 months. 5. Resuspension buffer: 20 mM Tris–HCl (pH 8), 50 mM NaH2PO4/Na2HPO4 (pH 7); 1 mM PMSF and complete protease inhibitor tablet (Roche) are added always fresh. 6. Elution buffer: 50 mM Tris–HCl (pH 8.5), 20 mM reduced glutathione. 7. 4× SDS buffer: 100 mM Tris–HCl (pH 7.4), 0.05% (w/v) bromophenolblue, 3% (w/v) SDS, 7.5% (v/v) glycerol, 5% (w/v) DDT (fresh). 8. Dialyzation buffer : 20 mM Tris–HCl (pH 7.5), 20% (v/v) glycine. 9. pGEX-5x-1 (Pharmacia), Lysozyme (Applichem), Triton-X (Sigma), Glutathione Sepharose 4B (Amersham), dialysis tube (GIBCO).

2.2.2. 35S-Labeled Protein In Vitro Translation

1. TNT® coupled reticulocyte lysate system (Promega).

2.3. Protein–Protein Interaction Techniques

1. Phosphobuffer: 50 mM Tris–HCl (pH 7.5), 2 mM EDTA (pH 8.0), 2 mM EGTA, 150 mM NaCl, 10 mM Na2PO4, 1% (v/v) Triton-X. The buffer is prepared always fresh before scraping the cells. 0.5 mM DTT, 0.2 mM PMSF, and protease inhibitor cocktail tablet (Roche)(1 per 10 ml) are added right before use.

2.3.1. CoIP

2. [35S] Methionine (Amersham) >1,000 Ci/mmol at 10 mCi/ml.

2. Protein-G Sepharose (Amersham).

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3. Monoclonal antibody against the V5 tag (1:10,000; Invitrogen) and a monoclonal antibody against hemagglutinin (HA; 1:500; Santa Cruz). 2.3.2. GST Pull-Down

1. Buffer 1: 50 mM Tris–HCl (pH 8), 120 mM NaCl, 0.5% (v/v) NP-40. 2. Buffer 2: 20 mM Tris–HCl (pH 8), 100 mM NaCl, 1 mM EDTA, 0.5% (v/v) NP-40. 3. Gluthation Sepharose 4B (Amersham). 4. Phosphoimager Storm 860 & ImageQuant (Molecular Dynamics) or equivalent.

2.4. Hydroxylation 2.4.1. Hydroxylation Activity Assay 2.4.1.1. HIF Hydroxylation

software

1. Lysis buffer: 250 mM sucrose, 50 mM Tris–HCI (pH 7.5) always freshly supplemented with complete protease inhibitors cocktail tablet (Roche) – 1 tablet/10 ml buffer. 2. Reaction buffer: 40 mM Tris–HCl (pH 7.5), 50 mM FeSO4, 1 mM ascorbate, 0.4 mg/ml catalase, 0.1 mM 2-oxoglutarate (unlabeled), 0.5 mM DTT, 2 mg/ml BSA. 3. [5-14C]2-oxoglutarate (Amersham).

2.5. Acetylation

1. [3H]acetyl-CoA (Amersham) – 137 GBq/mmol.

2.5.1. Acetyltransferase Assay

2. Human adrenocorticotropic hormone (Calbiochem), corticotropins fragment 1–24, 0.5 mM. 3. SP Sephadex (Sigma) – 50% slurry in 0.5 M acetic acid.

2.5.2. In Vitro Acetylation Assay

1. Acetylation buffer: 50 mM Tris–HCl (pH 8.0), 0.1 mM EDTA, 1 mM DTT, 10 mM sodium butyrate, 200 µM acetylCoA, 10% glycerol. 2. Polyclonal Signaling).

2.6. Phosphorylation 2.6.1. Growth Factor Treatment/Inhibitory Studies

anti-acetyl-lysine

antibody

(1:1,000,

Cell

1. Human IGF-1 (Sigma) is resuspended in 0.1 M acetic acid and stored at −4°C. 2. Acetic acid (0.1 M). 3. The PI(3)-kinase inhibitor LY294002 and the MEK inhibitor U0126 (Cell Signaling) are dissolved in DMSO as 10 mM stocks, and aliquots are stored at −20°C for up to 3 months. 4. Cell lysis buffer: 50 mM Tris–HCl, 5 mM EDTA (pH 8.0), 150 mM NaCl, 0.5% (v/v) NP-40. The lysis buffer without DTT and protease inhibitors can be stored at 4°C for several mounts; fresh 0.5 mM DTT, 0.2 mM PMSF, and protease inhibitor cocktail tablet (Roche) are always added before use.

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1. Kinase buffer: 0.2 M MOPS (pH 7.4), 0.5 M EDTA, 0.1 M (CH3COO)2. 2. Active GSK-3b (Cell Signaling). 3. ATP (10 mM). 4. [g-32P] ATP (Amersham). 5. GSK substrate (YRRAAVPPSPSLSRHSSPHQS*EDEEE) (1 mg/ml).

3. Methods 3.1. Cell Culture Techniques

3.2. Recombinant Proteins 3.2.1. GST-Recombinant Protein Expression in Bacteria and Purification

For transfection of HepG2 and HEK293T cells with DNA expression vectors, we typically employ either commercially available transfection reagents or the calcium phosphate method (Graham et al. 1973) following the manufacturer’s instructions or the standard protocol, respectively. We normally use 5–10 µg of plasmid DNA for each transfection performed in 10 cm Ø Petri-dishes. 1. A single colony of E. coli BL21 transformed with a vector expressing the GST-fusion protein is picked up and incubated overnight in 5 ml LB-Amp medium at 37°C on a shaker (220 rpm) as a preculture. 2. 1 ml of the preculture is transferred to 250 ml LB-Amp, incubated on shaker until the OD550 reached 0.4–0.5 (~3–4 h), and IPTG is added to induce fusion protein expression. To check the level of expression proteins, reference samples of 1 ml are taken before and after the IPTG induction. 3. The bacteria are further cultivated at 37°C on a shaker for additional 3 h. 4. The bacteria are collected by centrifugation at 2,500 × g at 4°C for 10 min, and the pellet may be stored at −20°C, if necessary, until further purification of the protein. 5. The pellet is placed on ice and completely resuspended in 10 ml of ice-cold resuspension buffer. 6. 10 mg lysozyme is added, and the suspension is gently mixed at 4°C for 30 min. 7. NaCl is added to a final concentration of 300 mM, followed by an ultrasonication on ice carried out in short bursts – 6 × 45 s pulses of 400 W with 1 min intervals between pulses. 8. Triton X-100 is added into the lysate to a final concentration of 1%, and the lysate is further incubated at 4°C for 30 min with gentle mixing.

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9. After centrifugation at 10,000 × g at 4°C for 10 min, the supernatant (reference samples have to be taken from the supernatant and the pellet) containing the expressed soluble fusion protein is incubated with 380 µl 80% Glutathione Sepharose 4B (per 10 ml dissolved fusion proteins) for 2 h at RT at a rocking platform. 10. The suspension is left at RT to sediment, and the supernatant is collected and stored at 4°C in case the protein was not bound to the beads. The beads pellet is washed 4× with cold 1× PBS, followed each time by passive sedimentation. 11. The GST-fusion protein is subsequently eluted 2–3× from the beads with elution buffer at RT for 1 h, in an Eppendorf thermomixer (750 rpm), followed by centrifugation at RT for 2 min, 350 ´ g in a tabletop centrifuge. At each washing/eluation step, a 100 µl reference sample is taken from the supernatant. The eluates and the beads are stored at −20°C. 12. All reference samples, the eluates and the beads are mixed with 2× SDS sample buffer, heated at 95°C for 5 min, separated by SDS–PAGE, and stained with Coomassie Blue R250. 13. The eluates containing the recombinant protein with the correct size are combined and transferred into a dialysis tube, and dialyzed at 4°C overnight against dialyzation buffer. 14. The protein concentration is estimated using the standard Bradford method (Bradford 1976), and protein samples are aliquoted and stored at −80°C. 3.2.2. 35S-Labeled Protein In Vitro Translation

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S-labeled protein is synthesized by incorporation of S-methionine using the TNT® coupled reticulocyte lysate system (Promega) and a suitable plasmid construct as a template according to the manufacturer’s instructions.

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1. The in vitro translation reaction for 35S-human VHL containing 25 µl TNT rabbit reticulocyte lysate, 2 µl TNT reaction buffer, 1 µl TNT T7 RNA polymerase, 1 µl 1 mM amino acid mixture minus Methionine, 2 µl [35S] Methionine, 1 µl RNasin ribonuclease inhibitor (40 u/ml), 1 µg pCMV-HA-VHL is incubated at 30°C for 90 min, and then stored at −20°C. 2. To determine the incorporation of the labeled 35S-methionine, 2 µl of the mixture are removed from the reaction and added to 98 µl of 1 M NaOH/2% H2O2. After a short vortexing, the sample is incubated at 37°C for 10 min. Each 50 µl is dropped onto a small Whatman 3MM filter paper and airdried. To measure the incorporation of 35S, one of the filter papers is washed once with 10% TCA, three times with 5% TCA, two times with ether/ethanol (1:1), and, after drying, is then soaked in 5 ml scintillation mixture. To measure the

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total counts present in the reaction, the other filter paper without any washes is soaked in 5 ml scintillation mixture. The radioactivity counts are measured with a liquid scintillation counter. The following calculation is performed to determine percent incorporation: Percent incorporation = (cpm of washed filter / cpm of unwashed filter) × 3.3. Protein–Protein Interaction Techniques 3.3.1. Coimmunoprecipitation

1. HEK293T cells are transiently transfected with wild-type or mutant forms of V5-tagged HIF-1alpha and the wild-type HA-tagged VHL by the standard calcium phosphate method using 5 µg plasmid DNA per plate. 2. 16 h after the transfection cells are cultured further for 4 h under hypoxia or normoxia before harvesting. 3. The cell pellet is resuspended in 300 µl phosphobuffer and incubated rotating at 4°C for 20 min. 4. After centrifugation at maximum speed in a table-top centrifuge at 4°C for 15 min, the supernatant is transferred to a new cup and the protein content is measured. 5. Subsequently, 300 µg of total cellular protein are incubated with 2 µg antibody against V5-tag at 4°C for 1 h (see Note 1). 6. 100 µl of 30% slurry of protein G Sepharose beads (see Notes 2 and 3) in phosphobuffer are added, and samples are incubated overnight at 4°C under rotation. 7. After short centrifugation, the Sepharose pellets are washed five times with phosphobuffer without protease inhibitors and after the last wash the beads are drained completely using a syringe and a needle. 8. Coimmunoprecipitated proteins and total cell extracts (see Note 4) are mixed with 2× SDS loading buffer, boiled at 95°C for 5 min, and analyzed by SDS–PAGE and Western blot with anti-HA-tag antibodies following the standard protocol.

3.3.2. GST Pull-Down Assay

GST Pull-down assay represents a form of affinity purification, and it is very similar to coimmunoprecipitation (Subheading 3.3.1) except that a bait protein (see Note 5), purified as described in Subheading 3.2.1 is used instead of an antibody. Here we describe the pull-down of the VHL protein to a GST-fusion protein containing a part of the ODDD, which is subjected to hydroxylation by PHDs. Only hydroxylated HIF-1 can bind VHL and thus the amount of VHL pulled down represents a measure of PHD hydroxylase activity. In addition, pull-down assays have been used in a number of other applications where protein–protein interactions are important.

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1. 20 µg purified GST-HIF-1alpha-TADN, or GST (as negative control) “bait” proteins are mixed with 50,000 dpm of [35S]-labeled human VHL as a prey (see Note 6) in buffer 1 and incubated at 20°C for 2 h with gentle shaking in the presence of 100 µl of 80% slurry of glutathione-Sepharose beads. 2. The resin is shortly centrifuged in a table-top centrifuge (350 ´ g) and washed four times with cold buffer 2 to remove the unbound material followed each time by centrifugation. 3. The GST-HIF-1alpha-VHL complexes (see Note 4) are eluted with 10 mM reduced glutathione. 4. Eluted proteins are mixed with 50 µl of 2× SDS loading buffer, denaturated at 95°C for 10 min, and loaded onto a 15% SDS–PAGE (see Note 7). 5. After electrophoresis, the gel is stained with Coomassie Blue R250. 6. Thereafter, it is placed in between two sheets of Whatman paper, wrapped in foil, and dried with a gel-dryer at 70°C for 2 h. 7. For autoradiography, the dried gel is exposed to a Phosphoimager screen overnight, and thereafter, signals are scanned and can be quantified with the ImageQuant software. An example of the results produced is shown in Fig. 1.

Fig. 1. Inhibition of HIF-hydroxylase activity by CoCl2. (a) In vitro prolyl hydroxylase activity assay. The GST-HIF1a-TADN fusion protein or the GST protein was incubated with HepG2 cell extract, cofactors, and [5-14C]2-oxoglutarate in the presence of CoCl2 (10 mM). The radioactivity associated to 14C-succinate was determined. In each experiment, the basal HIF-TADN-dependent activity (control) was set to 100% after being normalized by subtracting the GST-associated activity. Values are means ± SEM of three independent culture experiments. Statistics, Student’s t-test for paired values: *P £ 0.05 vs. control. (b) GST pull-down assay. HepG2 cells were treated with or without CoCl2 (10 mM). Cell extracts were prepared and incubated with the GST-HIF1a-TADN fusion protein supplemented with cofactors. Glutathione-Sepharose beads and [35S]VHL were then added and the bound VHL was recovered, subjected to SDS–PAGE, and visualized by phosphoimaging. The input remains from directly loaded [35S]VHL. The two bands represent the 213 and 160 amino acid VHL translation products (105, 106).

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In addition to the GST-HIF VHL pull-down assay, PHD activity can be measured also in a hydroxylation assay with GST-HIF proteins or even peptides encompassing the region containing the prolines, which are subjected to hydroxylation. Thus, the activity of the hydroxylating PHDs can be calculated from the formation of succinate out of 2-oxoglutarate. 1. HepG2 cells are cultured for 24 h under normoxia, the medium is aspirated, and the cells are homogenized at 4°C in lysis buffer. 2. The homogenate is centrifuged at 1,000 × g for 10 min to remove cellular debris and nuclei. The supernatant is centrifuged at 3,000 × g for 10 min. Again, supernatant is centrifuged at 18,000 × g for 10 min, and the resulting pellet is resuspended in 40 mM Tris–HCl (pH 7.5). 3. Cellular extracts (300 mg/ml) are incubated at 37°C for 30 min in reaction buffer supplemented with 50,000 dpm [5-14C]2-oxoglutarate and 20 µg purified GST or GST-HIF1alpha-TADN protein. 4. Radioactivity associated with succinate is determined in a liquid scintillation counter (101) (see Note 8). The basal GST-dependent activity is subtracted from the GST-TADNdependent activity. Further, a number of assay modifications have been published (102), and they are based on the same principle as above or hydroxylase activities are determined by the amount of radioactive 4-hydroxyproline formed when wild-type, Pro402 → Ala, Pro564 → Ala, or double mutant HIF-1alpha-ODDD translated in the presence of L-[2,3,4,5-3H]proline are used as a substrate (103).

3.5. Acetylation 3.5.1. Acetyltransferase Assay

To estimate the activity of acetyltransferases, which may modify HIF-1alpha, it is advisable to perform an acetyltransferase assay. This N-terminal acetyltransferase assay is basically performed as described by (104) with slight modifications. 1. Pure His-tagged ARD1 protein purified by nickel affinity chromatography or a complex of in vitro translated NAT1 (N-terminal acetyltransferase 1) and His-tagged ARD1 are used. 2. For immunoprecipitation of the NAT1–ARD1 complex, 60 µl of NAT1 translation mixture, 20 µl of ARD1 translation mixture, 15 µg of anti-His antibody, 160 µl of RIPA buffer, and 30 µl of 80% protein-G Sepharose are incubated on ice for 16 h. 3. The complex bound to the beads is collected by centrifugation in a table-top centrifuge (350 ´ g) or by passive sedimentation and washed three times with RIPA buffer containing 1% Nonidet P-40 and once with RIPA buffer without Nonidet P-40.

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4. The beads with the NAT1–ARD1 complex are incubated at 30°C for 3 h in a 150 µl reaction containing 136 µl of 0.2 M K2HPO4 (pH 8), 4 µl of 4.5 mM [3H]acetyl-CoA, and 10 µl of 33 µM human adrenocorticotropic hormone with constant agitation. 5. After centrifugation, the supernatant is applied to 150 µl of SP Sephadex and incubated for 5 min with rotation. 6. The mixture is centrifuged shortly in a table-top centrifuge (350 ´ g), and the resin is washed three times with 1 ml of 0.5 M acetic acid and finally with 300 µl of methanol. 7. Radioactivity in the corticotropin-containing pellet is determined by scintillation counting. 3.5.2. In Vitro Acetylation Assay

1. GST-HIF-1alpha-ODDD protein (1 µg) purified from bacteria (Subheading 3.2.1) and immobilized on gluthatione sepharose beads is coincubated with His-tagged ARD1 (1 µg) purified by nickel affinity chromatography in acetylation buffer at 30°C for 1.5 h. 2. The beads with coimmunoprecipitated protein complex are collected by short centrifugation in a table-top centrifuge (350 ´ g) or by passive sedimentation, followed by four washes with RIPA buffer. After the last wash, the beads are drained completely using a syringe and a needle, mixed with 2× SDS loading buffer, denaturated at 95°C for 10 min, and loaded on a SDS–PAGE. 3. Whether or not GST-HIF-1alpha-ODDD has undergone acetylation by ARD1 is identified by immunoblotting with an anti-acetyl-lysine antibody.

3.6. Phosphorylation 3.6.1. Growth Factor Treatment/Inhibitor Studies

1. HepG2 cells, plated in 1.5 ml MEM in 6 cm Ø Petri-dishes, are maintained for 16–18 h in serum-free medium, and then pretreated with a protein kinase inhibitor diluted into culture medium to yield 10 µM end concentration for 30 min. 2. Cultures are either treated or not with agonist and exposed to normoxia or hypoxia for 4 h or 15 min depending on the protein levels to be detected. 3. The medium is aspirated, cells are washed twice with ice-cold 1× PBS, and scraped off the plate into 150 µl lysis buffer. 4. The cells are thoroughly destroyed by ultrasonication in short pulses on ice. 5. After centrifugation at 10,000 × g at 4°C for 20 min, the supernatant is collected into a new tube and the protein concentration is measured.

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Fig. 2. Inhibition of the IGF-1-mediated HIF-1alpha induction by the PI(3)-kinase inhibitor LY294002 and the MEK inhibitor U0126. Serum-starved HepG2 cells were pretreated with 10 µM LY294002 or 10 µM U0126 for 30 min and then treated either with or without 100 nM human IGF-1 (Sigma) and exposed to normoxia (16% O2) or hypoxia (8% O2) for 4 h. Acetic acid was used in controls at a final concentration of 100 nM to keep the pH constant. 100 µg of protein from HepG2 cell lysates were analyzed by Western Blotting with antibodies against HIF-1alpha (Novus Biological Transduction Lab, 1:2,000), or against phospho-ERK1/2 (cell signaling, 1:1,000) where HepG2 cells were stimulated for 15 min with IGF-1. Autoradiographic signals were detected by chemiluminescence (77).

6. 100 µg of protein from HepG2 cell lysates are analyzed by Western Blotting following the standard protocol. An example of the results produced is shown in Fig. 2. 3.6.2. Phosphorylation of GST-Fusion Proteins by Recombinant GSK3b In Vitro

1. 10 mM ATP is diluted with 3× assay buffer 1:40 to make 250 mM ATP. 2. [g-32P] ATP is diluted to 0.2 mCi/ml [g-32P] ATP with 250 mM ATP solution. 3. Enzyme is transferred from −80°C to ice, and after thawing, GSK-3b is diluted to the desired concentration with 1× assay buffer. 4. Wild-type or mutant GST-fusion proteins (20 mg) purified as described in Subheading 3.2 are incubated in kinase buffer in the presence of 50 mU active GSK-3b and 1 mCi [g-32P]ATP at 30°C for 30 min. 5. As a positive control a peptide being a GSK-3b substrate (YRRAAVPPSPSLSRHSSPHQSEDEEE) should be used in a separate reaction. 6. After finishing the reaction samples are loaded onto a 10% SDS gel, and after electrophoresis and blotting onto a polyvinyldiene difluoride membrane, phosphorylated proteins are visualized by phosphorimaging. 7. After autoradiography, the membrane can be used to detect the respective GST-fusion proteins with an antibody against GST. An example of results produced is shown in Fig. 3.

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Fig. 3. HIF-1alpha is phosphorylated by GSK-3. (a, b) The positive control peptide (CP), the GST and the GST-HIF-1a-TADN wild-type fusion proteins were incubated with 50 mU active GSK-3b and 1 µCi (32P-g ATP) for 30 min at 30°C. Afterwards the phosphorylated proteins were separated from unbound radioactivity by electrophoresis on a 10% SDS gel. Radioactive proteins were visualized by phosphoimaging. After autoradiography, the membrane was used to detect the respective GST-fusion proteins with an antibody against GST (86).

4. Notes 1. An optional step is preclearing – addition of sepharose beads to the protein mixture in order to reduce nonspecific binding of proteins to the uncoated sepharose. 2. Superparamagnetic beads can be used instead of sepharose beads. 3. It is possible that the Abs/bait protein are first immobilized on the beads and then added to the protein mixture (direct method), or as described, Abs/bait protein can be added directly to the protein mixture (indirect method). 4. Negative controls and positive controls are absolutely necessary to be prepared in order to eliminate “false” positive results as a result of unspecific binding to the beads and to prove the proper functioning of the method.

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5. Bait proteins can also be generated by linking an affinity tag to proteins. 6. As prey protein cell lysates, recombinant purified proteins, or in vitro transcription/translation reactions can be used. 7. For visualization of the protein–protein interaction, a SDS– PAGE followed by Western blot or Coomassie, or silver staining, and [35S] radioisotopic detection can be used. 8. HIF hydroxylation can also be determined by MALDI-TOF.

Acknowledgments Our studies were supported by grants from Fonds der Chemischen Industrie and Deutsche Krebshilfe 106929. References 1. Kewley RJ, Whitelaw ML, Chapman-Smith A (2004) The mammalian basic helix-loophelix/PAS family of transcriptional regulators. Int J Biochem Cell Biol 36:189–204 2. Kvietikova I, Wenger RH, Marti HH, Gassmann M (1995) The transcription factors ATF-1 and CREB-1 bind constitutively to the hypoxia-inducible factor-1 (HIF-1) DNA recognition site. Nucleic Acids Res 23:4542–4550 3. Wenger RH, Stiehl DP, Camenisch G (2005) Integration of oxygen signaling at the consensus HRE. Sci STKE 2005:re12. doi:re12 4. Semenza GL (2007) Life with oxygen. Science 318:62–64 5. Wang GL, Jiang BH, Rue EA, Semenza GL (1995) Hypoxia-inducible factor 1 is a basichelix-loop-helix-PAS heterodimer regulated by cellular O2 tension. Proc Natl Acad Sci USA 92:5510–5514 6. Kallio PJ, Okamoto K, O’Brien S, Carrero P, Makino Y, Tanaka H et al (1998) Signal transduction in hypoxic cells: inducible nuclear translocation and recruitment of the CBP/ p300 coactivator by the hypoxia-inducible factor-1alpha. EMBO J 17:6573–6586 7. Jiang BH, Zheng JZ, Leung SW, Roe R, Semenza GL (1997) Transactivation and inhibitory domains of hypoxia-inducible factor 1alpha. Modulation of transcriptional activity by oxygen tension. J Biol Chem 272:19253–19260 8. Pugh CW, O’Rourke JF, Nagao M, Gleadle JM, Ratcliffe PJ (1997) Activation of hypoxiainducible factor-1; definition of regulatory

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Dimova and Kietzmann similarity to hypoxia-inducible factor 1alpha regulates the VEGF expression and is potentially involved in lung and vascular development. Proc Natl Acad Sci USA 94:4273–4278 Flamme I, Frohlich T, von Reutern M, Kappel A, Damert A, Risau W (1997) HRF, a putative basic helix-loop-helix-PAS-domain transcription factor is closely related to hypoxia-inducible factor-1 alpha and developmentally expressed in blood vessels. Mech Dev 63:51–60 Hogenesch JB, Chan WK, Jackiw VH, Brown RC, Gu YZ, Pray GM et al (1997) Characterization of a subset of the basichelix-loop-helix-PAS superfamily that interacts with components of the dioxin signaling pathway. J Biol Chem 272:8581–8593 O’Rourke JF, Tian YM, Ratcliffe PJ, Pugh CW (1999) Oxygen-regulated and transactivating domains in endothelial PAS protein 1: comparison with hypoxia-inducible factor-1 alpha. J Biol Chem 274:2060–2071 Morita M, Ohneda O, Yamashita T, Takahashi S, Suzuki N, Nakajima O et al (2003) HLF/ HIF-2alpha is a key factor in retinopathy of prematurity in association with erythropoietin. EMBO J 22:1134–1146 Warnecke C, Zaborowska Z, Kurreck J, Erdmann VA, Frei U, Wiesener M et al (2004) Differentiating the functional role of hypoxia-inducible factor (HIF)-1alpha and HIF-2alpha (EPAS-1) by the use of RNA interference: erythropoietin is a HIF-2alpha target gene in Hep3B and Kelly cells. FASEB J 18:1462–1464 Scortegagna M, Ding K, Zhang Q, Oktay Y, Bennett MJ, Bennett M et al (2004) HIF2alpha regulates murine hematopoietic development in an erythropoietin-dependent manner. Blood 105(8):3133–3140 Wiesener MS, Turley H, Allen WE, Willam C, Eckardt KU, Talks KL et al (1998) Induction of endothelial PAS domain protein-1 by hypoxia: characterization and comparison with hypoxia-inducible factor-1alpha. Blood 92:2260–2268 Hu CJ, Wang LY, Chodosh LA, Keith B, Simon MC (2003) Differential roles of hypoxia-inducible factor 1alpha (HIF1alpha) and HIF-2alpha in hypoxic gene regulation. Mol Cell Biol 23:9361–9374 Raval RR, Lau KW, Tran MG, Sowter HM, Mandriota SJ, Li JL et al (2005) Contrasting properties of hypoxia-inducible factor 1 (HIF-1) and HIF-2 in von Hippel-Lindauassociated renal cell carcinoma. Mol Cell Biol 25:5675–5686

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Chapter 14 The Basic-Helix-Loop-Helix-Leucine Zipper Gene Mitf : Analysis of Alternative Promoter Choice and Splicing Kapil Bharti, Julien Debbache, Xin Wang, and Heinz Arnheiter Abstract The activity of transcription factors is often regulated by Post-translational modifications. A precondition for such modifications is the presence, in the corresponding mRNAs, of the exons that either directly encode the modifiable residues in question, or encode protein domains that influence their modification indirectly. The inclusion or exclusion of coding exons is regulated predominantly by alternative splicing but can also depend on promoter choice and polyadenylation site selection. Information about exon inclusion and exclusion, both qualitatively and quantitatively, is particularly important for experiments designed to mutate endogenous codons because such mutations can alter splicing patterns. Therefore, we here describe methods employed to quantitate exon inclusion and exclusion, using as example a mouse transcription factor gene, Mitf. Key words: Reverse transcriptase polymerase chain reaction, Real-time PCR, Serine phosphorylation, Knock-in allele

1. Introduction A prerequisite for Post-translational modifications is the incorporation of the specific exons coding for the modifiable residues. It is particularly important to determine the relative efficiency of exon inclusion or exclusion in cases where endogenous genes are mutated in codons that affect modifiable residues as such mutations can lead to the absence rather than the (intended) presence of the modifiable residue. It is for this reason that we here describe methods that can be employed to analyze and quantitate promoter and splice choices that determine the presence or absence of specific exons. Our description focuses on a model gene, Mitf. This gene encodes a basic helix-loop-helix–leucine zipper transcription factor whose major role in vertebrates is the regulation of the Paul J. Higgins (ed.), Transcription Factors: Methods and Protocols, Methods in Molecular Biology, vol. 647, DOI 10.1007/978-1-60761-738-9_14, © Springer Science+Business Media, LLC 2010

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development and function of melanin-bearing pigment cells (that is, melanocytes and retinal pigment epithelium or RPE cells) (1). Mutations in Mitf can lead to severe deficiencies in neural crestderived melanocytes which in mammals are not only associated with loss of coat pigmentation but also with deafness (2). In man, for instance, about 20% of congenital hearing deficiencies of the type “Waardenburg syndrome II” are associated with heterozygosity for mutations in MITF (3). In birds and rodents, homozygosity for Mitf mutations can also be associated with the “transdifferentiation” of retinal pigment epithelium cells into a retina-like tissue, a developmental aberration associated with small eyes (called microphthalmia, hence the name, Mitf = microphthalmia-associated transcription factor) (4). Nevertheless, although Mitf is expressed in many more cell types besides pigment cells, most of them do not display overt phenotypes when Mitf is mutated. Among those that are affected are mast cells, B cells, and osteoclasts. The latter cells show severe impairments when Mitf is mutated in such a way that the mutant protein, which forms obligatory homo- and heterodimers, acts in a dominant-negative manner. Mice with such mutations can have an osteopetrosis leading to premature death at weaning (5). Although encoded by a single gene, MITF is not a single protein but represents a family of isoforms generated by alternative promoter choice, alternative splicing, and a series of functionally relevant post-translational modifications (reviewed in ref. (2)). In fact, the 214,000 bp gene (mouse, human) has at least nine different promoters, at least six of which associated with

Fig. 1. Partial gene structure of Mitf, focusing on alternative promoters D and H and showing noncoding/coding parts of 5¢ exons, splice patterns and primer selection for quantitative determination of promoter choices. Note that isoform H utilizes an translational start codon in exon H, and isoform D a start codon in the 3¢ part of exon 1B, called exon 1B1b. For results obtained using these primers, see ref. (6).

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unique amino-terminal protein sequences. Some of these promoters show a precise tissue-specific and developmental regulation (ref. (6), see also Fig. 1). Moreover, modifications of splicing patterns have been associated with specific pigmentary phenotypes in mice (7). Lastly, serine phosphorylation, sumoylation, and acetylation have all been shown to affect MITF activity in vitro. For instance, sumoylation at lysine-182 and lysine-316 decreases MITF activity in a promoter-specific way that depends on the number of cognate promoter motifs (E-boxes) capable of interacting with MITF (8, 9). Moreover, it has been reported that MAPK-mediated phosphorylation at serine-73 increases the protein’s capacity to stimulate the promoter of tyrosinase, a melanocyte differentiation gene, and that double phosphorylation at serine-73 and serine-409 leads to increased MITF protein degradation (10, 11). Serine-73 is present in exon 2B (see Fig. 2), an

Fig. 2. Partial gene structure of mouse Mitf, focusing on exon 2A/2B and showing alternative splice products and primer choices for quantitative determination of exon inclusion. The top shows a partial gene structure for the region spanning exon 1B and 3. Note that two alternative promoters linked to either exon 1B or 1M and the common splice acceptor in exon 2A. Exon 2 is bipartite, with exon 2B either included (Mitf 2B+) or excluded (Mitf 2B−) from the mRNA. Primers overlapping the 2A/2B junction or the 2A/3 junction are shown, with reverse primers placed such that similar size products result for the two exon 2B splice versions. The schematic also shows the relative position of a serine-73-to-alanine codon mutation (S73A). This mutation favors exon 2B exclusion.

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exon that is normally absent in only 5–10% of Mitf mRNAs, but is absent in over 95% of mRNAs transcribed from a mutant Mitf allele characterized by a serine-73-to-alanine codon change (12). These observations suggest that the serine-73 codon is part of an exonic splice enhancer sequence that binds specific arginine/serine-rich proteins which are known to regulate mRNA splicing (for a recent review, see ref. (13)). This example highlights the importance of exonic sequences in determining splice choices and hence, ultimately, whether the protein can be Posttranslationally modified or not (see Fig. 3).

Fig. 3. Real-time PCR to quantify exon inclusion/exclusion. (a) Establishment of standard curves. Graded amounts of mouse Mitf standard cDNA were mixed with heterospecific cDNA, and real-time PCR was performed as indicated in the text. 2B+ primers correspond to primer pair a–c from Fig. 2, and 2B− primers correspond to primer pair b–d from Fig. 2. (b) Quantitation of exon 2B exclusion in RNA prepared from HEK293 cells transfected with the minigene as described in text and Fig. 2. Regular PCR followed by product identification by gel electrophoresis was done with a forward primer in exon 1M and a reverse primer in exon 3. This yields a product of 312 bp when exon 2B is included, and of 144 bp product when exon 2B is excluded. S73A corresponds to a minigene with a two base-pair change in codon 73 (AGC-to-GCC). Results are given as mean with bars indicating standard deviation from three measurements. (c) Similar quantitative assay performed with cDNA obtained from hearts of wild type mice or mice with the S73A codon change as indicated for (b). Note that the difference between wild type and mutant is more pronounced in vivo compared to minigene-transfected cells. wt wild type.

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2. Materials 2.1. Embryo Harvesting and Tissue Grinding

1. Pregnant mouse females, for instance C57BL/6J (Jackson Labs, Bar Harbor, ME). 2. RNaseZap (Ambion, Austin, TX), RNase inhibitor solution for cleaning surgical instruments. Store at room temperature. 3. Straight surgical scissors (27 mm for mouse dissection and 15 mm for removing embryos from the uterus, ROBOZ Scientific, Gaithersburg, MD). 4. Two pairs of microdissection tweezers (tips 0.05 × 0.01 mm) (ROBOZ Scientific, Gaithersburg, MD). 5. 10 cm and 3.5 cm Petri dishes (Becton Dickinson and Company, Franklin Lakes, NJ). 6. DEPC-treated Molecular Biology grade water (Quality Biologicals, Inc., Gaithersburg, MD). Store at room temperature. 7. 10× PBS (Quality Biologicals, Inc., Gaithersburg, MD). Mix 100 ml of 10× PBS with 900 ml of DEPC-treated water to make 1× PBS. Store at 4°C. 8. 1.5 ml microcentrifuge tubes (Denville Scientific, Metuchen, NJ). 9. Plastic tissue grinders (Bel-art Products, Pequannock, NJ).

2.2. Cell Culture and Transfection

1. Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 1% Penicillin/ Streptomycin (Gibco/BRL, Carlsbad, CA). Store all reagents at 4°C. 2. Solution of 0.05% trypsin and ethylenediamine tetraacetic acid (EDTA) (Gibco/BRL, Carlsbad, CA). Store at 4°C. 3. 10 and 6 cm tissue culture grade dishes (Becton Dickinson and Company, Franklin Lakes, NJ). 4. Tris/EDTA (TE): 10 mM Tris-Hcl, 1 mM EDTA, pH 7.4 (Quality Biologicals, Inc., Gaithersburg, MD). Store at room temperature. 5. Effectene Transfection Kit (Qiagen, Valencia, CA). Store at 4°C. 6. 15 ml centrifuge tubes (Denville Scientific, Metuchen, NJ).

2.3. RNA Extraction and cDNA Preparation

1. RNeasy Mini RNA Extraction Kit (Qiagen, Valencia, CA). Store at room temperature.

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2. Ethanol 200 proof (The Warner-Graham Company, Cockeysville, MD). Mix 70 ml of 200 proof ethanol with 30 ml of DEPC-treated water to make 70% ethanol. 3. RNase-free-DNase (3,000 Kunitz units/ml, Qiagen, Valencia, CA). 4. Nanodrop Spectrophotometer (Nanodrop Technologies, Wilmington, DE). 5. SuperScript First-Strand Synthesis System for RT-PCR (Invitrogen, Carlsbad, CA). Store at −20°C. 2.4. Polymerase Chain Reaction

1. Taq 2000 Polymerase (5 units/µl) (Stratagene, La Jolla, CA). Store at −20°C. 2. 2 mM dNTPs (MBI Fermentas, Glen Burnie, MD). Store at −20°C. 3. Primers, final concentration 10 pmole/µl (Genelink, Hawthorne, NY). Store at −20°C. 4. PCR tubes 0.2 ml (Denville Scientific, Metuchen, NJ). 5. Molecular biology grade water (Quality Biologicals, Inc., Gaithersburg, MD). Store at room temperature. 6. Dyad DNA Engine Themocycler (MJ mini, now Bio-Rad, Hercules, CA). 7. 5× DNA loading dye (Teknova, Hollister, CA).

2.5. Agarose Gel

1. Ultra pure agarose (Invitrogen, Carlsbad, CA). Store at room temperature. 2. 50× TAE (Quality Biologicals, Inc., Gaithersburg, MD). 50× TAE is diluted to 1× by mixing 20 ml of 50× TAE in 980 ml of deionized water. 3. Ethidium bromide 10 mg/ml (Sigma, St. Louis, MO). 4. Sub-Cell GT agarose gel electrophoresis unit (Bio-Rad, Hercules, CA). 5. Pharmacia EPS 250/200 power supply (GE Healthcare, Pharmacia, Uppsala, Sweden). 6. Molecular weight marker, DNA ladder mix (MBI Fermentas, Glen Burnie, MD). Store at −20°C.

2.6. Real-Time PCR Analysis

1. Power SYBR Green Mix (2×) (Applied Biosystems, Foster City, CA). Store at 4°C. 2. 96-well optical reaction plate (Applied Biosystems, Foster City, CA). 3. ABI Prism 7000 Sequence Detection System (Applied Biosystems, Foster City, CA).

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3. Methods 3.1. Sample Preparation for Isoform Analysis 3.1.1. Harvesting Embryonic Tissue for Detection of Alternate Promoter Isoforms of Mitf at Different Developmental Stages. Example Eye Tissue

1. Pregnant C57BL/6J females: To calculate the developmental time point for harvesting embryos, the morning that the vaginal plug is first detected is considered embryonic day (E) 0.5. For instance, E11.5 embryos are collected 11 days following the day of plug detection (E0.5). For detailed information about embryo collection, see ref. (14). 2. Harvesting tissue for RNA extraction requires precautions to avoid RNase contamination. Before starting to dissect the pregnant female, treat all instruments with RNase inhibitor solutions (RNaseZap), and prepare the solutions needed for this procedure with DEPC-treated molecular biology grade water. A separate set of tweezers, which has not been used during mouse dissection, is used for the dissection of the eye tissue. Two sets of appropriately labeled 1.5 ml microcentrifuge tubes are stored in dry ice. 3. Euthanize the pregnant females according to methods approved by your Animal Care and Use Committee (for instance CO2). Harvest embryos using 27 mm scissors and tweezers and place the embryos in cold 1× PBS in a 10-cm dish. Remove the placenta and the chorionic membranes from each embryo, one at a time, using a pair of 15 mm scissors, and transfer the embryos to cold 1× PBS in a second 10 cm dish. 4. Using the clean set of tweezers, carefully remove both eyes from an embryo under a dissection scope and transfer the eyes to cold 1× PBS in a 3.5 cm dish. While holding the eyeball with one tweezer, use the other tweezer to enlarge the optic pit hole at the back of the eye. Push the retina/lens through the optic pit hole by squeezing/pushing at the anterior part of the eye. Collect the separated RPE/mesenchyme and retina/lens tissue into separate 1.5 ml microcentrifuge tubes stored on dry ice (see Note 1). Freeze these tubes at −80°C until further processing. 5. For other tissues, proceed accordingly.

3.1.2. Harvesting Transfected Cells for Assaying Alternative Splicing of Different Mitf Exons

1. Human embryonic kidney cells (HEK293) are cultured in DMEM medium with 10% FBS and 1% Penicillin/ Streptomycin at 37°C in an incubator with 6% CO2/air. Passage the cells as they approach 70% confluency, using 0.05% trypsin/EDTA, and replate at a dilution of 1:5 (see Note 2). 2. The methods described here are applicable to analyzing the relative and absolute levels of different splice isoforms of

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endogenous Mitf mRNAs. Because one of the goals is to analyze the sequence requirements for regular and alternative splicing, we use wild type and various mutated minigene constructs that are transfected into cultured cells. Generally, these minigene constructs contain a part of the gene where the exon to be tested for alternative splicing is flanked, at least, by the two neighboring introns and adjacent exons. For instance, to analyze alternative splicing of exon 2B of Mitf, the reporter construct contains part of the gene from exon 1M to exon 3, including introns 1 and 2 (see Fig. 2). For expression, these minigenes are placed under the control of a CMV promoter and contain a 3¢ untranslated region and polyadenylation signal. The strength of alternate splicing is analyzed by comparison with constructs where splice junctions or splice enhancer and silencer sequences are mutated. 3. For transfection, plate one million of freshly trypsinized cells in a 6 cm dish and transfect them the following morning using the Effectene Transfection Kit (Qiagen). Dilute 1 µg of the minigene construct dissolved in TE into 150 µl of DNA condensation buffer (Buffer EC) in a 1.5 ml microcentrifuge tube. Add 8 µl of enhancer solution and vortex the tubes for 1 s (see Note 3). 4. Incubate at room temperature for 2–5 min, then briefly spin the tubes to collect drops from the tube top. 5. Add 25 µl of effectene reagent to the tubes and vortex for 10 s. Incubate tubes at room temperature for 10 min. 6. While the DNA-transfection complexes are forming, aspirate the medium from the cells and gently wash cells with 4 ml 1× PBS. Add 4 ml fresh DMEM medium to the cells and return the dishes to the incubator. 7. Add 1 ml DMEM medium to the tube containing the DNAtransfection complex and mix gently by pipetting up and down. Add the transfection mix to the cells and gently swirl the dish to ensure uniform distribution of the DNAtransfection mix. Return the dishes to the incubator. 8. After 16 h of incubation, collect cells for RNA extraction. Aspirate the medium from the dish and wash cells with 4 ml 1× PBS. Trypsinize cells with 1 ml 0.05% trypsin/EDTA. Add 5 ml of fresh medium to the cells and transfer detached cells to a 15 ml centrifuge tube and pellet by centrifugation at 300 × g for 5 min. Aspirate the medium from the cell pellet without disturbing the pellet. Wash the cells with 1× PBS and spin the tubes again at 300 × g for 5 min. Aspirate the supernatant fluid and flash freeze the tubes in dry ice. Store tubes at −80°C until further processing.

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1. To grind embryonic tissue, use plastic tissue grinders fitting the conical bottom of a 1.5 ml microcentrifuge tube. Before grinding, clean the grinders with RNase inhibitor solution (RNaseZap) and store at −20°C to keep them cold. Add 50 µl of RLT buffer from the Kit (add 10 µl of b-mercaptoethanol to 1 ml RLT buffer before use) to the frozen tissue and grind it on dry ice using the prechilled plastic grinders. After grinding, add an additional 300 µl of RLT buffer to the tissue and freeze on dry ice until other tubes are processed. Once all the tubes are processed, thaw them on ice and process for RNA extraction starting at step 3. 2. To lyse frozen tissue culture cells, add 350 µl RLT buffer to the cell pellet immediately after taking it from the freezer. Vortex the tube to ensure a homogenous suspension of the cells and proceed to step 3. 3. Pipette the tissue lysate from step 1 or the cell suspension from step 2 onto a QIAshredder column placed in a 2 ml collection tube. Centrifuge tubes for 2 min at maximum speed to lyse the cells. 4. Add 350 µl of 70% ethanol to the homogenized cell lysate and mix thoroughly by pipetting up and down. Do not vortex or centrifuge the samples at this step. 5. Pipette all 700 µl of the sample mixture, including any precipitates that may have formed in the previous step, to an RNeasy mini spin column placed in a 2 ml collection tube. Centrifuge the samples for 30 s at 9,300 × g. 6. Discard the flow-through. Add 500 µl of buffer RW1 from the kit to the RNeasy column and spin the columns for 30 s at 9,300 × g. 7. Discard the flow-through along with the collection tube and transfer the column to a new collection tube. 8. Add RNase-free-DNase, in a mixture of 75 µl of buffer RDD from the kit and 5 µl of DNase (15 Kunitz units totally), onto each column. Make sure that the DNase solution covers the entire surface of the membrane and incubate columns at room temperature for 30 min. 9. Add another 500 µl of buffer RW1 to the column and spin the columns for 30 s at 9,300 × g. 10. Transfer RNeasy column to a new collection tube, add 500 µl of buffer RPE from the kit to the column, and spin for 30 s at 9,300 × g. 11. Add another 500 µl of buffer RPE onto each column and spin for 2 min at 9,300 × g to dry the column.

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12. Transfer the column to a 1.5 ml microcentrifuge tube and add 50 µl of 65°C RNase-free water onto the membrane. Incubate at room temperature for 5 min and spin the columns for 30 s at 9,300 × g. 13. The eluate contains pure RNA. Measure concentration using a nanodrop spectrophotometer using 1.3 µl of the undiluted RNA solution. 3.2.2. cDNA Preparation Using Superscript First-Strand Synthesis System for RT-PCR

1. Mix 1 µg of RNA with 1 µl of 10 mM dNTP mix, and 2 µl of random hexamers (50 ng/µl) in a 0.2 ml PCR tube. Adjust the final volume of the mixture to 10 µl with DEPC-treated water (see Note 4). 2. Incubate tubes at 65°C for 5 min and then transfer them to ice for 2–3 min. 3. While the samples are cooling on ice, prepare a second reaction mix containing 2 µl of 10× RT buffer, 4 µl of 25 mM MgCl2, 2 µl of 0.1 M DTT, 1 µl of RNaseOUT recombinant (Ribonuclease inhibitor) and 1 µl of Superscript II Reverse Transcriptase (50 units). Add 10 µl of the reaction mixture to each RNA/primer mixture. Mix the solutions gently and spin the tubes briefly to collect all of the solution at the bottom. 4. Incubate the tubes for 2 min at 25°C. Transfer the tubes to 42°C for 1 h. 5. Terminate the reaction by incubating tubes at 70°C for 15 min and then place tubes on ice. 6. Add 1 µl of RNase H to each tube and incubate at 37°C for 20 min. 7. Dilute the cDNA 1:5 in water. The cDNA is now ready for PCR. 8. Before using this cDNA for real-time analysis, check the quality of each sample by PCR amplification of an unrelated cDNA.

3.2.3. Setting Up a PCR

1. For a 20 µl PCR reaction, mix 2 µl of 10× reaction buffer, 2 µl of 2 mM dNTPs, 2 µl of 10 pmole/µl of each forward and reverse primers, 1 µl of the cDNA template, 11.75 µl of molecular biology grade water, and 1 unit (0.25 µl) of Taq Polymerase in a 0.2 ml PCR tube. For primer choice, see Figs. 1 and 2 and ref. (6). 2. Place the tubes in the thermocycler block and set up a cycling program as follows: 92°C, 2 min (step 1), 92°C, 20 s; 55°C, 30 s; 72°C, 1 min (step 2, repeat for 29 cycles), 72°C, 7 min (last step). Start the cycling program. The program is designed for DNA Engine DYAD thermocycler from MJ mini (see Note 5).

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3. Once the PCR is finished, add 5 µl of 5× DNA gel loading buffer to each tube. The samples are now ready to load on an agarose gel. 3.2.4. Agarose Gel Electrophoresis

1. The agarose gel set up is described for Sub-Cell GT agarose gel electrophoresis unit from Bio-Rad. 2. To prepare a 1.5% agarose gel for a chamber that holds 150 ml of gel solution, weigh 2.25 g of agarose in a 250 ml Erlyenmeyer flask and add 150 ml of 1× TAE. Heat the solution with intermittent shaking until agarose dissolves completely. Let the gel solution cool to about 50°C. Add ethidium bromide at a final concentration of 1 µg/ml. Pour the gel solution into the gel tray with stoppers on each side. Add the combs to the tray and let the gel solidify. 3. Remove the combs and the stopper from the solidified gel and submerge the gel into 1× TAE running buffer in the gel electrophoresis unit. 4. Load 10 µl total of sample and 1× loading buffer per well. 5. In a corner well, load 3 µl of DNA ladder mix as a molecular weight marker. 6. Run the gel at constant 75 V. Stop the gel when the dye front has reached 2/3rd of the total running distance. Take a picture of the gel, using a UV-gel documentation unit.

3.3. Quantitative Real-Time PCR 3.3.1. Primer Design for Alternative Promoter Isoforms 3.3.2. Primer Design for Alternatively Spliced Exons

To detect alternative promoter choice for the Mitf gene in different tissues at various developmental stages, forward primers specific for the alternative exons are used in combination with a reverse primer that is common to all isoforms. Forward primers are designed in a way that the PCR products amplified for the different isoforms are of similar size. For details, see ref. (6). To detect alternatively spliced exons, forward primers are unique to the exon–exon junctions of the differently spliced products. For the situation where the alternative exon is spliced-in, the forward primer is positioned at the exon junction in a way that part of the primer hybridizes with the 3¢ end of the upstream exon, and part with the 5¢ end of the downstream exon. For the situation where the alternative exon is spliced-out, forward primers are placed accordingly in a way that they span the expected junction formed by the absence of the alternative exon. Hence, the 5¢ parts of both types of forward primers are identical, but because their 3¢ parts are different, cross-priming is not usually a problem. For reverse primers, sequences in the downstream common exon are chosen at locations that assure a similar length of the two expected products (Fig. 2).

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3.3.3. Generation of Standard Curve and Real-Time Analysis of Alternative Promoter Isoforms

1. Amplify standard cDNA products corresponding to each isoform, using primer pairs as described in Subheading 3.3.1, step 1. Using amplified DNA allows for the generation of linear standard DNAs of a size similar to the cDNAs present in the test samples and is to be preferred over the use of plasmids for standard curve purposes. 2. Quantify the amounts of standard DNA products both spectrophotometrically and by agarose gel electrophoresis (see below). 3. Dilute appropriate concentrations of the standard DNA, for instance in steps of tenfold dilutions, into a cDNA source that does not contain the isoforms to be tested. For instance, in the case of Mitf-promoter isoforms, cDNA from hearts of Mitf mi-rw/mi-rw animals was used (6). These mutant mice lack the exon corresponding to the common reverse primer (exon 1B) but provide a complexity of cDNAs similar to that expected in the test sample. This is critical as primer efficiency differs depending on the amplification environment. In the absence of appropriate mutants, use heterospecific cDNA and species-specific primers. 4. Perform real-time PCR on these samples using a set of primers that represent a nested set to the original pair used to amplify the standard DNAs (for details, see ref. (6). An ABI Prism 7000 real-time PCR machine is used for the real-time analysis. 5. A log of the amount of standard DNA (weight/reaction) is plotted against the threshold cycle (cT) values obtained for each concentration. A best-fit regression plot is drawn for each standard curve. Using the average molecular weight of a basepair as 660 Daltons, the weight can be converted into the number of DNA molecules. 6. Generate standard curves for each isoform, and separately for each repeat assay performed with the test samples. 7. Perform the real-time PCR of the test samples (for instance, cDNA obtained from E11.5 RPE and retina), by using the same set of nested primers as used in step 4. Normalize each sample by quantifying an unrelated cDNA, again using realtime PCR (see Note 6). The absolute amount of cDNA molecules in test samples is determined by using the appropriate previously determined isoform-specific standard curves and linear regression plots.

3.3.4. Generation of Standard Curve and Real-Time Analysis of Alternatively Spliced Exons

1. The principles of standard curve determination are according to the methods described above for promoter choice determination (see Note 7). 2. Standard templates of mouse cDNA are diluted in a cDNA mixture from a human cell line (see Notes 2 and 7).

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The primers used for this analysis are made specific to mouse sequences so that human Mitf cDNA is not amplified in this mixture of mouse standard cDNAs and human cell line cDNA. An example of a standard curve determination is shown in Fig. 3A. Note that inappropriate cross-priming (for instance, 2B+ primers on 2B− cDNA) for cDNA concentrations above 1 pg/reaction gives results that are 1–4 logs below those obtained for appropriate priming (for instance 2B+ primers on 2B+ cDNA). 3. Using the above methods of standard curve determination, test samples are subjected to real-time PCR as mentioned. Use quantification of an unrelated cDNA for normalization of the test samples (see Note 6). Results can be expressed in absolute numbers of cDNA molecules, or, as shown in Fig. 3B and 3C, as relative amounts of exon 2B-lacking cDNAs compared to the sum of exon 2B-lacking and exon 2B-containing cDNAs.

4. Notes 1. If embryos need to be genotyped, separate each embryo into an individual tube and remove a portion from each embryo for PCR or other appropriate genotyping reaction. 2. Instead of using heterospecific cells to prepare cDNA to be used during standard curve determination, one may also use cell types in which the gene of interest is not expressed (or expressed below threshold levels for the chosen PCR conditions). 3. The method of transfection has to be adapted to the type of cell line used. 4. The amount of RNA used for the RT reaction can be lower but each test sample should contain the same amount of total RNA. 5. The annealing temperature may vary with the efficiency of the primer sets, and the number of cycles for step 2 may vary depending on the abundance of the cDNA of interest in the sample. An optimum number of cycles is considered to be that which gives amplification in the linear range. 6. For normalization, use an unrelated cDNA whose threshold cycle time is similar to that of the test cDNA. For the analysis of exon 2B splice variants, for instance, we used USF, a cDNA whose relative abundance is similar to that of Mitf exon 2B− cDNA in wild type, and does not change significantly between samples obtained from different genotypes.

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7. If one wants to avoid the use of nested primer set-PCR to generate standard DNAs from appropriate plasmids, one can use the respective plasmids directly as standard cDNAs. In this case, only one primer set is used both for the generation of the standard curve and for the analysis of the test samples.

Acknowledgments This work was supported by the Intramural Research Program of the National Institutes of Health, NINDS. References 1. Hodgkinson CA, Moore KJ, Nakayama A, Steingrimsson E, Copeland NG, Jenkins NA, Arnheiter H (1993) Mutations at the mouse microphthalmia locus are associated with defects in a gene encoding a novel basic-helixloop-helix-zipper protein. Cell 74:395–404 2. Arnheiter H, Hou L, Nguyen MTT, Bismuth K, Csermely T, Murakami H, Skuntz S, Liu W, Bharti K (2006) Mitf – A matter of life and death for the developing melanocyte. In: Hearing V, Leong SPL (eds) From melanocytes to malignant melanoma. Humana, Totowa, NJ 3. Tassabehji M, Newton VE, Read AP (1994) Waardenburg syndrome type 2 caused by mutations in the human microphthalmia (MITF) gene. Nat Genet 8:251–255 4. Bharti K, Nguyen MT, Skuntz S, Bertuzzi S, Arnheiter H (2006) The other pigment cell: specification and development of the pigmented epithelium of the vertebrate eye. Pigment Cell Res 19:380–394 5. Steingrimsson E, Copeland NG, Jenkins NA (2004) Melanocytes and the microphthalmia transcription factor network. Annu Rev Genet 38:365–411 6. Bharti K, Liu W, Csermely T, Bertuzzi S, Arnheiter H (2008) Alternative promoter use in eye development: the complex role and regulation of the transcription factor MITF. Development 135:1169–1178 7. Hallsson JH, Favor J, Hodgkinson C, Glaser T, Lamoreux ML, Magnusdottir R, Gunnarsson GJ, Sweet HO, Copeland NG, Jenkins NA, Steingrimsson E (2000) Genomic,

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transcriptional and mutational analysis of the mouse microphthalmia locus. Genetics 155:291–300 Murakami H, Arnheiter H (2005) Sumoylation modulates transcriptional activity of MITF in a promoter-specific manner. Pigment Cell Res 18:265–277 Miller AJ, Levy C, Davis IJ, Razin E, Fisher DE (2005) Sumoylation of MITF and its related family members TFE3 and TFEB. J Biol Chem 280:146–155 Hemesath TJ, Price ER, Takemoto C, Badalian T, Fisher DE (1998) MAP kinase links the transcription factor Microphthalmia to c-Kit signalling in melanocytes. Nature 391:298–301 Wu M, Hemesath TJ, Takemoto CM, Horstmann MA, Wells AG, Price ER, Fisher DZ, Fisher DE (2000) c-Kit triggers dual phosphorylations, which couple activation and degradation of the essential melanocyte factor Mi. Genes Dev 14:301–312 Bismuth K, Skuntz S, Hallsson JH, Pak E, Dutra AS, Steingrimsson E, Arnheiter H (2008) An unstable targeted allele of the mouse Mitf gene with a high somatic and germline reversion rate. Genetics 178:259–272 Lin S, Fu XD (2007) SR proteins and related factors in alternative splicing. Adv Exp Med Biol 623:107–122 Nagy A, Gertenstein M, Vintersten K, Behringer R (2003) Manipulating the mouse embryo. A laboratory manual, 3rd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY

Chapter 15 Phosphorylation Control of Nuclear Receptors Sébastien Lalevée, Christine Ferry, and Cécile Rochette-Egly Abstract Most transcription factors including nuclear receptors (NRs) act as sensors of the extracellular and intracellular compartments. As such, NRs serve as integrating platforms for a variety of stimuli and are targets for Post-translational modifications such as phosphorylations. During the last decade, knowledge of NRs phosphorylation advanced considerably because of the emergence of new technologies. Indeed, the development of a wide range of phosphorylation site databases, high accuracy mass spectrometry, and phospho-specific antibodies allowed the identification of multiple novel phosphorylation sites in NRs. New and improved methods also emerge to connect these data with the downstream consequences of phosphorylation on NRs structure (computational prediction, NMR), intracellular localization (FRAP), interaction with coregulators (proteomics, FRET, FLIM), and affinity for DNA (ChIP, ChIP-seq, FRAP). In the future, such integrated strategies should provide data with a treasure-trove of information about the integration of numerous signaling events by NRs. Key words: Phosphorylation, Nuclear receptors, Phosphoproteomics, Mass spectrometry, Phosphorylation sites databases, Phosphorylation site-specific antibodies, FRAP ChIP

1. Introduction Protein phosphorylation is one of the most relevant and ubiquitous Post-translational modifications. It is an integral part of cell signaling and is involved in virtually all-eukaryotic cellular processes. It has been estimated that 30% of cellular proteins are phosphorylated at a given time, representing the phosphoproteome, and that over 100,000 potential sites of phosphorylation exist in the human proteome (1). Phosphorylation is a rapid and reversible modification that critically regulates most cellular events through altering protein structure, protein–protein interactions, protein’s activity, localization, and stability. Moreover, aberrant phosphorylation has been linked to a variety of disease states. Thus, the elucidation of protein phosphorylation is of great value Paul J. Higgins (ed.), Transcription Factors: Methods and Protocols, Methods in Molecular Biology, vol. 647, DOI 10.1007/978-1-60761-738-9_15, © Springer Science+Business Media, LLC 2010

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to understand signaling mechanisms and cellular networks in most biological systems and to reveal potential drug targets. Among the phosphoproteome, there are nuclear receptors (NRs), which are members of a large superfamily of evolutionarily related DNA-binding transcription factors (2, 3). In humans, 48 members of the family have been identified, 24 being liganddependent receptors (4). They include the nuclear receptors for steroid hormones such as the estrogen receptor (ER), the androgen receptor (AR), the progesterone receptor (PR), the glucocorticoid receptor (GR), and the mineralocorticoid receptor (MR) as well as for nonsteroidal ligands such as the vitamin D receptor (VDR), the thyroid hormone receptors (TRs), the retinoid receptors (RARs and RXRs), and the peroxysome proliferator activated receptors (PPARs). NRs share a well-defined organization, consisting mainly of a central DNA-binding domain (DBD) linked to a C-terminal Ligand-Binding domain (LBD) and an N-terminal domain (NTD), each domain containing phosphosites. Both the DBD and the LBD are highly folded with structures, which have been determined by nuclear magnetic resonance (NMR) and crystallographic studies. The phosphorylation sites identified in these domains are located in flexible regions such as loops, which are more accessible for molecular recognition and modifications (5, 6). In contrast, the NTD is natively unstructured (7, 8) and therefore contains the majority of the phosphorylation sites identified to date (9–12). In this chapter, we present an overview on how NR phosphorylation sites can be identified and how the consequences of phosphorylation on NRs activity can be analyzed. As most phosphorylation studies have been performed with ligand-dependent NRs, we will focus essentially on these receptors.

2. Identification of Nuclear Receptors Phosphorylation Sites

The analysis of NRs phosphorylation has been revealed as a challenging task due not only to its highly dynamic nature but also to the low ratio of phosphorylated versus nonphosphorylated NRs found in vivo. Early studies of NRs phosphorylation used radiolabeling, immunoprecipitation, 2D-PAGE, phosphoamino acid mapping, phosphopeptide mapping, protein sequencing, and sitedirected mutagenesis to identify candidate sites. Such classical approaches are clearly depicted and reviewed in details in (13–16). These classical approaches have resulted in the identification of multiple phosphorylation sites in steroid nuclear receptors (ER, PR, GR, AR) as well as in nonsteroid receptors (RARs, RXR, PPAR, etc.), most of them being located in the N-terminal domain (9–12). However, though informative, they are technically limited as they require radioactive material and large amounts of recombinant

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NRs overexpressed in cultured cells or of bacterially expressed NRs purified and phosphorylated in vitro with different kinases. Therefore, they are not efficient for identifying phosphorylation of endogenous NRs in response to their cognate ligand or to signaling pathways. Over the past 5 years, the emergence of new phosphoproteomic tools such as mass spectrometry-based analytical methods has exploded the field, and many excellent reviews have been written on the subject (17, 18). Briefly, large scale and high throughput analysis of the phosphoproteome by high performance tandem mass spectrometry (MS/MS) combined with methods for enrichment of samples at the phosphoprotein (immunoprecipitation with phosphospecific antibodies) or at the phosphopeptide levels (immobilized metal affinity (IMAC) or titanium dioxide chromatography (19)) and for fractionation of the enriched samples (nanoHPLC or capillary electrophoresis (17, 20)), allowed unambiguously to obtain large-scale phosphorylation data sets. A limitation of MS-based techniques is the requirement of large amounts of material due to the low stoichiometry of protein phosphorylation. However, when coupled to stable-isotope labeling, it presents the advantage of being a fundamental tool for quantifying changes in phosphopeptide abundance (1, 18, 21). Nevertheless, new and improved methods to conduct unbiased analysis of protein phosphorylation and to detect phosphorylated residues emerge every month. An updated summary of the locations and proposed functions of experimentally verified phosphorylation sites can be found at several phosphorylation site databases such as http://www. phosphosite.org and http://phospho.elm.eu.org. Most of them have been incorporated in the Swiss-prot database (http://ca. expasy.org/sprot) (22). In addition, a wide range of computational servers have been developed for prediction of phosphorylation sites for some kinase families and are available on Internet: Scansite (http://scansite.mit.edu/motifscan_seq.phtml) (23), Predikin (http://predikin.biosci.uq.edu.au) (24, 25), NetPhosK (http://www.cbs.dtu.dk/services/NetPhosK), Pred Phospho (http://pred.ngri.re.kr/PredPhospho.htm), and KinasePhos (http://kinasephos.mbc.nctu.edu.tw). More recently, developments in large-scale and high confidence quantitative MS-based phosphoproteomics allowed to extract thousands novel phosphorylation sites as well as novel motifs for specific kinases (21, 26). Therefore, new phosphorylation site databases such as Phosida (http://www.phosida.com) (27, 28) have been developed and can match kinase motifs to thousands of phosphosites. They also integrate structural and evolutionary information on each phosphosite. MS/MS analysis coupled with optimized database search strategies has allowed the discovery of novel phosphorylation sites with low abundance in ER (29), PR (30), and AR (31).

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However, it must be kept in mind that, before considered valid, the predicted phosphorylation sites must be experimentally verified using mutation of phosphorylated residues and/or phosphorylation site-specific antibodies (29, 32–35). Such antibodies provide promising tools to evaluate in immunoprecipitation and/ or immunoblotting experiments the phosphorylation profile of individual sites in response to a particular stimulus or during disease processes (33, 36, 37) and to reveal intricate interplays between the different phosphorylation sites within an NR (33).

3. Identification of Protein Kinases Involved in NRs Phosphorylation

4. Dynamics of NRs Phosphorylation

The kinase family responsible for the phosphorylation of a serine, threonine, or tyrosine residue can be predicted by analysis of the sequence containing the phosphosite, using the prediction servers mentioned above. However, several individual NR phosphorylation sites can be potentially modified by multiple kinases. Indeed, serine or threonine residues followed by a proline match a consensus motif recognized by either mitogen-activated protein kinases (MAPKs) or cyclin-dependent kinases (CDKs). Moreover, both kinases families are large and include several members that respond to a variety of different stimuli. As an example, in vivo, phosphorylation of ERa at serine 118 (38, 39), and of RARa at serine 77 (40) can involve either cyclinH/cdk7 in response to the ligand or p42/p44 MAPKs in some cancer cells independently of the ligand. In early in vivo studies, the search for the kinases involved in NRs phosphorylation was performed using pharmacological inhibitors that target specifically the different proline-directed kinases (33, 37, 39, 40). However, experiments with such inhibitors are sometimes controversial and may lack specificity. Today the development of new tools such as small interfering RNAs (41) coupled to kinase assays and microarrays improved significantly the identification of the kinases involved in NRs phosphorylation (33). High throughput screening of the kinases using siRNAs banks targeting the kinome can also be performed (Flexiplate siRNA gene family lists provided by Qiagen, for example). Such an approach requires automated cell transfection coupled to fluorescence microscopy analysis of NRs using phosphospecific antibodies.

Phosphorylation is a dynamic and cell-specific process. Therefore, equally challenging as the identification of the phosphosites is the comparison of phosphorylation profiles in response to the

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ligand or to specific signaling pathways. When combined with stable-isotope-labeling by amino acid in cell culture (SILAC) or stable-isotope-tagged amine-reactive reagents (iTRAQ), MS can follow the dynamics of phosphorylation (1, 18, 21). Albeit such a strategy provided valuable information concerning the regulation of phosphorylation sites, only the most abundant sites are generally monitored, due to loss of the phosphate groups during collision-induced dissociation (CID). More efficient fragmentation techniques such as electron capture dissociation (ECD) or electron transfer dissociation (ECD) should represent improvement to assign more precisely phosphosites in low abundance peptides with multiple charges (17). Now experiments such as flow cytometry can be designed to analyze signaling networks in single cells following stimulation in complex samples containing multiple cell types. This technique can analyze ten thousands fixed cells per second and measure up to 19 fluorescent parameters simultaneously, using antibodies to cell type specific surface markers and to phosphorylation sitesspecific antibodies (21, 42). It provided a dynamic picture of STATs phosphorylation in acute myeloid leukemia cells in response to different treatments. Due to the increasing number of NR phosphospecific antibodies available, flow cytometry should indicate how NRs respond to their cognate ligand and/or to different signaling pathways.

5. Phosphorylation and NRs Subcellular Localization

In general, nuclear receptors are located in the nucleus, but some of them such as AR, GR, and MR are cytoplasmic and undergo a ligand-induced nuclear import. These NRs also share the property of undergoing cycles of nucleocytoplasmic shuttling. Finally, NRs not only move between nuclear and cytosolic compartments but also within the nucleus between transcriptionally active or inactive clusters. A number of studies provided evidence that these movements would be controlled by phosphorylation. Immunofluorescence experiments performed with GFPtagged NRs (either WT or mutated at the phosphorylation sites) indicated that phosphorylation of specific sites enhances nuclear localization while phosphorylation of others increases nuclear export (11, 43, 44). However, forcing the localization of NRs to the nucleus or the cytoplasm upon fusion to a nuclear localization signal or a nuclear export signal, followed by analysis with phosphosite-specific antibodies, revealed that phosphorylation of certain sites would be compartment sensitive, whereas phosphorylation of other ones is not (45). Such approaches coupled to the use of kinase pharmacological inhibitors or siRNAs should establish

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correlations between NRs localization and pathologies characterized by aberrant kinase activities (46). During the last years, Fluorescence Recovery After Photobleaching (FRAP) improved considerably and allowed the researcher to visualize the dynamic behavior of NRs in a live-cell scenario. It has been used to examine nucleocytoplasmic shuttling using GFP-tagged AR or MR (47, 48). FRAP also proved to be a strong approach to study the intranuclear dynamics of NRs (49–51). Similar approaches using phospho-mutants should confirm the role of phosphorylation in these movements (52).

6. NRs Phosphorylation and Transcription Regulation

7. How Phosphorylation Regulates NR Transcriptional Activity

Initial studies of the role of NR phosphorylation in the regulation of NR-dependent transcription relied on transient transfection assays using receptors with alanine substitutions for the phosphorylation sites and reporters for artificial promoters containing hormone response elements (16, 40). In some cases, aspartic or glutamic acid substitutions have been used to mimic the negative charge of a phosphate. However, changes in structure as a result of the size of the phosphate group are not always reproduced by an acidic residue. Moreover, overexpressed mutants can work as dominant negatives competing with the endogenous receptors. To circumvent such inconvenient, a good alternative is to stably express the NR of interest in cells that are negative for the receptor. In line with this, stable HeLa cells expressing ERa WT or mutated at the phosphorylation sites have been established (53). Another alternative is to stably reintroduce the WT or mutated NRs in cells that have been invalidated for the receptor by homologous recombination (33, 54). Such cell lines are now used to study the effect of phosphorylation on gene expression by measuring variations in mRNA levels of endogenous established NR-target genes by quantitative RT-PCR, using gene-specific primer pairs (33, 37).

Despite the huge amount of research related to phosphorylation, the detailed role that specific phosphosites play in the function of NRs as of most individual proteins remains poorly understood. This is a challenging task as phosphorylation can alter either the structure of NRs, ligand binding, NRs interaction with coregulators, or their affinity for DNA.

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7.1. Phosphorylation and NRs Structure

In general, the phosphorylation sites are located in the functional domains of NRs, the well structured Ligand and DNA binding domains or the highly unstructured N-terminal domain. The motifs that are associated with phosphorylation sites occur predominantly within flexible regions (5, 6) such as loops between the LBD helices (55) or within intrinsically disordered regions such as the NTD (56). Unfortunately, no structural information is available on conformational changes due to phosphorylation due in part to the difficulty of obtaining sufficient purified phosphorylated NRs. However, nuclear magnetic resonance (NMR), electron paramagnetic resonance (EPR), and circular dichroism (CD) experiments should provide some information (57). Indeed such approaches indicated that phosphorylation of unstructured flexible domains within other transcription factors may induce changes in the structural properties of the domain with profound impact on its interaction with coregulators (5) and/or on the dynamics of adjacent structured domains (58). Recently, databases of 3D structures of protein phosphorylation sites have been developed (59) such as Phospho3D (http:// cbm.bio.uniroma2.it/phospho3d) (60) and DISPHOS (http:// core.ist.temple.edu/pred/pred.html) (6). Therefore, computational studies started to play a central role to predict how phosphorylation can induce relatively small conformational changes (61). Simulations performed with peptides bearing a phosphorylated versus nonphosphorylated serine have shown that phosphorylation stabilizes alpha-helix formation when located at the N-terminus while it destabilizes at the interior (62). Phosphorylation can also induce cis-trans isomerization of the proline residues following phosphorylated serines (61, 63). Within peptides with random conformation (37, 59), phosphorylation rather confers a more structured conformation. One of the major determinants of stabilization might be the formation of hydrogen bonds between the phosphate moiety on the serine and side chains of basic adjacent residues (37, 59).

7.2. NR Phosphorylation and Ligand Binding

Ligand binding acts as a switch on and is therefore one of the most important events in the control of NRs activity. In general, ligand binding is analyzed in in vitro equilibrium-based ligand binding assays, using tritiated or fluorescein-conjugated hormones (36, 64). Phosphorylation was found to differentially modulate affinity for the ligand, depending on the phosphorylated domain. As an example, phosphorylation of the ERa LBD was found to increase affinity for estradiol (11, 36), while phosphorylation of the PPARg NTD rather reduces ligand binding (64). No molecular or structural mechanism has been correlated yet to the former effect, but the latter has been correlated to phosphorylation-dependent modifications of the LBD conformation.

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7.3. Phosphorylation and Coregulators Interactions

As the NTD contains most phosphorylation sites, the challenge was to isolate coregulators interacting specifically with the phosphorylated or nonphosphorylated domain. Unfortunately, the classical Yeast Two Hybrid screening (65) could not be efficiently used because of the intrinsic transcriptional activity of this domain. Therefore, researchers rather used pull down experiments or phage display screens with immobilized GST-tagged NTDs (66, 67) or far western blotting with biotin-tagged NTD probes (68). Such strategies allowed the isolation of several phospho-dependent interactants for the NTD of ERa (66–68), RARa (69), or GR (37). Coregulators interacting specifically with the nonphosphorylated NTD have also been isolated (70). Note that for such coregulators, phosphorylation impedes or disrupts the interaction. Now mass spectrometry is again the tool of choice to identify proteins that bind to the NTD of NRs in a phosphorylationdependent manner. Such interactions can be determined by peptide pull downs where the unmodified and modified peptides are immobilized on a resin and each incubated with extracts derived from cells of interest (71). Quantification is also possible when combined to SILAC. Together with software advances, these new strategies should give rise to the discovery of a battery of new phosphorylation-dependent coregulators. Once such partners are isolated, the influence of phosphorylation on their interaction with NRs has to be further validated. Today, the standard GST-pull down and coimmunoprecipitation experiments using receptors with alanine or phosphomimetic substitutions are not recommended due to steric hindrance and charge differences between a phosphate group and an acidic residue. Now, real time biophysical techniques are preferred such as Plasmon resonance (Biacore) (72), provided that the coregulator or its interacting domain can be covalently immobilized or captured to the sensor surface in the active form. Then, synthetic peptides in which the serine (threonine or tyrosine) residue is phosphorylated or not, are injected. The inconvenient of this technique is the requirement of highly purified coregulators. However, it has the advantage of determining in real time the kinetic and affinity parameters of the interaction (73). Finally, a recent and unique approach to monitor the dynamic association–dissociation of proteins within living cells is Fluorescence Resonance Energy Transfer (FRET), associated to Fluorescence Lifetime Imaging Microscopy (FLIM). These techniques are based on energy transfer from a fluorophore in an electronic excited state serving as a donor to an acceptor chromophore, using pairs of CFP-YFP or GFP-DsRed tagged molecules. It becomes increasingly used to study NR domain interactions with cofactors motifs (50, 74) and will be promising to analyze the influence of phosphorylation on coregulators binding (36).

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In contrast to the NTD, the LBD interacts with a huge number of coregulators and for most of them the interaction relies on ligand-induced conformational changes of the interacting surface. However, according to recent studies, phosphosites located in flexible regions of the LBD such as loops, do not seem to control directly the binding of coregulators to the domain but rather have profound effects on coregulators binding at adjacent regions very likely through allosteric control. As an example, phosphorylation of RXRa at a residue located in the omega loop alters the conformation of the nearby coactivator’s interaction surface, and therefore impedes the recruitment of coactivators (75). In contrast, phosphorylation of RARa at S369 in loop 9–10 increases the binding of the cyclin H subunit of the general transcription factor TFIIH at the nearby N-terminal end of H9 (33, 55). It must be noted that these phosphorylation sites are not conserved between NRs indicating the existence of receptor specific, phosphorylation-dependent fine-tuning. 7.4. Phosphorylation and NRs DNA Binding

NRs regulate gene expression through binding to specific response elements located in the promoters of target genes. While steroid receptors bind DNA exclusively as homodimers, nonsteroid receptors bind as heterodimers with RXRs. Whether phosphorylation plays a role in NRs binding to DNA was initially studied in Electro Mobility Shift Assays (EMSA) using a radiolabeled oligopeptide corresponding to a response element and recombinant NRs phosphorylated with several kinases either in vitro or in transfected cells. In some cases, the results were corroborated in super shifts experiments performed with phosphosite-specific antibodies (55). Such approaches indicated that phosphorylation of serine residues involved in the recognition of the cognate response elements or located within the DBD or LBD dimerization surfaces decrease DNA binding (76–78). In contrast, phosphorylation of residues located in other domains such as the NTD rather increased the receptor-DNA interaction (36, 55), highlighting the possibility of interdomain communication. Recent investigations using Fluorescein-labeled oligonucleotides and increasing concentrations of control ER or phosphorylated ER suggested that phosphorylation of these residues might alter the conformation or apparent size of the NR-DNA complex (36). However, such assays do not take into account the need to modify the chromatin of target genes integrated in the genome nor interactions with other transcription factors. Therefore, chromatin immunoprecipitation experiments (ChIP) are now used to study the recruitment of different phospho NRs at endogenous genes promoters using antibodies against total NRs or phosphorylated NRs. Using NR null or negative cells expressing NRs mutated at the phosphorylation sites often completes such a strategy. Such approaches indicated that phosphorylation controls the recruitment of RARa, ERa, and GR to target promoters (33, 79, 80).

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There is an increasing evidence that phosphorylation controls the expression of NRs target genes with a promoter context dependence (54). However, there are no data indicating whether this promoter specificity reflects differences in NR interactions with response elements. Currently the most commonly used high-throughput method for identifying NR binding sites is chromatin immunoprecipitation followed by microarray hybridization (ChIP-chip) (81). However, new methods have recently been developed to take advantage of the next-generation highthroughput sequencing technologies. In one such method, ChIP-seq, immunoprecipitated DNA fragments are directly sequenced, and the short sequence reads are then mapped to the reference genome (82, 83). Combined with the use of phosphosite-specific antibodies, ChIP-seq should be a promising technique for the identification of DNA sequences binding specifically the phosphorylated NRs. Finally, it is now possible to study in real time the dynamics of the interaction of NRs with a DNA template taking advantage of the FRAP technology (50, 84). The use of fluorescently tagged GR combined with that of cells containing tandem arrays of the MMTV promoter with GR binding sites, permitted to visualize the rapid exchange rates of GR with specific DNA binding sites (49, 85). The FRAP strategy should address the influence of phosphorylation at specific sites or of pharmacological kinase or phosphatase inhibitors on most NRs dynamics.

8. Phosphorylation and Other NR Modifications

NRs are also targets for other modifications such as ubiquitination, sumoylation, acetylation, and methylation (9, 86, 87). Today it is admitted that interplay between different Post-translational modifications is an important mechanism to achieve an integrated regulation of NRs activity. Only a few studies reported the influence of phosphorylation on sumoylation (88) and acetylation (89). However, the best example of cross-talk between modifications is the phosphorylation-dependent ubiquitination and subsequent proteasomal degradation of most NRs such as ER, PR, GR, and RAR (56, 90). On the basis of this function of phosphorylation was the observation that NRs with the phosphorylation sites substituted with alanine residues are more stable. Additionally, these mutants exhibit reduced ubiquitination and degradation by the 26S proteasome upon cognate ligand binding. The influence of phosphorylation on ubiquitination was originally investigated in NR immunoprecipitation experiments followed by immunoblotting analysis of ubiquitin. Today, the increasing number of phosphosite-specific antibodies should facilitate the investigations.

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Whether phosphorylation controls the recruitment of the ubiquitin–proteasome machinery directly or indirectly through conformational changes requires further investigations taking advantage of the recent MS-based strategies described above.

9. Conclusion Since the original classical experiments, considerable progress has been made in the identification of NRs phosphorylation sites and in our understanding of the role of these modifications in the control of NRs activity. Indeed, new technologies such as chromatography for phosphopeptide enrichment and high accuracy mass spectrometry allowed the identification of multiple phosphorylation sites in NRs, with some residues being constitutively phosphorylated, while others become phosphorylated in response to the ligand or to specific cell signaling pathways. It also allowed to decipher fine-tuned interplays between some phosphorylation sites. Indeed, phosphorylation of each site can occur separately from the others or depend on a priming phosphorylation event. In the future, improvement of the purification and quantification methods, combined with advances in automation and with development of more robust and specific software tools, should allow the identification of numerous new NR phosphorylation sites, even with low abundance. Today, an integrated strategy for analysis of a phospho NR would include the following consecutive steps: prediction by computational analysis, phosphorylation (in vivo or in vitro by activated kinases), separation of the phosphorylated receptor followed by tandem MS/MS analysis of the phosphosites, and finally validation of the phosphorylation sites (22) (Fig. 1). Such an integrated strategy should provide data with a treasure-trove of information about the integration of numerous signaling events by NRs. Now the future challenges are to connect these data directly with new highly sensitive, real time or large-scale technologies, in order to get novel critical information about the influence of each phosphosite on the regulation of NRs activity (Fig. 1). New biophysical approaches such as NMR, FRET, FRAP, and FLIM are promising tools to investigate how phosphorylation fine-tunes the structure and the intracellular localization of NRs as well as their interactions with new coregulators. Large-scale microarrays, ChIP-seq, and quantitative proteomics should also provide interesting information about the downstream gene-expression and protein complexes changes controlled by NRs phosphorylation. Finally, large-scale and quantitative phosphorylation screens of NRs combined with other large-scale data sets should pave the way to breakthroughs in disease-related research.

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Fig. 1. Strategies for integrated analysis of nuclear receptors phosphorylation.

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Lalevée, Ferry, and Rochette-Egly Kremer R (2008) Phosphorylation of the human retinoid X receptor alpha at serine 260 impairs coactivator(s) recruitment and induces hormone resistance to multiple ligands. J Biol Chem 283:4943–4956 Yoshimura K, Muto Y, Shimizu M, Matsushima-Nishiwaki R, Okuno M, Takano Y, Tsurumi H, Kojima S, Okano Y, Moriwaki H (2007) Phosphorylated retinoid X receptor alpha loses its heterodimeric activity with retinoic acid receptor beta. Cancer Sci 98:1868–1874 Chen D, Pace PE, Coombes RC, Ali S (1999) Phosphorylation of human estrogen receptor alpha by protein kinase A regulates dimerization. Mol Cell Biol 19:1002–1015 Delmotte MH, Tahayato A, Formstecher P, Lefebvre P (1999) Serine 157, a retinoic acid receptor alpha residue phosphorylated by protein kinase C in vitro, is involved in RXR. RARalpha heterodimerization and transcriptional activity. J Biol Chem 274:38225–38231 Blind RD, Garabedian MJ (2008) Differential recruitment of glucocorticoid receptor phospho-isoforms to glucocorticoid-induced genes. J Steroid Biochem Mol Biol 109:150–157 Valley CC, Metivier R, Solodin NM, Fowler AM, Mashek MT, Hill L, Alarid ET (2005) Differential regulation of estrogen-inducible proteolysis and transcription by the estrogen receptor alpha N terminus. Mol Cell Biol 25:5417–5428 Carroll JS, Brown M (2006) Estrogen receptor target gene: an evolving concept. Mol Endocrinol 20:1707–1714 Dietz SC, Carroll JS (2008) Interrogating the genome to understand oestrogen-receptormediated transcription. Expert Rev Mol Med 10:e10

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Chapter 16 Regulation of Krüpple-Like Factor 5 by Targeted Protein Degradation Ceshi Chen Abstract Transcription factors are usually unstable proteins. The degradation of the majority of transcription factors is through the ubiquitin proteasome pathway and is tightly regulated by E3 ubiquitin ligases. KLF5 is an important transcription factor regulating cell proliferation, cell cycle, survival, migration, differentiation, angiogenesis, and stem cell self-renewal. We have shown that the WWP1 E3 ligase targets KLF5 for ubiquitin-mediated degradation. Several methods to determine whether a protein is ubiquitinated have been described [Kaiser, Tagwerker (Methods Enzymol 399:243–248, 2005); Bloom, Pagano (Methods Enzymol 399:249–266, 2005)]. This chapter focuses on experimental approaches testing KLF5 transcription factor ubiquitination and degradation by its E3s. Key words: Degradation, KLF5, Proteasome, Ubiquitination, WWP1

1. Introduction Ubiquitination is a common protein Post-translational modification which initiates protein degradation and signaling. Ubiquitin is a small conserved protein with 76 amino acids. Ubiquitin can be conjugated to a substrate’s lysine residue through a covalent isopeptide bond via a three-step cascade mechanism that is sequentially mediated by three enzymes: the ubiquitin-activating enzyme (E1), ubiquitin-conjugating enzyme (E2), and ubiquitin ligase (E3) (1, 2). Most E3s contain either a RING finger domain or a HECT domain. The E3 controls substrate specificity. Multiple ubiquitin molecules can form a polyubiquitin chain through the Lys (K) residues of ubiquitin. The K48-linked polyubiquitin chain will target proteins for degradation by the 26S proteasome. Many known transcription factors, such as p53, HIF-1a, b-catenin, and

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KLF5, are tightly controlled by the ubiquitin proteasome pathway. Degradation of these proteins is tightly regulated by their E3 ubiquitin ligases, such as Mdm2, VHL, SCFb-TrcP, and WWP1. These transcription factors and their E3s are well documented to play important roles in human cancers (3). The KLF family consists of over 20 members in humans, and is structurally characterized by three tandem zinc-finger domains at the C-terminus (4). The KLF5 protein is a ~55 kDa protein with two proline rich transactivation domains (TAD) (5, 6) (Fig. 1). The KLF5 protein undergoes various Post-translational modifications, such as phosphorylation (6), acetylation (7), sumolyation (8), and ubiquitination (9, 10). Like other KLF transcription factors, KLF5 binds to GC-rich DNA sequences, such as an Sp1 site, GC box, or CACCC box (6), through the zinc finger domains. The KLF5 protein has been shown to associate with numerous cofactors, such as RAR (11), NFkB (12), and C/EBP (8), to regulate gene transcription. KLF5 has been reported to regulate several target genes, such as the platelet-derived growth factor a (PDGF-a) (13), Cyclin D1(14), EGFR (15), PPARg (16), and Nanog (17), in different cell models. Accumulated evidence suggests that KLF5 promotes cell proliferation, cell cycle progression, survival, migration and invasion, differentiation, angiogenesis, and stem cell self-renewal. WWP1 belongs to the C2-WW-HECT-type E3 family, which comprises eight other members (18). All family members share a distinctive domain structure: a C2 domain at the N-terminus for calcium-dependent phospholipid binding, 2–4 WW domains in the middle for protein–protein interaction with PY motifs, and a HECT domain at the C-terminus for the ubiquitin transfer. Several studies suggest that WWP1 negatively regulates the transforming growth factor-b (TGF-b) signaling by targeting its molecular components, including the TGF-b receptor 1 (TbR1) (19), Smad2 (20), and Smad4 (21), for ubiquitin-mediated degradation. In addition, WWP1 has been reported to target the p53 (22) and p63 (23) transcription factors for ubiquitin-mediated proteolysis. After we demonstrated that KLF5 is ubiquitinated and degraded through the proteasome (9), we further mapped the

Fig. 1. Schematic showing the structural organization of the KLF5 protein and its Posttranslational modifications. TAD represents the transactivation domain. The PY motif represents the PPXY sequence. The three black boxes at the C-terminus represent zinc finger domains.

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destruction domain and found that the PY motif controls KLF5 ubiquitination and degradation. Following that, we showed that WWP1 targets KLF5 for ubiquitin-mediated proteasomal degradation via the WW-PY motif interaction (10). Here, we use KLF5 ubiquitination by WWP1 as an example to illustrate how to study transcription factor ubiquitination and degradation by its E3s. The following protocols are primarily based on our earlier publications (9, 10).

2. Materials 2.1. Cell Culture and Transfection

1. The 22Rv1 prostate cancer cell line (ATCC, Manassas VA) RPMI-1640 medium supplemented with 5% fetal bovine serum (FBS), HEPES (0.1 M), sodium pyruvate (1 mM), sodium bicarbonate (0.15%), glucose (0.45%), and penicillin and streptomycin (1%) 2. DMEM (met/cys-free) (Invitrogen, Carlsbad, CA) 3. Dialyzed FBS (Invitrogen) 4. MG132 (Sigma, St. Louis, MO) is dissolved at 20 mM in dimethylsulfoxide (DMSO) and stored at −20°C 5. Cycloheximide (Sigma) 6. Lipofectamine 2000 (Invitrogen) 7. pcDNA3.1-KLF5-FLAG and pcDNA3.1-KLF5DPY-FLAG (a mutant that cannot interact with WWP1 due to the lack of the PY motif) 8. pLenti6-WWP1 and pLenti6-WWP1C890A (catalytic inactive mutant) pcDNA3.1-HA-Ubiquitin 9. DC protein assay kit (Bio-Rad, Hercules, CA) 10. The anti-b-actin mouse monoclonal antibody (Sigma) 11. Anti-KLF5 antibody (9) 12. Anti-WWP1 antibody (1A7, Novus Biologicals, Littleton, CO) 13. Anti-FLAG M2 mouse monoclonal antibody (Sigma) 14. Anti-FLAG M2 Affinity Gel (Sigma) 15. Anti-HA antibody (Cell Signaling, Danvers, MA) 16. The Ubiquitin–Protein Conjugation Kit (BostonBiochem, Cambridge, MA) 17. TNT Quick Coupled Transcription/Translation Systems (Promega, Madison, WI) 18. Protein Lysis Buffer: 50 mM Tris–HCl (pH 7.4), 150 mM NaCl, 1 mM EDTA, 1% Triton X-100, and 1% protease inhibitor cocktail I (Sigma)

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19. Denaturing Lysis Buffer: 50 mM Tris–Cl, pH 6.8, 1.5% SDS 20. 3×SDS Sample Buffer: 0.5 M Tris–HCl (pH 6.8), 3% SDS, 30% glycerol, 3% b-Mercaptoethanol, 0.01% bromophenol blue 21. 10× SDS–PAGE Running Buffer (Bio-Rad) 22. Transfer Buffer: 25 mM Tris base, 192 mM glycine, 20% methanol (pH 8.3) 23. 10×Phosphate Buffered Saline (PBS, Hyclone) 24. Ponceau S (Sigma) 25. SuperSignal West Pico Substrate (Pierce, Rockford, IL) 26. EBC/BSA Buffer (with 1% protease inhibitor cocktail I): 50 mM Tris–Cl, pH 6.8, 180 mM NaCl, 0.5% CA630, 0.5% BSA 35

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S-L-Methionine, in vitro Translation Grade, >1,000 Ci/ mmol; >37 TBq/mmol (MP Biomedicals)

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S-Trans label, ~10 mCi/ml (ICN, Irvine, CA)

3. Methods 3.1. Chase Assays for the KLF5 Degradation by WWP1 In Vivo

3.1.1. The CHX Chase Assay

The most common method of testing whether an interesting transcription factor is degraded by its E3 via the ubiquitin proteasome pathway in cells is to test whether the protein stability is decreased after the coexpression of the wild type E3 and increased after knocking down the endogenous E3 by RNA interference. The transcription factor destruction by its E3 should be blocked by treating the cells with cell-permeable proteasome inhibitors, such as MG132, Epoxomicin, and Lactacystin. The protein stability is usually examined by the cycloheximide (CHX) assay or the pulse chase assay. 1. Seed 22Rv1 cells in four 12-well plates at the 5 × 105/well density, incubate overnight. 2. On the second day, cotransfect the cells with plasmids expressing KLF5 (WT KLF5-FLAG and the mutant KLF5DPYFLAG) and WWP1 (WT WWP1, WWP1m, and the vector control) by using Lipofectamine 2000 (six groups × eight wells for each group). 3. Two days later, add MG132 (20 mM) and DMSO (four wells for MG132 and the remaining four wells for DMSO for each group). Add cycloheximide (CHX, 20–100 mg/ml) into the wells labeled with 3 h, 2 h, and 1 h at the corresponding time points (see Note 2). Skip the time zero wells.

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4. Wash the cells with the PBS buffer once, collect the cell lysate using the protein lysis buffer (120 ml per well). 5. Measure the protein concentration by using the DC protein assay kit. We use BSA (0–4 mg/ml) as the standard. 6. Load 20–50 mg protein to SDS–PAGE gels (8% acrylamide) and transfer to PVDF membranes in transfer buffer for 1 h at 100 V in a cold room. 7. Disassemble the sandwiches, put the blots in 0.5% ponceau S, shake 1 min, wash blots with milli-Q H2O, cut blots according to the molecular standard, and wash blots twice with milli-Q H2O. 8. Block the blots with 5% milk in PBST (PBS buffer with 0.1% Tween 20) for 1 h at room temperature. 9. Incubate the blots with primary (anti-KLF5, anti-b-actin, and anti-WWP1) antibodies (see Note 3) diluted with 3% BSA in PBST overnight at 4°C. 10. Wash the blots twice for 10 min each with PBST. 11. Incubate the blots with secondary antibodies diluted in 3% milk for 1 h at room temperature. 12. Wash the blots three times with PBST. 13. Incubate the blots with the SuperSignal West Pico Substrate for 5 min. 14. Collect images by using the Fujifilm Imaging system LAS3000. The advantage of the CHX assay is that neither radioactive material nor immunoprecipitation is required. However, the results should be interpreted cautiously because the protein synthesis for E3 is also blocked by CHX. Transcription factors could be stabilized due to the rapid degradation of the E3 ligase. If E3 has a short half-life, the pulse chase assay is recommended to measure the transcription factor degradation. 3.1.2. The Pulse Chase Assay

1. Seed 3 × 106 22Rv1 cells into 6-cm dishes (eight in total), incubate overnight. 2. The next day, cotransfect the cells with plasmids expressing KLF5 (WT KLF5-FLAG) and WWP1 (WT WWP1 and the empty vector control) by using Lipofectamine 2000 (two groups × four wells for each group). 3. 48 h later, aspirate the media from each plate and wash the cells twice with 5 ml warm PBS. 4. Add 1 ml warm DMEM (met/cys-free) with 5% dialyzed FBS to each dish and incubate the cells for 30 min at 37°C. This is the starvation period.

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5. Add 33 ml of 35S-Trans-label to each plate in a cell culture hood (at ~10 mCi/ml) to label proteins in vivo. 6. Carefully place the cells in an incubator and incubate for 40 min at 37°C. 7. Carefully remove the radioactive media into 35S liquid waste bottles using pipets with a cotton plunge. Wash the cells on each plate twice with 5 ml warm PBS. 8. Add 5 ml warm DMEM+FBS+2 mM Methionine+2 mM Cysteine to all plates except the zero “0” samples. Start timer. 9. Immediately remove all “0” plates. Return the other plates to 37°C. 10. Remove the media from the “0” plates. Wash the cells with 2 ml PBS twice. 11. On ice, add denaturing lysis buffer (0.2 ml for each dish) and collect the proteins into test tubes with screw caps by scraping. 12. Perform similar collections at the different time points during the chase (1–3 h). 13. Boil the samples for 15 min on a heat block. 14. Take all the protein, add 1.25 ml EBC/BSA buffer. 15. Add 20 ml prewashed FLAG-M2 affinity gel and rotate overnight in a cold room. 16. Spin down the beads at 10,000 × g, 5 s at 4°C. 17. Remove the supernatant and wash the beads three times with 1 ml ice-cold EBC/BSA buffer, vortex each time. 18. Resuspend the beads in 30 ml 3× SDS–PAGE sample buffer. 19. Denature the proteins and resolve the samples in SDS–PAGE gels. 20. Dry gels and autoradiography. 3.2. The Ubiquitination Assay In Vitro for KLF5

To demonstrate that the degradation of a transcription factor proceeds in a ubiquitin-dependent manner, it is essential to demonstrate the ubiquitin-conjugated intermediates. Typically, incubation of the 35S-methinone-labeled transcription factor protein in a cell extract supplemented with ubiquitin, ATP, and the deubiquitinase inhibitor ubiquitin aldehyde will cause the accumulation of high molecular mass adducts in vitro. Addition of the purified wild type E3 enzyme, but not the ligase dead mutant, will increase the amount of ubiquitinated transcription factor species. These ubiquitin conjugated proteins can be detected by autoradiography or immunoblotting after SDS–PAGE.

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1. Amplify the full length KLF5 gene by PCR using a forward primer with the T7 promoter sequence and a backward primer. The forward primer should be purified by PAGE after synthesis because of its length. 2. Synthesize the KLF5 protein by in vitro translation in the presence of 35S-methinone using the TNT Quick Coupled Transcription/Translation Systems by following the protocol provided in the manual (see Note 4). 3. Resolve 2 ml translated samples by SDS–PAGE to examine the synthesized KLF5 protein by autoradiography. 4. Mix 8 mg Fraction A (predominantly E1 and E2 enzymes), 8 mg Fraction B (predominantly E3 and deubiquitinating enzymes), 26 mg ubiquitin, 4 mM ubiquitin aldehyde, and 2.5 ml energy solution (10×) in 25 ml volume in Eppendorf tubes. GST-WWP1 (2.5 mg), GST-WWP1m, and GST are added into three different reactions (see Note 5). Addition of all the reagents should be carried out on ice. 5. Add 2 ml of translated 35S-labeled KLF5 into each reaction. 6. Incubate the mixture at 37°C for 30 min. 7. Stop the reaction by adding 10 ml 3×sample buffer and denature samples. 8. All samples are subjected to SDS–PAGE (10% acrylamide), load 2 ml of 35S-labeled KLF5 as the input. 9. Dry the gels and expose the dried gels to film (see Note 6). 3.3. Immunoprecipitation (IP) Under Denaturing Conditions for the KLF5 Protein Ubiquitination by WWP1 In Vivo

It is essential to demonstrate the transcription factor is also ubiquitinated by its E3 in cultured cells and in the cell-free system. After the E3 ligase is overexpressed or silenced in cells, proteasome inhibitors are added to accumulate the ubiquitinated transcription factor proteins. Following that, the transcription factor proteins are immunoprecipitated under denaturing conditions. The ubiquitinated transcription factor proteins can be detected by immunoblotting using an antiubiquitin antibody (see Note 7). The denaturing conditions are used to avoid the artifact that the ubiquitinated proteins are the transcription factor associated proteins. 1. Seed 1 × 106 22Rv1 cells into 6-well dishes (7-wells in total), incubate overnight. 2. The next day, co-transfect the cells with plasmids expressing KLF5 (WT KLF5-FLAG and KLF5DPY-FLAG), WWP1 (WT WWP1, WWP1m, and the empty vector control), and HA-Ubiquitin (see Note 8) by using Lipofectamine 2000 (one dish transfected with WT KLF5 and WWP1 but no HA-Ubiquitin is used as a negative control).

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3. Two days later, treat the cells with 20 mM MG132 for 4 h to block protein degradation. 4. Rinse the cells in the dishes once with the PBS buffer. 5. Add 150 ml denaturing lysis buffer to each well. 6. Collect the proteins by scraping. 7. Boil the samples for 15 min in a heat block. 8. In 1.5 ml Eppendorf tubes, put 70 ml of the denatured proteins into 1.2 ml EBC/BSA buffer to avoid denaturing the antibodies by high concentrations of SDS. 9. Add 20 ml prewashed FLAG-M2 affinity gels and rotate overnight in a cold room. 10. Spin down the beads at 10,000 × g, 5 s at 4°C. 11. Remove the supernatant and wash the beads three times with 1 ml ice-cold EBC/BSA buffer, vortex each time. 12. Resuspend the beads in 30 ml 3× SDS–PAGE sample buffer. 13. Denature the proteins and resolve the samples in SDS–PAGE gels. 14. Perform Western blotting with the anti-HA-Ab, anti-KLF5 Ab, and anti-WWP1 Ab (see Notes 9–12).

4. Notes 1. The example chase results are shown in Fig. 2. The degradation rate of KLF5 is slightly faster in pulse chase experiments than in CHX chase experiments. 2. The chase time should be predetermined for different transcription factors by performing a pilot experiment with multiple time points. The time points should include the time when the protein is reduced by half in the presence of its E3.

Fig. 2. Measuring the KLF5 protein half-life by CHX and pulse chase assays in 22Rv1. (a) The CHX chase assay. The stable protein b-actin served as a control. CHX (100 mg/ml) was used to block protein synthesis. (b) The pulse chase assay (see Note 1).

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3. The use of epitope tags on transcription factors can lead to artifacts. If possible, use the antibodies directly against the transcription factor. In the case of KLF5, a FLAG tag at the N-terminus, but not the C-terminus, of KLF5 stabilizes the protein (24). 4. The rabbit reticulocyte ubiquitin conjugate system does not contain any proteasome activity so no proteasome inhibitors need to be added. 5. The transcription factor ubiquitin conjugation assay could be performed with purified E1, E2, and E3. However, other Post-translational modifications and/or binding partners may be essential for KLF5 ubiquitination by WWP1. 6. To further demonstrate that the high molecular mass adducts are indeed ubiquitin conjugates of the transcription factor, several additional experiments could be performed: (a) ATP dependence: the adducts will not be generated without ATP (energy solution). (b) Ubiquitin dependence: the adducts will not be generated without Ubiquitin. (c) MeUb: the adducts will be inhibited by adding MeUb (a methylated derivative of ubiquitin lacking free amino groups) so polyubiquitin chains cannot be efficiently formed. 7. The KLF5 ubiquitination can be detected by direct immunoblotting. The high molecular species are dramatically increased in the presence of WWP1 and MG132 (10). 8. If the His-Ubiquitin is used instead of HA-Ubiquitin, the proteins can be purified by Ni-NTA agarose beads under denaturing conditions (1). The ubiquitinated proteins can be detected by an antibody directly against the transcription factor. 9. The type of polyubiquitin chain can be distinguished by polyubiquitin linkage-specific antibodies (25). 10. One example result is shown in Fig. 3b. In this case, WT WWP1 clearly increases the KLF5 ubiquitination in the 22Rv1 cells. 11. Transcription factor ubiquitination and degradation by its E3 may be regulated by signals, such as phosphorylation (e.g., b-catenin by SCFb-TrcP) and hydroxylation (e.g., HIF1a by VHL). 12. Once the transcription factor has been shown to be ubiquitinated and degraded by a candidate E3, the techniques described should be applied to test the endogenous protein.

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Fig. 3. Measuring the KLF5 ubiquitination in vitro and in vivo. (a) WWP1 ubiquitinates KLF5 in vitro. The recombinant GST-WWP1 increases KLF5 polyubiquitination. The in vitro translated KLF5 protein is used as input (lane 1). It is worth pointing out that KLF5 is also ubiquitinated without adding exogenous purified GST-WWP1 because of other KLF5 E3s in the cell-free system. (b) WWP1 ubiquitinates KLF5 in 22Rv1 cells. HA-Ubiquitin was co-transfected. The cells were treated with 20 mM MG132 to block protein degradation.

Acknowledgments This work was supported in part by a grant (RSG-08-199-01) from the American Cancer Society, a grant (BCTR0503705) from Komen for the Cure, and a grant (W81XWH-07-1-0191) from the Department of Defense. References 1. Kaiser P, Tagwerker C (2005) Is this protein ubiquitinated? Methods Enzymol 399: 243–248 2. Bloom J, Pagano M (2005) Experimental tests to definitively determine ubiquitylation of a substrate. Methods Enzymol 399: 249–266 3. Chen C, Seth AK, Aplin AE (2006) Genetic and expression aberrations of e3 ubiquitin ligases in human breast cancer. Mol Cancer Res 4:695–707 4. Black AR, Black JD, Azizkhan-Clifford J (2001) Sp1 and Kruppel-like factor family of transcription factors in cell growth regulation and cancer. J Cell Physiol 188:143–160 5. Kojima S, Kobayashi A, Gotoh O, Ohkuma Y, Fujii-Kuriyama Y, Sogawa K (1997) J Biochem (Tokyo) 121:389–396 6. Zhang Z, Teng CT (2003) Phosphorylation of Kruppel-like factor 5 (KLF5/IKLF) at the CBP interaction region enhances its transactivation function. Nucleic Acids Res 31: 2196–2208

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Chapter 17 Post-translational Control of ETS Transcription Factors: Detection of Modified Factors at Target Gene Promoters Li Li, Janice Saxton, and Peter E. Shaw Abstract ETS transcription factors are implicated in gene regulation during cell proliferation and in the development of the haematopoietic cell lineage. Characteristically, ETS proteins act in concert with other transcription factors and are regulated by post-translational modifications, most frequently phosphorylation. These events have been shown to modulate the DNA binding affinity and interactions of ETS transcription factors with co-activators, events that can ultimately determine the formation of productive transcription complexes on target gene promoters. However, direct implication of a transcription factor or one of its post-translational modifications in the regulation of a given gene requires detection of the modified factor at the target gene promoter. Chromatin immunoprecipitation assays were originally adopted to probe modifications to histone proteins associated with transcriptionally active genes in yeast. They have since been used to confirm the presence of numerous proteins at diverse gene promoters including, for example, recruitment of the mitogen-activated protein (MAP) kinases ERK1 and ERK2 to the promoters of mitogen-responsive genes. Here chromatin immunoprecipitation is used to demonstrate the inducible appearance of phosphorylated Elk-1 at the human c-fos promoter. Keywords: Chromatin immunoprecipitation (ChIP), DNA binding, Elk-1, MAP kinase, Mitogens, Phosphorylation, Semi-quantitative polymerase chain reaction (PCR)

1 Introduction The 50 or so known ETS transcription factors share in common a winged helix-turn-helix DNA-binding domain and are implicated in the regulation of gene expression in diverse contexts, including cell proliferation and lineage development [1, 2]. The activity of many ETS transcription factors is subject to modulation by post-translational modifications (PTMs), commonly but by no means exclusively phosphorylation [3]. This is exemplified by the profound change in transcriptional activity that follows the phosphorylation of ternary complex factors such as Elk-1 by Paul J. Higgins (ed.), Transcription Factors: Methods and Protocols, Methods in Molecular Biology, vol. 647, DOI 10.1007/978-1-60761-738-9_17, © Springer Science+Business Media, LLC 2010

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the mitogen-activated protein kinases ERK1 and ERK2. Elk-1 phospho-activation was first observed as an increase in its binding affinity for serum response factor (SRF) bound at the c-fos SRE [4]. Phospho-peptide mapping and functional analyses of Elk-1 point mutants subsequently identified key phosphorylation sites in Elk-1 [5–7] and informed the development of phospho-specific antibodies against Elk-1 with which its activation can be monitored under a range of experimental circumstances. The diversity of PTMs with the potential to influence gene expression is illustrated by a consideration of those detected at histone tails [8]. The development of reagents to analyse these dynamic modifications has mirrored the example of phosphospecific antibodies and a battery of antibodies now exists for both generic modifications, such as acetyl-lysine or mono/dimethyl arginine, and unique modified substrates, such as ERK1/2 phosphorylated at the TEY motif or Elk-1 phosphorylated at S383. The development of further antibodies for the detection of PTMs to transcription factors and co-activators will continue to be an important contribution to their study, but direct implication of any such modification in the regulation of a specific gene requires detection of the appropriately modified factor at the target gene promoter. This can be achieved by chromatin immunoprecipitation (ChIP) assay, a technique that gained widespread recognition through the development of antibodies directed at specific histone modifications [9]. In this scenario, an antibody raised against a modified transcription factor would be used to pinpoint the appropriately modified factor to a particular promoter. Moreover, if the modification is induced and/ or labile, time-resolved ChIP assays can follow the appearance and/or disappearance of the modification from the promoter. This approach is described here showing the appearance of phospho-Elk-1 at the c-fos promoter following mitogen stimulation of HeLa cells.

2 Materials 2.1 Cell Culture, Cross-Linking, Lysis and Sonication

1. Dulbecco’s Modified Eagle’s Medium (DMEM) (Lonza) supplemented with 10% foetal calf serum (FCS) (Autogen Bioclear), 2 mM l-glutamine (Sigma), 100 U/mL penicillin and 100 mg/mL streptomycin (Sigma). 2. Falcon 10 cm cell culture dishes (Becton Dickinson). 3. Phorbol 12-myristate 13-acetate (PMA, 10 mg/mL in DMSO, Calbiochem). 4. Formaldehyde (36.5%, Sigma).

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5. Pre-lysis buffer (PLB): Phosphate buffered saline (PBS) containing 125 mM glycine, 1 mM EDTA, 1 mM phenylmethanesulphonyl fluoride (PMSF) (freshly added). 6. Protease inhibitor cocktail (complete, EDTA-free, Roche). 7. Microfuge tubes, 1.5 mL. 8. Lysis buffer: 50 mM Tris–HCl, pH 8.0, 1% sodium dodecyl sulphate (SDS), 10 mM EDTA containing 1× protease inhibitor cocktail (freshly added). 9. Dilution buffer: 20 mM Tris–HCl, pH 8.0, 1% Triton X-100, 2 mM EDTA, 150 mM NaCl, containing 1× protease inhibitor cocktail (freshly added). 10. Cooled bench-top microfuge (Eppendorf, model 5417R). 11. Sonicator (Sonics Vibracell VC 50T). 2.2 Chromatin Immunoprecipitation and Washing Immune Complexes

1. Antibodies: the B-4 pElk-1 mouse monoclonal, recognising phospho-S383 of Elk-1, was from Santa Cruz. The anti-Elk-1 antibody I-20 from Santa Cruz (sc-355, lot D2007, 200 mg/ mL) precipitates total Elk-1, i.e. both unphosphorylated and phosphorylated forms. However, the anti-Elk-1 antibody I-20 (sc-355×, lot E2606, 2 mg/mL) precipitates only the unphosphorylated form of Elk-1 [10]. 2. Vortex mixer (Vortex Genie 2, Scientific Industries). 3. Vertical wheel mixer (Stuart Scientific SB2 Rotator). 4. Sonicated salmon sperm DNA (GE Healthcare). 5. Protein G sepharose beads (Fast Flow, GE Healthcare). 6. Sepharose bead wash buffer (BWB): 10 mM Tris–HCl, pH 8.0, 1 mM EDTA, 120 mM NaCl. 7. Bovine serum albumin (BSA, Sigma). 8. 5 M NaCl. 9. Triton X-100 (Alfa Aesar). 10. Nonidet P 40 (NP-40, Fluka BioChemika). 11. 8 M LiCl. 12. Deoxycholate, sodium salt (Sigma). 13. 0.5 M EDTA, pH 8.0. 14. TSE I: 20 mM Tris–HCl, pH 8.0, 0.1% SDS, 1% Triton X-100, 2 mM EDTA, 150 mM NaCl. 15. TSE II: 20 mM Tris–HCl, pH 8.0, 0.1% SDS, 1% Triton X-100, 2 mM EDTA, 500 mM NaCl. 16. Buffer III: 10 mM Tris–HCl, pH 8.0, 1% NP-40, 1 mM EDTA, 1% deoxycholate, 0.25 M LiCl. 17. TE buffer: 20 mM Tris–HCl, pH 8.0, 1 mM EDTA.

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2.3 Elution, Cross-Link Reversal and DNA Purification

1. 10% SDS. 2. 1 M NaHCO3. 3. 5 M NaCl. 4. Proteinase K, 20 mg/mL (Sigma). 5. 1 M Tris–HCl, pH 6.5. 6. 0.5 M EDTA. 7. 3 M sodium acetate (NaAc), pH 5.2. 8. Elution buffer: 1% SDS in 0.1 M NaHCO3. 9. Heated shaker (Thermomixer compact, Eppendorf). 10. Bench-top centrifuge (Eppendorf, Model 5415C). 11. PCR purification (QIA-Quick Purification Kit, Qiagen).

2.4 PCR Analysis

1. PCR tubes (0.5 mL, Axygen Scientific). 2. Primers for human c-fos promoter (−472 to −276): 5¢-GGGTCCGCATTGAACCAGGTGC (forward) 5¢-GCCGTGGAAACCTGCTGACGCA (reverse). 3. Primers for human c-fos gene (+1,711 to +1,865): 5¢-CTGGGAACTCGCCCCACCTGTGTC (forward) 5¢-CACTGCAGGTCCGGACTGGTCGAG (reverse). 4. Pfu DNA polymerase (Stratagene). 5. 10× Pfu buffer (Stratagene). 6. Deoxyribonucleotide triphosphates (dNTP set, 100 mM solution, GE Healthcare). 7. PCR Thermocycler (TRIO-Thermoblock, Biometra). 8. 50× Tris–Acetate–EDTA buffer (TAE): 2 M Tris, 1 M Acetic acid, 50 mM EDTA, pH 8.0. 9. Ethidium bromide (10 mg/mL, Amresco). 10. Agarose gel apparatus (sub-cell GT wide mini, BioRad). 11. Molecular Biology Agarose (BioRad) and Ultrasieve Agarose (Biogene). 12. Gel loading buffer (6×): 15% Ficoll 400 (Sigma), 0.25% Xylene Cyanol FF in ddH2O. 13. UV trans-illuminator (BioDoc-It™ Image System, UVP).

3 Methods The detection of transient PTMs in a dynamic system requires a robust cell model in which the PTM is at first absent and can be induced in the majority of cells in a synchronised manner. For

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Elk-1 phosphorylation, this can be achieved in HeLa (and other) cells by serum withdrawal for 20–24 h to halt cells in G0, followed by mitogen stimulation to trigger G1 entry, for which we use PMA. For time-resolved ChIP assays involving time-spans of a few minutes, it is best to process time points individually. Although the phospho-serine modification is not particularly labile, it is nonetheless important after cross-linking to harvest cells swiftly in ice-cold buffers. Perhaps the most critical step in ChIP assays is shearing the DNA/chromatin. If the average size of DNA fragment is too long, spatial resolution along the gene will be lost as the IP will capture large DNA fragments stretching beyond the promoter of interest, which are a source of background signals. Conversely, if the DNA fragments are shorter on average than the intended PCR product, amplification will be compromised. When introducing new antibodies into ChIP analyses, it is important to assess the antigen accessibility in sheared chromatin samples. A good antibody will precipitate its antigen completely. This can be assessed by immunoblotting samples of the immunoprecipitate and the post-IP chromatin together for the antigen, which should be present in the immunoprecipitate (obviously) but absent from the post-IP chromatin. If a substantial proportion of the antigen remains in the chromatin, the antibody is unsuitable for ChIP assays, although the problem may be alleviated by increasing the amount of antibody used or by affinity purification. 3.1 Mitogen Stimulation, CrossLinking, Cell Lysis and Sonication

1. In preparation for a time course experiment, HeLa cells are cultured in 10 cm dishes for 48 h in DMEM with 10% FCS, 2 mM l-glutamine, 100 U mL penicillin and 100 mg/mL streptomycin to 70–80% confluence (corresponding to ~5 × 106 cells per dish) and then starved for 20–24 h in serumfree DMEM until stimulation with PMA (100 ng/mL) for the required time points, here 0, 10, 30, and 60 min. 2. At the appropriate times after PMA stimulation, add formaldehyde drop-wise directly into culture medium to a final concentration of 1%, mix by swirling gently and incubate at 37°C for 10 min. 3. Aspirate medium from cells and wash them twice with 15 mL of ice-cold PLB. 4. Scrape cells from dishes in 1 mL ice-cold PLB with a rubber policeman, transfer into 1.5 mL tubes and centrifuge immediately at 2,600 rpm (720 × g) at 4°C for 10 min. Carefully remove the supernatants from the cell pellets. At this point the cell pellets can be frozen at −80°C for later use. 5. Add 200 mL of ice-cold lysis buffer to the cell pellets and resuspend by drawing cells into the pipette tip 5–6 times. Incubate on ice for 10 min.

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6. To each 200 mL cell lysate, add 100 mL ice-cold Dilution buffer, mix and then divide each sample into two equal aliquots in 1.5 mL microfuge tubes, to optimise the volume for sonication. 7. To shear the chromatin/DNA, pack each tube with a 150 mL aliquot of cell lysate in ice (e.g. in a small plastic beaker) and sonicate for 4 × 15 s at 40% power with 30-s intervals. The sonicator tip (3 mm ø) should be immersed as far as possible, without touching the side of the tube, and samples must remain cold throughout. These conditions have been determined empirically to produce DNA fragments of 200–500 base pairs (see Note 1). 8. Sediment the residual insoluble chromatin by centrifugation in a bench top microfuge at 14,000 rpm (20,800 × g) for 10 min at 4°C. Carefully take the supernatants and recombine each pair of samples in a fresh tube. Set aside 10–20 mL of each chromatin preparation as the input fraction. The remaining soluble chromatin can be used right away or frozen at −80°C (see Note 2). 3.2 ChIP and Washing Immune Complexes

1. Dilute 140 mL of each soluble chromatin sample to 1 mL with ice-cold Dilution buffer in fresh 1.5 mL microfuge tubes (thereby diluting the SDS from the lysis buffer to 0.1%). Add 4 mg of antibody to each sample and incubate in a vertical wheel mixer over night in the cold room (4°C) (see Note 3). 2. The immune complexes are precipitated with protein G sepharose beads, which are supplied and stored as a 1:1 slurry and must be washed and pre-blocked with BSA before use. Re-suspend the bead slurry thoroughly and transfer 30 mL for each immunoprecipitation into a fresh 1.5 mL microfuge tube. (To improve accuracy when pipetting beads, 5–6 mm can be cut off from the end of the pipette tip with a scalpel or razor blade to widen the bore of the tip.) Collect beads from slurry by centrifugation at 4,000 rpm (1,700 × g) for 2 min. Wash the beads three times in 1 mL sepharose BWB for 5 min on wheel, collecting each time by centrifugation at 4,000 rpm (1,700 × g) for 2 min. (The washing steps can be expedited by aspirating the buffer from the tubes; to avoid losing beads use a glass pasteur pipette drawn out to a fine tip in a bunsen flame.) Pre-block the beads in 1 mL BWB containing 1% BSA for 2 h at 4°C on the vertical wheel. After blocking, wash the beads another three times in 1 mL BWB for 5 min on wheel, collecting by centrifugation at 4,000 rpm (1,700 × g) for 2 min. Take up the beads in their own volume of Dilution buffer to obtain a 1:1 slurry.

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3. To the immune complexes (formed over night), add 5 mg of sheared salmon sperm DNA and incubate for a further hour at 4°C. Aliquots (30 mL) of the pre-blocked and washed protein G sepharose bead slurry (thoroughly re-suspended) are then added to the immune complexes and incubated for 1 h at 4°C on the wheel. 4. Collect the immunoprecipitates/beads by centrifugation at 4,000 rpm (1,700 × g) for 2 min. Remove the supernatants and wash the beads sequentially for 10 min in 1 mL each of ice-cold TSE I, TSE II, Buffer III and twice with ice-cold TE buffer on the vertical wheel in the cold room (4°C), collecting the beads each time at 4,000 rpm (1,700 × g) for 2 min. 3.3 Elution, Cross-Link Reversal and DNA Purification

1. Elute the DNA–protein complexes from the beads in 75 mL Elution buffer at RT for 15 min in a shaker (600 rpm). Sediment the sepharose beads by centrifugation at 4,000 rpm (1,700 × g) for 2 min at RT and carefully transfer the supernatants to fresh 1.5 mL tubes. Avoid contamination with sepharose beads. Repeat the elution procedure once and pool the two eluates from each sample. 2. Mix each eluate (150 mL) with 6 mL of 5 M NaCl (to 0.2 M) and incubate at 65°C for 6 h to reverse the cross-linking. Collect the liquid by brief centrifugation. These reversed ChIP eluates can be retained in the same tubes. In parallel, reverse the input samples (dilute to 50 mL with Dilution buffer and add 2 mL of 5 M NaCl). Centrifuge input samples at 13,000 rpm (17,900 × g) for 2 min to sediment debris and transfer the supernatants to clean 1.5 mL tubes. At this stage, the input samples can be stored at −20°C. 3. To the reversed ChIP eluates add 1 mL proteinase K (20 mg/ mL), 6 mL 1 M Tris–HCl, pH 6.5, 3 mL 0.5 M EDTA and incubate for 1 h at 45°C. Centrifuge tubes briefly to collect the solution before DNA purification. At this stage, the reversed ChIP eluates may be stored at −20°C. 4. The DNA is purified from the ChIP samples with the QIAQuick PCR Purification Kit (Qiagen) as described in the manufacturer’s protocol. To the ~150 mL reversed ChIP eluates add 750 mL of buffer PBI (Qiagen) and mix. To ensure the pH remains below 7.5, which aids the recovery of DNA, add 10 mL 3 M NaAc, pH 5.2 to the mixture and mix thoroughly. 5. Apply the samples to QIA-Quick columns and centrifuge at 13,000 rpm (17,900 × g) for 1 min. Wash the columns with 750 mL of buffer PE (Qiagen), centrifuging the columns at 13,000 rpm (17,900 × g) for 1 min.

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6. Centrifuge the column once more at 13,000 rpm (17,900 × g) for 1 min to remove residual buffer PE and transfer the columns to clean 1.5 mL microfuge tubes. 7. Add 50 mL of buffer EB (Qiagen) to each column, allow to stand for 1 min and centrifuge at 13,000 rpm (17,900 × g) for 1 min. The eluted DNA samples can be used directly for PCR or stored at −20°C. 3.4 PCR Analysis

1. Set up the PCRs to detect the presence of target promoter DNA in 0.5 mL reaction tubes to a final volume of 25 mL. Each reaction will contain the following: 10× Pfu buffer, 2.5 mL dNTP mix (5 mM), 1 mL Forward primer (5 mM), 1 mL Reverse primer (5 mM), 1 mL Pfu polymerase (2.5 U/mL), 0.1 mL Eluted DNA samples, 2 mL ddH2O to 25 mL For input samples, the DNA is further diluted 1:10 with ddH2O and 2 mL are used per reaction. If the thermocycler to be used lacks a lid, samples must be overlaid with mineral oil (50 mL). 2. For the c-fos promoter and gene primer pairs indicated here, suitable thermocycler settings are as follows: Step 1: 95°C for 3 min Step 2: 93°C for 1 min Step 3: 60°C for 1 min Step 4: 72°C for 2.5 min Number of cycles: 34 (promoter) or 28 (gene) Step 5: 72°C for 5 min Step 6: 4°C indefinite 3. After the PCR, mix 10 mL of each reaction with 2 mL of 6× Gel loading buffer (see Note 4) and load onto a 2% Agarose gel (1% Molecular Biology Agarose and 1% Ultrasieve Agarose cast in 1 × TAE). The gel is run in 1 × TAE at 70 V for 50 min. 4. Stain the gel in Ethidium Bromide solution (0.5 mg/mL in 1 × TAE) for 15–30 min and wash in distilled water for 5–10 min. 5. Images of gels are obtained with a UV trans-illuminator and digital camera. An example of the results produced is shown in Fig. 1. The use of a digital imaging system allows subsequent

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Fig. 1 Appearance of phospho-Elk-1 at c-fos promoter in response to mitogens. HeLa cells were serum starved or starved and stimulated with TPA (100 ng/mL) for the times indicated. The presence of Elk-1 and phospho-Elk-1 at the c-fos promoter was then determined by ChIP as described. PCRs containing primer pairs amplifying a promoter region containing the SRE (upper panel) or a region of the gene (lower panel) were performed after immunoprecipitation of DNA/promoter complexes with the Elk-1 antibodies indicated in Subheading 2.2

quantification of band intensities within a limited linear range (see Note 5).

4 Notes 1. It is advisable to establish sonication conditions for each cell type used (the DNA fragment size is influenced by cell type, cell number, volume and sonication power). To check the sheared DNA size, dilute 10–20 mL aliquots of sheared soluble chromatin to 50 mL with Dilution buffer, add 2 mL 5 M NaCl and heat at 65°C for 6 h to reverse cross-links. Mix samples with 6 × Gel loading buffer and load onto a 2% Agarose gel together with a suitable DNA molecular weight ladder. Visualise sheared DNA by staining with Ethidium Bromide. 2. DNA from 2 to 3 × 106 cells is needed to perform each ChIP reaction. 3. In some cases the chromatin requires pre-cleaning. After diluting the chromatin to 1 mL with Dilution buffer, add 2 mg of sheared salmon sperm DNA and protein G sepharose beads and incubate for 2 h at 4°C on wheel. Sediment the beads by centrifugation, transfer the chromatin supernatant to fresh tubes and then add the antibody for the immunoprecipitation step. 4. The Gel loading buffer lacks bromophenol blue (BPB) because in the gel system used it would migrate to the same position as the PCR product and reduce the fluorescent signal.

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Fig. 2 Densitometric quantification of PCR efficiency. Serial dilutions of DNA extracted from sonicated chromatin (4, 16, 64-fold, as indicated), were amplified in standard PCRs with the c-fos promoter primer pair (n = 3). After 34 cycles, samples were back-diluted to give arithmetically equal amounts of amplicon from each input dilution and analysed by gel electrophoresis. Band intensities were quantified by densitometric analysis of digital gel images

5. To determine the relationship between target DNA and amplicon yield, serial dilutions of DNA extracted from sonicated chromatin (input fraction) should be assayed with each primer pair in standard PCRs. After amplification for a set number of cycles, the PCR products are inversely diluted with respect to the initial serial dilution of input DNA to obtain arithmetically equal amounts of amplicon, which should yield bands of equal intensity upon gel electrophoresis. In this way, limitations associated with DNA staining and the restricted linear range of the imaging device can be circumvented. As shown in Fig. 2, each fourfold reduction in input DNA gave rise to a corresponding decrease in amplification product, demonstrating a linear response to target DNA concentration across a 64-fold range. Alternative methods for relative and absolute quantification of input DNA by PCR include the introduction of radionucleotide triphosphates in the amplification reactions or the development of fluorescent probes for real time quantitative PCR [11].

Acknowledgements We thank Wendy Solis for secretarial assistance. This work was supported by grants to PES from the BBSRC (refs. C17917 and C19734).

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References 1. Sharrocks AD (2001) The ETS-domain transcription factor family. Nat Rev Mol Cell Biol 2:827–837 2. Shaw PE, Saxton J (2003) Ternary complex factors: prime nuclear targets for mitogenactivated protein kinases. Int J Biochem Cell Biol 35:1210–1226 3. Buchwalter G, Gross C, Wasylyk B (2004) Ets ternary complex transcription factors. Gene 324:1–14 4. Gille H, Sharrocks AD, Shaw PE (1992) Phosphorylation of transcription factor p62TCF by MAP kinase stimulates ternary complex formation at c-fos promoter. Nature 358:414–417 5. Gille H, Kortenjann M, Thomae O, Moomaw C, Slaughter C, Cobb MH, Shaw PE (1995) ERK phosphorylation potentiates ELK-1-mediated ternary complex formation and transactivation. EMBO J 14: 951–962 6. Janknecht R, Ernst WH, Pingoud V, Nordheim A (1993) Activation of ternary

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complex factor ELK-1 by MAP kinases. EMBO J 12:5097–5104 Marais R, Wynne J, Treisman R (1993) The SRF accessory protein Elk-1 contains a growth factor- regulated transcriptional activation domain. Cell 73:381–393 Kouzarides T (2007) Chromatin modifications and their function. Cell 128:693–705 Kuo MH, Brownell JE, Sobel RE, Ranalli TA, Cook RG, Edmondson DG, Roth SY, Allis CD (1996) Transcription-linked acetylation by Gcn5p of histones H3 and H4 at specific lysines. Nature 383:269–272 Zhang HM, Li L, Papadopoulou N, Hodgson G, Evans E, Galbraith M, Dear M, Vougier S, Saxton J, Shaw PE (2008) Mitogen-induced recruitment of ERK and MSK to SRE promoter complexes by ternary complex factor Elk-1. Nucleic Acids Res 36:2594–2607 VanGuilder HD, Vrana KE, Freeman WM (2008) Twenty-five years of quantitative PCR for gene expression analysis. Biotechniques 44:619–626

Chapter 18 Integration of Protein Kinases into Transcription Complexes: Identifying Components of Immobilised In Vitro Pre-initiation Complexes Hong-Mei Zhang, Stéphanie Vougier, Glenn Hodgson, and Peter E. Shaw Abstract Regulation of gene expression is essential for coordinated cell growth and development. The de-regulation of certain genes is also recognised to contribute to both heritable and acquired disease. Transcription factors influence the assembly and activity of transcription complexes, which they achieve in part by recruiting co-activators to gene promoters to participate in the dynamic cycle of polymerase binding, initiation and escape from the promoter. Co-activator recruitment and accompanying post-translational modifications to components of promoter complexes appear to differ between genes and as a consequence of varying signal input. Thus a full understanding of transcriptional initiation and control will ultimately require the elucidation of these processes. The method described here was designed to detect the presence of proteins and post-translational modifications in complexes formed in vitro on gene-specific promoters. It has been used, among other things, to detect the recruitment of the Mitogen-Activated Protein (MAP) kinases ERK1 and ERK2 to the promoters of mitogen-responsive genes. Key words: Mitogens, MAP kinase, DNA binding, Biotin–streptavidin, Magnetic beads, Protein mass spectrometry, Immunoblotting

1. Introduction The coordinated expression of around 25,000 protein-coding genes in the human genome underlies the diversity of cells and the complexity of form readily apparent in our own bodies. Regulation occurs primarily at the point of initiation of transcription by RNA polymerase II (RNAP), an event requiring a multitude of basal transcription factors and RNAP-associated co-activators (1). Transcription is both preceded and accompanied by chromatin rearrangements that allow transcription factors to access gene promoters and direct the formation of pre-initiation complexes Paul J. Higgins (ed.), Transcription Factors: Methods and Protocols, Methods in Molecular Biology, vol. 647, DOI 10.1007/978-1-60761-738-9_18, © Springer Science+Business Media, LLC 2010

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(PICs), as well as facilitating the progression of RNAP along genes (2). In the case of many transcription factors involved in acute gene regulation, phosphorylation has long been recognised to have a profound effect on their activity (3). Protein phosphorylation also plays a critical role in at least one other level of transcriptional control. Several basal transcription factors and co-activators present in PICs have associated protein kinases, including CDK7/cyclin H present with MAT1 in TFIIH, CDK8/cyclin C present with MED230/240 in the negative regulatory ARC/Mediator-like complex, and Cdk9/cyclin T/K, part of the P-TEFb (positive transcription elongation factor) complex. Their principal role appears to be controlling the phosphorylation status of the RNAP carboxy-terminal domain (CTD), which oscillates between phosphorylation of serine 2 and serine 5 during the transcription cycle (reviewed in (4)). Several reports have described the association of yeast MAPKs with specific gene promoters (5–8). Furthermore, human p38a was recently shown to occupy gene promoters during myogenesis (9). Similarly, ERK and MSK (Mitogen and Stress-Activated Kinase) were found in complexes with the progesterone receptor on the MMTV promoter (10) and with Elk-1 on the mitogenresponsive c-fos and egr1 promoters (11). These findings are consistent with the proposal that MAPKs and other acutely regulated kinases may be frequent occupants of signal-dependent gene promoters (8, 12) and imply that they serve additional roles during transcriptional activation beyond the phosphorylation of target transcription factors (13–15). A number of proteins recruited into PICs have been detected in transcription complexes assembled in vitro on immobilised DNA promoter templates (16, 17). This precept indicates that the analysis of PICs isolated on specific gene promoter templates is likely to reveal participating protein kinases when and where they are present. Screening isolated PICs for the presence of protein kinases can take an unbiased approach, involving mass spectrometry (MS) and peptide mass fingerprinting (or MS-MS peptide sequencing). Alternatively, a predictive approach can be taken, involving the separation of PIC components by SDS-PAGE and immunoblotting with antibodies to screen for individual protein kinases. Here we describe the latter, although the protocol is also suitable for the isolation of PICs for MS analysis.

2. Materials 2.1. Cell Culture, Mitogen Stimulation and Nuclear Extract Preparation

1. MEM-Joklik (Biochrom AG), supplemented with 5% new born calf serum (NBCS) (Biochrom AG), 5% foetal calf serum (FCS) (Autogen Bioclear), 100 U/mL penicillin and 100 µg/ mL streptomycin (Sigma).

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2. Dulbecco’s Modified Eagle’s Medium (DMEM) (Biochrom AG) supplemented with 10% FCS, 2 mM L-glutamine (Biochrom AG), 100 U/mL penicillin and 100 µg/mL streptomycin. 3. Glass flasks (Schott Duran), stir bars and magnetic stirrers (Ikamag REO), 37°C incubator (Sanyo MIR-253). 4. Cooled centrifuge with 6 × 1 L buckets (Sorvall RC3C Plus) and at least 6 × 1 L plastic bottles (Nalgene). 5. Phosphate Buffered Saline (PBS), containing freshly added 2 mM Na3VO4, 10 mM NaF and 0.5 mM benzamidine. 6. Cooled centrifuge (8 × 50 mL rotor) (Heraeus, Megafuge 1.0R or similar). 7. Vortex mixer (Vortex Genie 2, Scientific Industries). 8. Buffer A: 10 mM HEPES pH 7.9, 1.5 mM MgCl2 and 10 mM KCl, containing freshly added 2 mM Na3VO4, 10 mM NaF, 20 mM b-glycerophosphate, 10 mM p-nitrophenyl phosphate (pNPP) and 0.5 mM dithiothreitol (DTT). 9. Glass Dounces (40 and 15 mL chambers with type B pestles) (Kontes Glass Co.). 10. Buffer C: 20 mM HEPES pH 7.9, 25% Glycerol, 0.42 M NaCl, 1.5 mM MgCl2 and 0.2 mM EDTA, containing freshly added 2 mM Na3VO4, 10 mM NaF, 20 mM b-glycerophosphate, 10 mM pNPP and 0.5 mM DTT. 11. Buffer D: 20 mM HEPES pH 7.9, 20% Glycerol, 20 mM KCl, 1.5 mM MgCl2 and 0.2 mM EDTA, containing freshly added 2 mM Na3VO4, 10 mM NaF, 0.5 mM phenylmethanesulphonyl fluoride (PMSF), 0.5 mM benzamidine and 0.5 mM DTT. 12. High speed centrifuge (Beckman Coulter Avanti J26 XP) with 8 × 50 mL rotor (JA25.50). 13. Screw cap centrifuge tubes (50 mL) (Nalgene). 14. Dialysis tubing (MW cut off 3.5 kDa) (Spectrum Labs). 15. Cooled bench-top microfuge (Eppendorf, model 5417R). 16. Lowry protein concentration determination kit (BioRad, DC Protein Assay). 2.2. Preparation of Immobilised Templates

1. A promoter-specific PCR primer pair. The upstream primer (with respect to the promoter) has a biotin moiety coupled to the 5¢ end for subsequent binding to streptavidin beads. 2. DNA source from which the required promoter sequence can be amplified. 3. Thermocycler (Biometra, TRIO-Thermoblock). 4. DNA purification (QIA-quick PCR purification, Qiagen).

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5. Streptavidin-coated magnetic beads (Dynal, Dynabeads M-280 Streptavidin). 6. Magnet holder for handling beads (Dynal). 7. 1× Binding buffer: 1 M NaCl, 10 mM Tris pH 7.4, 0.2 mM EDTA. 8. Vertical wheel mixer (Stuart SB2 Rotator). 9. Heated shaker (Eppendorf Thermomixer compact). 10. Transcription buffer: 12 mM HEPES pH 8.0, 12% Glycerol, 60 mM KCl, 120 mM EDTA pH 8.0, 7.5 mM MgCl2, 1 mM DTT, 0.5 mM PMSF. 11. Transcription buffer containing 0.05% Nonidet P-40 (NP-40). 2.3. Assembly of Pre-initiation Complexes and Elution

1. Salmon sperm DNA (Sigma) sheared (1 mg/mL in TE). 2. Poly(dIdC) (1 mg/mL in TE) (GE-Healthcare). 3. MgCl2 (0.1 M). 4. Nuclear extract in D buffer (protein concentration ~10 mg/ mL). 5. Vertical wheel mixer (Stuart SB2 Rotator). 6. Heated shaker (Eppendorf Thermomixer compact). 7. Laemmli loading buffer (6×). 8. Restriction Enzyme XhoI, 20 U/µL (NE Biolabs).

2.4. SDS-PAGE and Immunoblotting

1. Acrylamide solution, 30% (29:1 acrylamide:bisacrylamide, Sigma). 2. Resolving (lower) buffer: 1.5 M Tris–HCl pH 8.8 containing 0.4% (w/v) sodium dodecylsulphate (SDS). 3. Stacking (upper) buffer: 0.5 M Tris–HCl pH 6.8 containing 0.4% (w/v) SDS. 4. Ammonium persulphate (APS) as a 10% solution in ddH2O (store cold for up to 1 week). 5. N,N,N,N¢-tetramethyl-ethylenediamine (TEMED) (BioRad). 6. Water-saturated isobutanol. 7. SDS-PAGE buffer: 25 mM Tris, 192 mM glycine, 0.1% SDS. 8. Pre-stained protein molecular weight markers (Page Ruler, Fermentas). 9. Transfer buffer: 25 mM Tris, 192 mM glycine, 20% (v/v) methanol. 10. Chromatography paper 3MM (Whatman). 11. Polyvinylidine fluoride (PVDF) membrane (Westran S, Whatman).

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12. Tris-buffered saline (10× TBS): 100 mM Tris–HCl pH 8.0, 1.5 M NaCl. 13. TBS-T: 1× TBS supplemented with 0.1% (v/v) Tween 20. 14. TBS-TM: TBS-T supplemented with 5% (w/v) instant dried skimmed milk powder. 15. Shallow glass or plastic dishes (~100 × 80 × 20 mm) and 50 mL Falcon tubes. 16. Flat bed shaker (GFL 3017 or similar). 17. Primary antibodies. 18. Roller bars (Stuart, Roller Mixer SRT1). 19. Secondary antibodies conjugated to horseradish peroxidase. 20. Enhanced chemiluminescence (ECL) reagents (GE-Healthcare). 21. Digital fluorescence imaging system (Fujifilm LAS3000).

3. Methods 3.1. Mitogen Stimulation and Nuclear Extract Preparation

1. For medium scale nuclear extract preparation, HeLa cells are grown as suspension cultures (up to 6 L) in MEM-Joklik medium supplemented with 5% NBCS and 5% FCS in 10-L, flat-bottomed, round glass flasks with magnetic stir bars. The flasks are placed on magnetic stirrers (heat insulated) in a 37°C cabinet. A 6-L culture yields about 7 × 109 cells and ~10 mL of nuclear extract. For smaller scale extracts, cells can be grown on plastic or in roller bottles in DMEM supplemented with 10% FCS. 2. To bring HeLa cells in suspension culture to quiescence, collect cells by centrifugation (Sorvall RC3C Plus) at 1,200 rpm (420 × g) for 20 min at RT, re-suspend the cell pellet gently in 1/10 of the original volume of starvation medium (MEMJoklik supplemented with 0.5% NBCS), dilute cells to 6 × 105 cells/mL and return to culture for 36–48 h. For cells on plastic or in roller bottles, replace the medium with DMEM supplemented with 0.5% FCS. 3. Just prior to mitogen stimulation, the cell number should be counted (haemocytometer) for future reference (step 9). To stimulate cells, pre-warmed FCS (to 15% of final volume) is added to the culture. 4. Suspension cultures are harvested 5–8 min after FCS addition by decanting the culture into pre-cooled centrifuge bottles placed in iced water and centrifugation at 1,200 rpm (420 × g) for 20 min at 4°C. Cells grown in plates or bottles are harvested after 15 min as they can be cooled rapidly by removing medium and immediately adding ice cold PBS.

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5. Decant the medium from the bottles (remove the last few drops with a pipette) and re-suspend each cell pellet (by swirling) in 5 mL ice cold PBS (+ inhibitors). Pool the cells in 50 mL Falcon tubes on ice. Rinse each bottle with a further 5 mL of ice cold PBS and add to pooled cells on ice. 6. Collect the cells in a cooled centrifuge (Megafuge 1.0R) at 2,000 rpm (690 × g) for 10 min at 4°C. Decant the PBS, resuspend each pellet in 5 mL fresh PBS (+ inhibitors) and combine all the cells into a single Falcon tube. Centrifuge the cells again at 2,000 rpm (690 × g) for 10 min at 4°C, pour off the supernatant and record the packed cell volume (PCV) and pellet weight (optional). 7. Add 5 × PCV of complete buffer A to the cells and re-suspend them with ten short pulses on a vortex mixer. Incubate the cells on ice for 10 min. Mix the cells again (ten short pulses) and centrifuge at 2,000 rpm (690 × g) for 10 min at 4°C. The cell pellet should have swollen about twofold. Discard the supernatant and re-suspend the cells in 2 × PCV (recorded earlier) of complete buffer A. 8. Transfer the cells to a cooled glass Dounce (40 mL chamber, type B pestle), placing a droplet (~100 µL) of the cell suspension onto a microscope slide. Wearing protective gloves hold the chamber tightly in ice and homogenise the cells with ten up and down strokes of the pestle. Place a droplet of the homogenate onto the glass slide and check for lysis under a microscope. More than 90% of the cells should be broken open. If not, further strokes of the pestle should be applied. Collect the nuclei by centrifugation at 2,500 rpm (1,080 × g) for 10 min at 4°C. 9. Carefully pipette the supernatant into a fresh tube (on ice) (see Note 1). To the nuclear pellet add 1 mL of complete buffer C per 109 cells, as recorded from the count in step 3, and quickly re-suspend the nuclei. Transfer to a cooled glass Dounce (15 mL chamber, type B pestle), again placing a droplet (~100 µL) of the suspension onto a microscope slide. Hold the chamber tightly in ice (protective gloves) and homogenise with ten hard, up and down strokes of the pestle. Place a droplet of the homogenate onto the glass slide and check for disruption under a microscope. Fewer than 10% of the nuclei should remain intact. 10. Pour the homogenate into a cooled 25/50 mL beaker with stir bar, cover with aluminium foil or plastic film and place in ice on a magnetic stirrer. Allow the homogenate to stir for 30 min. Transfer the homogenate into a 50 mL screw cap centrifuge tube and centrifuge (Beckman Coulter) at 15,000 rpm (27,200 × g) for 30 min at 4°C.

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11. Cut a piece of prepared dialysis tubing of sufficient length and rinse with ddH2O and then with D buffer (see Note 2). Clip or tie off one end and pipette the supernatant post centrifugation into the tube. Clip or tie off the other end such that the tubing is just taut. Dialyse the nuclear extract twice against 100 volumes of cooled D buffer for 90 min at 4°C (3 h in total). 12. Post-dialysis, unclip one end of the tubing and, with a pipette, transfer the extract into 1.5 mL microfuge tubes (the tubing can be progressively cut down with scissors to facilitate this step). Centrifuge in a cooled bench-top centrifuge at 14,000 rpm (20,800 × g) for 15 min at 4°C. Transfer supernatants into fresh 1.5 mL microfuge tubes (1 × 10 µL, 5 × 100 µL and 500 µL aliquots), snap freeze in liquid nitrogen and store at −80°C. The 10 µL aliquot is intended for measuring the protein concentration of the extract, which should lie in the range of 9–12 mg/mL. 3.2. Preparation of Immobilised Templates

1. The biotinylated DNA templates are prepared by PCR amplification and should be checked for specificity and homogeneity by agarose gel electrophoresis. 2. Purification of the PCR product(s) is carried out with Qspin columns (Qiagen). The DNA concentration is measured on a spectrophotometer (Nanodrop). The template is then ready for immobilisation on streptavidin-coated magnetic beads (see Note 3). 3. Pre-wash 300 µg beads (sufficient for ten PIC assembly reactions) in 500 µL binding buffer twice for 15 min at RT on a vertical wheel. Remove the binding buffer. 4. Add biotinylated promoter fragment (20–25 pmol in TE) to the beads; add 1/4 volume of 5 M NaCl (to 1 M NaCl) and then binding buffer to a final volume of 60 µL. Re-suspend beads and incubate for 1 h at RT in a heated shaker (350 rpm). Remove tubes occasionally and re-suspend any sedimented beads by flicking tube with index finger. 5. After incubation, remove the supernatant and run a 10 µL sample on a small, 1.5% agarose gel, loading an equivalent amount of pre-bead DNA solution as a control. More than 75% of the DNA should have bound to the beads. 6. Wash the beads (+ DNA) (immobilised templates) twice in 500 µL binding buffer for 15 min at RT on the vertical wheel. 7. Wash the beads (+ DNA) twice in 500 µL transcription buffer for 15 min at RT on the vertical wheel. After washing twice in transcription buffer, re-suspend the beads + DNA in 100 µL transcription buffer. The immobilised templates, sufficient for ten reactions, can be used directly for PIC assembly or frozen and stored at −20°C.

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3.3. Pre-initiation Complex Assembly and Elution

1. For each reaction, place a 1.5 mL microfuge tube on ice and add 300–360 µg HeLa nuclear extract (protein concentration ~10 mg/mL), 5 µg salmon sperm DNA, and 3 µg poly(dIdC). Add 0.1 M MgCl2 to a final concentration of 5 mM and transcription buffer to a final volume of 100 µL. Mix gently and pre-incubate on ice for 15 min. 2. To each aliquot of HeLa nuclear extract, add 10 µL of immobilised template (from step 7 in Subheading 3.2), mix gently and incubate at 30°C for 45 min with gentle shaking (heated shaker, 350 rpm). 3. Insert each reaction tube into magnetic holder, allow beads to collect at the magnet and transfer the nuclear extract to a fresh tube on ice (see Note 4). Wash the beads (immobilised PICs) three times in 500 µL transcription buffer containing 0.05% NP-40. The NP-40 serves to hinder aggregation of the beads during the washing stages. Remove the final wash (see Note 5). 4a. Two strategies are described for the elution of proteins in preparation for immunoblotting. For high salt elution, add 30 µL 1 M NaCl to the beads, mix and incubate at 30°C for 15 min with gentle shaking (heated shaker, 350 rpm). Insert each reaction tube into the magnetic holder, allow beads to collect at the magnet and transfer the eluate to a fresh tube on ice. Add 6 µL of 6× Laemmli loading buffer, mix and boil samples for 3 min. Complexes can also be eluted with successive steps of increasing NaCl concentration, as indicated by the overview presented in Fig. 1.

Fig. 1. Schematic diagram of PIC assembly and analysis. (1) Biotinylated promoter templates are immobilised on streptavidin-coated magnetic beads. (2) Promoter templates are incubated in nuclear extract under conditions that allow complex formation. (3) Pre-initiation complexes are collected on beads and washed to remove non-specific proteins. (4) Complexes are eluted from beads with successive NaCl steps and fractions are separated by SDS-PAGE.

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4b. As an alternative to step 4a PICs can be separated from the magnetic beads by restriction endonuclease (RE) cleavage (16), provided that a suitable site has been engineered into the immobilised template. An assessment of several REs indicated that XhoI is able to cleave efficiently in transcription buffer, thus obviating the need to change buffers and potentially disrupt the PICs. For RE cleavage, add 30 µL Transcription buffer (without NP-40) containing 50 U of XhoI and incubate at 30°C for 1 h with gentle shaking (heated shaker, 350 rpm). Insert each reaction tube into the magnetic holder, allow beads to collect at the magnet and transfer the eluate to a fresh tube on ice. This strategy releases considerably less protein from the beads and may thus reduce the level of non-specific contaminants, but it also leaves a significant residue of specific proteins at the beads, most likely as a consequence of incomplete digestion. These can be released by subsequent elution with 1 M NaCl, as described in step 4a. To the eluate(s), add 6 µL of 6× Laemmli loading buffer, mix and boil samples for 3 min and allow to cool to RT. The samples can be used directly or stored at −20°C. 3.4. SDS-PAGE and Immunoblotting

There are many descriptions of standard SDS-PAGE and electrotransfer of proteins onto membranes. This protocol assumes the use of the BioRad Mini-protean II and Trans-Blot Semi-Dry systems, but alternatives exist and can be used if preferred. 1. Insert clean glass plates (1 mm spacers) into gel assembly and tighten screws, ensuring plates and spacers are flush at the bottom. Insert into pouring stand. For a 10% resolving gel, mix 2 mL 30% acrylamide, 1.5 mL lower buffer, 2.5 mL ddH2O. Add 25 µL APS and 15 µL TEMED, mix and fill the gel mould to a depth of 55 mm. Overlay resolving gel with 1 mL isobutanol and allow to polymerise (ca. 20 min). 2. For the stacking gel, mix 0.5 mL 30% acrylamide, 0.94 mL upper buffer, 2.3 mL ddH2O. Pour isobutanol off the resolving gel and then rinse mould thoroughly with ddH2O. To the stacking gel mix add 10 µL APS and 10 µL TEMED, mix and fill the gel mould, insert comb and allow stacking gel to polymerise (ca. 20 min). 3. Assemble gel in electrophoresis tank, add SDS-PAGE buffer, remove comb and load samples (including MW ladder). Electrophorese at 120 V until dye front has entered resolving gel, then at 180 V until front reaches bottom of gel. 4. Disassemble the gel, gently prise one glass plate off the gel and use one edge of the free plate to separate the stacking gel from the resolving gel. Soak the resolving gel in transfer buffer for 20 min at RT. For each gel cut one piece of PVDF

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membrane and six pieces of 3MM paper the size of the gel (80 × 55 mm). Wet the PVDF membrane in 100% methanol and then soak the membrane and 3MM paper in transfer buffer. 5. For each gel, pile three sheets of wetted 3MM paper onto the lower plate (anode) of the semi-dry transfer cell. Avoid trapping air bubbles between the layers. Lay a piece of PVDF membrane on the pile of 3MM paper and lay the gel on the membrane (this order is critical!). Then lay another three pieces of wetted 3MM paper onto the gel, avoiding air bubbles. Lower the cathode plate gently onto the pile(s) and ensure it clips into place. Fit the cover, and at a potential of 30 min) may cause cells to aggregate and not sonicate efficiently. 3. Quench the cross-link reaction by direct addition of 10 ml 0.125 M Glycin and incubate for 10–15 min at RT, under gently rotation (rotating-platform). 4. Wash the cells twice with 10 ml of cold PBS, scrap the cells in 3 ml of cold PBS to allow complete recovery, and collect them in two 2-ml Eppendorf tubes. Spin the cells 5 min at 2,000 × g – 4°C, and resuspend each cell pellet with 250 ml of cold PBS. Pool the samples. You should end with only one Eppendorf tube containing 500 ml of cross-linked cells.

3.6.2. Chromatin Preparation (Day 1)

Chromatin is recovered following several steps of washingresuspension. 1. Spin the cells 5 min at 2,000 × g – 4°C (see Note 4). 2. Resuspend and wash the cells with 1 ml of cold PBS. 3. Spin the cells 5 min at 2,000 × g – 4°C, discard the supernatant. 4. Resuspend and wash the cells with 1 ml of cooled NCP buffer 1. 5. Spin the cells 5 min at 2,000 × g – 4°C, discard the supernatant. 6. Resuspend and wash the cells with 1 ml of cooled NCP buffer 2. 7. Spin the cells 5 min at 2,000 × g – 4°C, discard the supernatant. 8. Resuspend the cells in 1 ml Lysis Buffer at RT.

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9. Sonicate for 15 s at power 3 and 60% output, with 1-min refractory period between the three sonications. Keep the lysate on ice during this time, but remember the presence of SDS. So don’t leave the lysis more than 2 min on ice. 10. Spin 10 min at 10,000 × g – RT. 11. Remove 100 ml of lysate (Input), and store at −20°C until proceeded to the reverse cross-linking step. The immunoprecipitation will be performed on the remaining 900 ml. (see Notes 5 and 6). 3.6.3. Immunoprecipitation (Day 1)

1. Bead preparation: Prepare 500 ml of 50% protein A/G Sepharose Beads slurry. Wash three-times the beads with 1 ml cold PBS, twice with 1 ml IP-, and resuspend in 500 ml IP-buffer (see Note 7). 2. Split in two the 900 ml of chromatin suspension. To each 450 ml of chromatin suspension, add 900 ml of IP Buffer, and proceed to the preclearing step by adding 10 mg of ssDNA (salmon sperm DNA 1 mg/ml)) and 100 ml of the 50% protein A/G Sepharose Beads slurry. Incubate the two 2 mlEppendorf tubes 3–4 h at 4°C under rocking. 3. Pellet the unspecific protein A/G Sepharose Beads complexes by centrifugation at 800 × g for 1 min at 4°C. Recover the supernatant (1,400 ml per 2 ml-Eppendorf tube). 4. To proceed to the Immunoprecipitation: incubate under rocking overnight at 4°C, 700 ml of supernatant with each specific antibody. In our case, 15 ml of anti-USF1, 15 ml of anti-USF2, 15 ml anti-Tbx2, and 15 ml of Rabbit IgG, these later unrelated antibodies serve to see background pull-downs (Fig. 2).

3.6.4. Collection of Complexes (Day 2)

1. Add 2 mg of ssDNA and 40 ml of the 50% protein A/G Sepharose Beads slurry to each immuno-precipitated samples. 2. Incubate 2–3 h at 4°C under rocking. 3. Pellet the specific complexes by centrifugation at 800 × g for 1 min at 4°C, and remove the supernatant. 4. Wash seven-times, and spin successively the specific complexes with 300 ml of Washing Buffer 1, Washing Buffer 2, Washing Buffer 3, and finally three-times with Washing Buffer 4. Perform the seven spins at 800 × g for 1 min and 4°C, and discard the supernatant. 5. To recover the specific DNA/protein complexes from the beads: Extract the complexes by adding 50 ml of Extraction Buffer to bead-pellets, shake 10 min (vortex-vibrax: 1250g). Spin (800 × g – 5 min – 4°C). Remove the supernatant and store it on ice. Repeat twice the extraction step. You should end with 150 ml of extract protein/DNA complexes.

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Fig. 2. USFs supershift assays. Supershift assay determine which b-HLH-LZ proteins are responsible for the observed shifted bands. In this assay, USF1 and USF2 antibodies are able to shift alone nearly completely the protein/DNA complexes (lanes 2 and 3: a-USF1 and a-USF2). Together (lane 4: a-USF1/a-USF2), they shift completely the protein/DNA complexes. The mobility of the protein/DNA complexes is unchanged in the presence of an antibody raised against a transcription factor that does not belong to the bHLH-LZ family (anti-Tbx2), or with rabbit immunoglobulin (IgG). Taken together, these data clearly show that under the experimental conditions used, this E-box sequence binds specifically USF1 and USF2 b-HLH-LZ members.

6. To reverse the crosslinking of each protein/DNA complexes (Input 100 ml and supernantants150ml): incubate the samples overnight at 65°C, under gently agitation (500 rpm). Make a hole in the lid of the Eppendorf tube, using a syringe, to help evaporation and denaturation of the formaldehyde. 3.6.5. DNA Preparation and PCR (Day 3)

1. Recover DNA from the 150 ml reversed crosslink samples by using NucleoSpin Extract® II, columns (Macherey Nagel). Elute DNA in 50 ml H2O. 2. Standard and real time PCR assays are performed to measure genomic DNA promoter sequence enrichment present in the DNA extract within the ChIP samples. We initially used standard PCR to analyze in vivo promoter occupancy (Fig. 3a), and now we tend to use Real-Time PCR (Fig. 3b), which allows us to quantify sequence fold enrichment of the studied promoter. Sequence enrichment will be relative to IgG ChIPcontrol and to control-region amplification. For control-region amplification, we initially used an unrelated promoter region (the Hsp70 promoter region; Galibert et al. 2001; (13)), and now with real time PCR, we use distal regions of the studied promoter. In this case, the control-sequence should be about 2–3 kb from the E-box target promoter sequence (although it depends on sonication efficiency). Control-region amplification should also not contain conserved E-box motif.

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Fig. 3. USFs ChIP analysis. An example of typical results obtained with USFs antibodies in ChIP assays. Immunoprecipitated DNA was amplified by standard PCR (a) or by real time PCR (b). Accuracy of the PCR is given by the amplification of a specific single product, corresponding to the target gene in the Input and in the a-USF ChIP lanes, while no amplification is observed in the control ones (beads, IgG), and amplification of a control region (the Hsp70 gene (a) in this example) is limited to the input. In case of real time PCR fold enrichment is calculated, by normalization with a control region and a control ChIP.

3. At least two sets of primers should be designed; one centered on the target E-box sequence and the other amplifying a control-region. Sequence amplification should be of 100–150 nucleotides long. 4. Primer should be designed carefully and validated. Generally we use a length of about 20-bases, containing 40–60% GC. Avoid sequences that might produce internal secondary structure, avoid three G or C nucleotides in a row near the 3¢-end of the primer (nonspecific primer annealing), 3¢-ends of the primers should not be complementary (production of primers dimers). For real time PCR, Tm should be between 58 and 60°C (ideally both primers should have nearly identical melting temperature). Several primer-design programs can be

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used with comparable results; among them, we frequently use Primer3 (http://frodo.wi.mit.edu/). Primers efficiency needs to be checked and specific amplification assessed by the dissociation curve profile, in the case of real time PCR. 5. Real Time PCR reaction is obtained by mixing 3 pmoles of each designed Forward and Reverse primers (0.3 ml of the 10 mM stock), with 5 ml of Power SYBR® Green, PCR Master Mix (Applied Biosystem) to 2 ml of extracted DNA-ChIP, or Input DNA samples. Complete with H2O to a final volume of 10 ml-program. 6. Sequence fold enrichment will be calculated according to the DDCt method (http://www3.appliedbiosystems.com/ AB_Home/applicationstechnologies/Real-TimePCR/ AbsolutevsRelativeQuantitation/index.htm). Fold Induction = 2(DCt1−DCt2), where: DCt1 = (Ct Target sequence−Ct Control Region) Specific ChIP in our case USFs antibodies and: DCt2 = (Ct Target sequence−Ct Control Region) Control ChIP in our case IgG or unrelated antibody (anti-Tbx2).

4. Notes 1. Cell Lysis can be checked under the microscope by the addition of the Trypan Blue solution to an aliquot of cells. The dye is excluded from intact cells, but stains the nuclei of lysed cells. When trained you do not need to use Trypan Blue anymore. 2. It is well advised to prepare the gels on Day 1 and to perform the assay on Day 2. In this case to avoid that the acrylamide-gel dries, humidify the upper part with 0.5× TBE soaked-tissue, wrap in saran, and place in the cold room. On Day 2, remove the comb; complete the assembly of the gel unit; add running buffer in the upper and lower tanks; wash the wells with running buffer (0.5× TBE), and remove the bubbles that would interfere with the migration. 3. Each set of binding assays must include a control in which no protein is added (free-probe only). 4. Samples can be stored at this step for at least 3 days at −80°C; in this case, wash with cold PBS the cells previous follow up. 5. Sonication is used to disrupt the cross-linked chromatin fibers. It is crucial to generate the appropriate length of chromatin fragments as it can greatly affect the read-out obtained at the end of the procedure. A length of 1,000 bp is accepted as optimal. The sonication step has to be set up according to the

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sonicator model present in your lab. The present settings are to be used with a Sonifier Cell Disruptor II W450 from Branson, with a 3 mm microtip probe. 6. To check the efficiency of sonication, take 15 ml of the lysate. Load the sample onto a 1% agarose gel and visualize by Ethidium Bromide staining. Repeat these latter steps until the desired fragment size is obtained. 7. Bead preparation should be performed the previous day and stored at 4°C.

Acknowledgment This work was supported over the years by the LNCC – “Comités Départementaux du Grand Ouest” and the ARC cancer care fundings. We would also like to thank the CNRS, University of Rennes and Brittany Region for their support. References 1. Juven-Gershon T, Hsu JY, Theisen JW, Kadonaga JT (2008) The RNA polymerase II core promoter – the gateway to transcription. Curr Opin Cell Biol 20:253–259 2. Jones FS, Crossin KL, Cunningham BA, Edelman GM (1990) Identification and characterization of the promoter for the cytotactin gene. Proc Natl Acad Sci U S A 87:6497–6501 3. Blackwell TK, Huang J, Ma A et al (1993) Binding of myc proteins to canonical and noncanonical DNA sequences. Mol Cell Biol 13:5216–5224 4. Blackwell TK, Weintraub H (1990) Differences and similarities in DNA-binding preferences of MyoD and E2A protein complexes revealed by binding site selection. Science 250:1104–1110 5. Baxevanis AD, Vinson CR (1993) Interactions of coiled coils in transcription factors: where is the specificity? Curr Opin Genet Dev 3:278–285 6. Aksan I, Goding CR (1998) Targeting the microphthalmia basic helix-loop-helix-leucine zipper transcription factor to a subset of E-box elements in vitro and in vivo. Mol Cell Biol 18:6930–6938 7. Halazonetis TD, Kandil AN (1991) Determination of the c-MYC DNA-binding site. Proc Natl Acad Sci U S A 88:6162–6166 8. Galibert MD, Miyagoe Y, Meo T (1993) E-box activator of the C4 promoter is related to but distinct from the transcription factor upstream stimulating factor. J Immunol 151:6099–6109

9. Sirito M, Lin Q, Maity T, Sawadogo M (1994) Ubiquitous expression of the 43- and 44-kDa forms of transcription factor USF in mammalian cells. Nucleic Acids Res 22:427–433 10. Sirito M, Walker S, Lin Q, Kozlowski MT, Klein WH, Sawadogo M (1992) Members of the USF family of helix-loop-helix proteins bind DNA as homo- as well as heterodimers. Gene Expr 2:231–240 11. Corre S, Galibert MD (2005) Upstream stimulating factors: highly versatile stress-responsive transcription factors. Pigment Cell Res 18: 337–348 12. Braunstein M, Rose AB, Holmes SG, Allis CD, Broach JR (1993) Transcriptional silencing in yeast is associated with reduced nucleosome acetylation. Genes Dev 7:592–604 13. Corre S, Primot A, Sviderskaya E et al (2004) UV-induced expression of key component of the tanning process, the POMC and MC1R genes, is dependent on the p-38 activated upstream stimulating factor-1 (USF-1). J Biol Chem 279:51226–51233 14. Metivier R, Gallais R, Tiffoche C et al (2008) Cyclical DNA methylation of a transcriptionally active promoter. Nature 452:45–50 15. Galibert MD, Carreira S, Goding CR (2001) The Usf-1 transcription factor is a novel target for the stress-responsive p38 kinase and mediates UV-induced Tyrosinase expression. EMBO J 20:5022–5031

INDEX A Acetylation acetyltransferase assay ...............................220, 225–226 in vitro acetylation assay .................................. 220, 226 Activation transcription factor 3 (ATF3) ...........42, 67, 306 Activation transcription factor 4 (ATF4) ........................38, 41, 43, 45, 52, 61, 62, 67 Aflotoxin B1 aldehyde reductase (AFAR) ......................................................... 38, 40 Alanine ................ 44, 60, 144, 155, 239, 240, 256, 258, 260 Alkylation .....................................................52, 53, 65, 148 All-trans retinoic acid (ATRA) ....................................... 42 Alternative splicing .................................238, 239, 243–244 Antioxidant enzymes ........................................... 39–41, 43 Antioxidant response element (ARE) ....................... 37–68 Aphidicolin.....................................................114–119, 122 ARE inducers ..................................... 39, 40, 44, 45, 51–66 Arsenic ...................................................................... 55, 58 ATM ....... ................................................................ 33, 305 Autoradiography ...........................................224, 227, 228, 272, 273, 321, 333, 358, 365, 369, 393

B Bach 1(BTB and CNC homology 1) ........................ 41, 67 Basal condition .................................. 14, 41, 46, 48, 51–52, 60, 63, 66, 359, 365, 366 Basic leucine zipper (bZIP) ................................. 41–43, 45 Basic nuclear localization signal (bNLS) ................... 52, 63 b-catenin .......................................... 35, 187–197, 267, 275 BCL11B .................................................378–380, 382–389 Bifunctional inducers ....................................................... 39 Biotinylated DNA templates ......................................... 297 BJ1 cells ..................................................114, 116–119, 122 Blocking agents ............................................................... 39 Brahma-related gene 1(BRG1) ................................. 43, 45 BTB/POZ domain (Broad complex, Tramtrack and Bric-a-Brac/Poxivirus and zinc finger domain) ............................................... 45, 46 Butylated hydroxyanisole (BHA) .................................... 42

C Cadmium ........................................................................ 58 Calmodulin (CaM).................................................... 58, 67

CAND1 (Cullin-Associated NeddylationDissociated 1) ................................................ 49, 50 CARM1 (Coactivator-associated arginine methyltransferase 1) ....................................... 41, 67 Carnosol. See Diterpene Carotenoids ..................................................................... 39 Casein kinase 2 (CK-2) ..................................56, 58–59, 67 Catalase (CAT) .................. 38, 40, 127, 128, 131–134, 220 Catechins ........................................................................ 39 CBP/p300 (CREB-binding protein)............18, 41, 67, 318 Cell cycle ............................................. 9, 12, 19, 31, 48, 49, 96, 113, 114, 116, 117, 122, 188, 268, 305, 339, 340 Chromatin immunoprecipitation (ChIP) ChIP-ChIP ............................................................. 260 ChIP-seq ......................................................... 260, 261 Chromium ................................................................. 55, 58 Circular dichroism (CD) ............................................... 257 c-Jun...... ............................................................ 31–35, 216 c-Myc.. .................................................31–35, 96, 307, 308 Co-activators RAC3/SRC3 ....................................................... 41, 67 Cofactors-generating enzymes .................................. 38, 40 Co-immunoprecipitation (co-IP) ........................54, 57, 58, 219–220, 223, 258 Collagen ..........................162, 164, 165, 167, 201, 209, 215 Computational prediction ......................174, 253, 257, 261 Confocal microscopy .........................................85–87, 118, 146, 162, 163, 165–167, 189, 194, 195, 202 COP9 signalosome (CSN) ........................................ 49, 50 Co-repressors ............................................................ 9, 318 Coumarins ....................................................................... 39 Cr (VI). See Chromium CREB (c-AMP-response element-binding protein). See CBP/p300 (CREB-binding protein) CRM1 (chromosome region maintenance 1) ................. 44, 51, 60–63, 66, 67, 96, 143, 165, 167, 188, 191, 217 Crystallography ............................................................. 252 C-terminal region (CTR) .................................... 12, 44–46 Cul 3-based E3 ubiquitin ligase ................................ 46, 48 Cullin (Cul) ....................................................43, 48–50, 54 Cullin-RING ligases (CRLs) .......................................... 49 Cyclin B1................................................114, 116, 120, 121 Cyclooxygenase (COX) inhibitors ................................... 39 Cyclopentenone-containing molecules ............................ 54

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TRANSCRIPTION FACTORS 408 Index Cysteine (Cys) residue ....................... 38, 46, 51–53, 55, 63, 64, 65, 217 Cytochrome P-450 (CYP450) ........................................ 39

D Delayed/late response ...........................................61, 62, 66 Deneddylation ........................................................... 49, 50 Dephosphorylation ........................................23, 34, 58, 61, 62, 125–136, 140, 143, 148, 150–152, 318, 319 Detoxification enzymes ....................................... 38–43, 64 DIDLID element/DLG motif of the Neh2 domain of Nrf2 .............................................. 43, 45 Diterpene ........................................................................ 60 Dithiocarbamate ............................................................. 58 Dithiolethiones (DTs) ............................................... 39, 41 DMAT (2-dimethylamino-4,5,6,7-tetrabromo1H-benzimidazole).............................................. 58 DNA affinity ................................................................... 10 DNA repair enzymes ................................................ 38, 40 Double glycine repeat (DGR) .................. 43, 46, 50, 51, 53 Double glycine repeats (DGR) .................................. 43, 46 D3T (3H-1,2dithiole-3-thione). See Dithiolethiones (DTs)

E E1. See Ubiquitin-activating enzyme E2. See Ubiquitin-conjugating enzyme E3. See Ubiquitin-protein ligase Early cellular response ............................................... 61, 62 E-box .................................................... 239, 340, 341, 345, 348, 349, 353, 391–406 ECH (Erythroid-derived CNC Homology protein) ...............................................41, 43, 45, 58 Eicosapentaenoic acid (EPA) .......................................... 54 Electron paramagnetic resonance (EPR) ....................... 257 Electrophile response elements (EpREs) .......38, 41, 52, 61 Electrophoretic mobility shift assay (EMSA) ....................................154–155, 259, 340, 341, 343, 348, 392, 393, 395, 396, 399–400 Elf1.....................................................................5, 8, 11, 13 ELK-1 .........5, 7, 9, 10, 13, 15–23, 279–281, 283, 287, 292 Embryo mouse harvesting ..................................................... 241 Enhanced green fluorescent protein (EGFP) .................................. 44, 86, 143, 183, 189 EPA ..... ........................................................................... 54 Epoxide hydrolase (EH) ............................................ 38, 40 Er81.................................................. 5, 8, 10–11, 13, 15, 16 ERK. See Extracellular signal-regulated kinase (ERK) Erm .................................................................5, 7, 12, 16 ETGE motif of the Nh2 domain of Nrf2 ......43, 45, 51, 64 ETS transcription factors ETS-1 .................................... 4, 5, 7, 12–18, 20, 22, 23 ETS-2 ....................................................5, 7, 13, 21, 22 Euchromatin ................................................................. 318 Exportins ................. 44, 45, 51, 60, 62, 66, 67, 96, 185, 188

Extracellular signal-regulated kinase (ERK) .............. 7–10, 14, 16, 17, 19, 23, 32, 34, 57, 67, 116, 119–121, 216, 217, 227, 280, 292, 301, 302 Eye embryonic mouse .................................................... 243

F Ferritin ...................................................................... 38, 40 FLAG-M2 affinity gel ...................................269, 272, 274 Flavonoids ....................................................................... 41 Flow cytometry cellular subpopulations ............. 377, 378, 382, 383, 388 fixation..............................................381, 384–386, 388 flowJo software ........................................................ 386 fluorochrome-labeled antibodies ............................. 378 mean fluorescence intensity (MFI) ...........378, 383, 384 nuclear epitopes ................................378, 382, 383, 386 permeabilization ...............................378, 381, 383–389 staining ..................................... 378, 379, 381, 383–387 surface markers .................................377, 378, 384, 388 Fluorescence lifetime imaging microscopy (FLIM) ................................... 258, 261 Fluorescence recovery after photobleaching (FRAP) ....................................................86, 88–90, 152–153, 194, 195, 205, 206, 256, 260, 261 Fluorescence resonance energy transfer (FRET)............................... 201, 205–211, 258, 261 Forkhead box factors (FOXM1) ............................ 113–123 FRAP. See Fluorescence recovery after photobleaching Fyn kinase (Fyn) .......................................44, 60–62, 66, 67

G Gamma-glutamyl-cysteine synthetase (g-GCS)......................................................... 38, 40 Glutathione (GSH) .......................................38, 40–42, 53, 55, 60, 63, 219, 222, 224, 361 Glutathione reductase (GR) ..................................... 38, 40, 53, 200, 203–206, 209, 252, 255, 258–260 Glutathione S-conjugate efflux pumps ...................... 38, 40 Glutathione S-transferases (GSTs) .......................... 38–40, 42, 60, 63, 136, 219–228, 258, 273, 276, 314, 360, 361, 366, 372 Glycogen synthase kinase 3-b (GSK-3-b) ..................... 51, 60–62, 66, 67, 217, 221, 227–228 G2/M... ..................................................113, 114, 117, 118 Green fluorescent protein (GFP) ............................. 24, 86, 89, 99, 107–109, 130–132, 135, 141–146, 148, 150–155, 161–170, 173, 174, 183, 191, 193–195, 200–206, 209, 255, 256, 258, 378 GST-P. See Placental glutathione S-transferase GST-P enhancer 1 (GPE1)............................................. 42

H Heme oxygenase 1 (HO-1) ............................38–40, 56, 60 Heterochromatin ..................................................... 42, 318

TRANSCRIPTION FACTORS 409 Index Heterokaryon ........................................................ 161–170 High-affinity ETGE motif ............................................. 51 Histone deacetylases (HDACs) ...........................6, 16, 318, 319, 379, 380 Hrt1/Roc1/Rbx1 ...................................................... 48–49. See also Rbx1/Roc1/Hrt1 hTERT-BJ1................................................................... 114 Hydrogen peroxide (H2O2)...................................61, 62, 66 Hydroxylation activity assay .......................................... 220 4-Hydroxynonenal (4-HNE) .......................................... 56 Hypoxia-inducible factors (HIFs) HIF-1alpha (HIF-1a)...................................17–20, 35, 213–218, 223–228, 267, 275 HIF-2alpha (HIF-2a)..........................17–20, 213–218 HIF-3alpha (HIF-3a)..................................... 214–216 Post-translational regulation ............................ 213–229

I Immobilized metal affinity (IMAC) ............................. 253 Immunoblotting ............................................119, 120, 154, 226, 254, 272, 273, 283, 292, 294–295, 298–301, 306–309, 312, 327, 329, 330, 357–375 Immunofluorescence ................................................ 82, 83, 106–109, 152, 168, 191–192, 255 Immunoprecipitation (IP) .................................54, 57, 149, 156, 223, 225, 252–254, 258–260, 271, 281, 284, 287, 309–311, 322, 330, 334, 393, 397, 401, 402 Importation ....................................... 47, 52, 59–63, 65–67, 96, 97, 126, 128, 129, 140, 145–150, 194–197, 255 Importins karyopherins a and/ or b฀.....................45, 59, 66 Indoles ............................................................................. 39 Interferon ...............................................141–148, 150–156 Intervening region (IVR) ......................... 46, 47, 50, 51, 53 Intranuclear informatics .................................78, 85, 90–92 In vitro translation .........................................219, 222–223, 225, 270, 273, 276, 308–310, 312–314 Isoliquiritigenin ............................................................... 54 Isothiocyanate (ITC) ............................ 39, 41, 58, 146, 148 iTRAQ .......................................................................... 255

J Janus kinases ( JAKs) ............................................. 140–143 J3-isoprostanes ................................................................ 54 JNK1 (c-JunNH(2)-terminal kinase 1)/MAPK8 (mitogen-activated protein kinase 8) ........... 57, 216 JunD..... .............................................. 38, 41, 52, 61, 62, 67

K Karyopherins a and/ or b. See Importins karyopherins a and/ or b Keap1 (Kelch-like erythroid-cell-derived protein with CNC homology (CNC)-associated protein 1) ................... 38, 41–68 Keap1-Cul3 complexes........................................ 52–59, 65

Keap1-Cul3-Rbx1 E3 ubiquitin ligase complex .....................................38, 47, 52, 55 Keap1 cysteine residues ..................................52–59, 63, 65 Kelch repeats region. See Double glycine repeats (DGR) Keratinocyte culture HaCaT cells ............................................................ 127 Knockout .................................. 42, 141, 215, 340, 343, 395 Kruppel-like factor 5 (KLF5) .........................171, 268–276 Kruppel-like factor 8 (KLF8) .................171–174, 182, 183

L Leptomycin B (LMB) .........................................51, 60, 96, 99, 100, 104, 105, 107, 108, 188, 191–195 Leucine zipper (ZIP) domain ..............................41, 43–45, 63, 237–250, 340, 392 Leucokotriene B4 dehydrogenase (LTB4DH) ................ 38 Lipooxygenase (LOX) ..................................................... 39 Live cell imaging .................................................85, 86, 99, 106–109, 144, 194, 197, 201–210 Low-affinity DGL motif................................................. 51 Luciferase assay .....................................................120, 121, 203, 341, 343, 345, 348, 349, 352–353 Lysine (Lys) .....................................................6, 32, 43, 44, 46, 48, 51, 55, 65, 67, 98, 105, 115, 117, 118, 189, 197, 201, 209, 216, 220, 226, 239, 267, 280

M Maf, protein recognition element (MARE) .................... 42 Mass spectrometry (MS) .......................................253, 255, 258, 261, 292, 321, 326, 380, 387, 389 Matrix attachment regions (MARs) ...............317, 318, 331 MDM2................ 33, 35, 216, 268, 306–310, 312, 313, 314 MEK signalling MEK inhibitors ............................................... 220, 227 Raf/MEK/MAPK signaling ............................ 113–123 MHC class-I locus ........................................................ 317 Minigene ........................................................240, 244, 366 Mitogen activated protein kinase (MAPK) ................ 6–10, 14, 15, 19–24, 56–58, 67, 113–123, 216, 217, 239, 254, 280, 292 Molecular chaperones ................................................ 38, 40 Monofunctional inducers................................................. 39 Musculoaponeurotic fibrosarcoma (Maf ) .................................41, 42, 67. See also sMaf (small musculoaponeurotic fibrosarcoma) Mutagenesis PCR-based ...................................................... 175–176 site-directed ...............143, 174, 175, 200, 202, 252, 345

N NAD(P)H:quinone oxidoreductase 1 (NQO1) ......... 38, 40 Nedd8 modification of the cullin protein ........................ 49 Nedd8-specific E2 enzyme. See Ubc12 Neddylation ............................................................... 49, 50

TRANSCRIPTION FACTORS 410 Index Need8. See Neddylation NESTA ..........................................................44, 45, 52, 63 NESzip ...........................................................44, 45, 52, 63 Net phosphorylation ..............................................15, 16, 22 N-3 fatty acids. See EPA NF-E2-related factor 2 (Nrf2) .................................. 37–68 NF-E2-related factor 3 (Nrf3) .................................. 42, 67 Nickel-NTA agarose.............................................. 320, 324 Nicotinamide adenine dinucleotide phosphate (NADPH) ..................................................... 38, 40 Nrf2 cysteine modification ........................................ 59, 64 Nrf2-ECH homology (Neh) domains ..................... 43–46, 48, 51–54, 58, 63–65 Nrf2 knockout mice......................................................... 42 Nrf2 phosphorylation .....................................58–63, 65, 66 Nrf2 wild-type mice ........................................................ 42 Nuclear basic leucine zipper (ZIP) transcription factors ............................................................ 41, 43 Nuclear export signal (NES) ...............................10, 44–45, 47, 51, 59–61, 63, 64, 66, 67, 96, 188, 193, 217, 255 Nuclear import ............................ 45, 47, 52, 59–63, 65, 66, 96, 126, 128, 145–147, 149, 150, 152, 194, 195, 197 Nuclear localization bipartite ........................................................61, 62, 174 monopartite ..........................................44, 45, 173, 174 signal (NLS) ............................................10, 44, 45, 47, 59, 61, 62, 66, 169, 171–185, 194, 200, 255 Nuclear magnetic resonance (NMR) ..............252, 257, 261 Nuclear matrix ..............................................79, 82–84, 317 Nuclear microenvironment ....................... 77, 78, 82, 83, 88 Nuclear phosphatases ...............................58, 126, 140, 143 Nuclear receptors (NRs) ........................................ 251–262 Nucleocytoplasmic shuttling ...............................46, 48, 96, 128, 139, 144, 146, 162, 255, 256 Nucleolar organizing region (NORs) .............................. 80 Nucleoporins ....................................................96, 139, 149 Nucleosome remodeling ................................................ 318

O Oltipraz (OPZ) ......................................................... 39, 42 Oxidation .......................................................39, 52–54, 65 Oxidative stress........................................ 38, 40, 41, 43–53, 55, 56, 60, 62–64, 66, 67

P p53........................................................... 31–35, 42, 43, 55, 67, 267, 268, 305–314, 318 pCAF (p300/CBp-associated factors) .....................41, 216, 306, 318, 319 PDZ-like domain .......................................................... 318 Peroxiredoxins (Prxs) ................................................. 38, 40 Phase 1 xenobiotic detoxification enzymes...................... 40 Phase 2 xenobiotic detoxification enzymes...................... 40

Phenols ................................................ 39, 97, 98, 102, 109, 115, 163, 166, 169, 201, 203, 209, 219, 270, 287, 307, 320, 334, 364, 396, 399 Phenylalanine ............................................................ 44, 60 Phosphatase ........................................... 22, 23, 34, 58, 101, 115, 126–128, 133–134, 140, 143, 147, 151, 153, 260, 321, 334, 363, 395, 397 Phosphoglycerate mutase5 (PGAM5)..................47, 48, 67 Phosphorylation growth factor treatment ............................220, 226–227 GST-fusion proteins by GSK3b ...............221, 227–228 kinase inhibitors ...............................151, 220, 226, 227 phosphomimetics ..................................................... 258 phosphopeptide mapping ........................................ 252 phosphoproteomics.................................................. 253 phosphorylation site data bases................................ 253 site-specific antibodies ..................................... 254, 255 Photoactivation photoactivatable GFP ...............................165–167, 169 PI3K (phosphatidylinositol 3-kinase) ....................... 60, 66, 122, 216, 217 PKC (protein kinase C) .....................................7, 8, 11, 15, 56–58, 61, 62, 67, 319, 321, 322, 326, 327, 330, 331 PKR-like endoplasmic reticulum-resident kinase (PERK) ........................................57, 67, 121 Placental glutathione S-transferase (GST-P) .................. 42 p38 MAPK. See Mitogen activated protein kinase (MAPK) PMF-1 (Polyamine-modulated factor-1 protein) ..................... 38, 41, 52, 61, 62, 67 Polymerase chain reaction real time (qPCR) .............................................240, 242, 247–249, 340–347, 349–352, 397, 403–405 standard curve...................................240, 248–250, 344 Post-translational modifications (PTMs) ................... 3–25, 35, 52–59, 216, 218, 237, 238, 251, 260, 267, 268, 275, 279, 280, 282, 302, 305–314, 318, 319 Poxivirus and zinc finger (POZ) domain. See BTB/POZ PPM1A .......................................... 127, 128, 133, 134, 136 Pre-initiation complexes (PICs) .............291–302, 340, 391 PRMT1 (Protein arginine methyl-transferase 1) ...... 41, 67 Proteasome system proteosome inhibitor (MG132) .........................58, 269, 270, 274–276 Protein degradation cycloheximide (CHX) ..............................270–271, 274 half-life ............................................................ 271, 274 pulse chase ................................................270–272, 274 Protein kinase C ................................................7, 8, 11, 15, 56–58, 61, 62, 67, 319, 321, 322, 326, 327, 330, 331 Protein-protein interactions co-immunoprecipitation (co-IP) ..................... 219–220 GST pull-down assay ...............................220, 223–224 Protein trafficking

TRANSCRIPTION FACTORS 411 Index Q Quinone ................................................... 38, 40, 42, 55, 56

R RAC3 ....................................................................... 41, 67 Raf kinase ...................................................23, 24, 113–123 Raloxifene ........................................................................ 39 Ran-binding protein 3 (RanBP3) ...................126, 128–136 Rbx1/Roc1/Hrt1, 48–49. See also Hrt1/Roc1/Rbx1 Reactive nitrogen species (RNS) ..................................... 63 Reactive oxygen species (ROS).................................. 40, 63 Reactive thiol groups ....................................................... 52 Receptor associated coactivator (RAC3) ................... 41, 67 Recombinant proteins GST-recombinant protein expression and purification...................................219, 221–222 35 S-labeled protein in vitro translation ......219, 222–223 Redox-insensitive nuclear export signal in the ZIP domain of Nrf2. See NESzip Redox-sensitive nuclear export signal in the Neh5 transactivation (TA) domain of Nrf2. See NESTA Reduction .....................................................10, 39, 53, 288 Restriction endonuclease (RE) ...................................... 299 Resveratrol ....................................................................... 39 Retinoic acid (RA) .................................................... 42, 67 Retinoic acid receptor alpha (RARa) ....................... 42, 67, 254, 258, 259 Retinoids ..............................................................39, 42, 67 RING E3 ligases. See Cullin-RING ligases (CRLs) RNA..... ....................................16, 39, 80, 89, 91, 116, 128, 132, 133, 134, 222, 240–247, 249, 270, 291, 340–343, 346, 347, 349–351, 353, 384, 388 RNAP carboxy-terminal domain................................... 292 RNA polymerase II (RNAP)................................. 291–292 Runx factors ....................................... 78, 80, 83, 85, 86, 91

S Semi-quantitative polymerase chain reaction (PCR) .................................................. 279 Serine 40 (S40) .......................................................... 56, 57 SERM (selective estrogen receptor modulator) ............... 39 Serum deprivation ..................................114, 116–118, 122 Serum response factor (SRF)......................9, 280, 301, 302 10-Shogaol ...................................................................... 54 Signalosome. See COP9 signalosome Signal transducer and activator of transcription (STATs) .............. 139–143, 148–150, 155, 156, 255 Single nucleotide polymorphisms (SNPs) ............... 43, 398 siRNA (small interfering RNA) ..............................50, 191, 254, 255, 343 Skp1........................................................................... 12, 49 SKp1-Cullin-F-box protein (SCF) .....................12, 48–50, 268, 275

SMADS export assay ...............................................128, 131–133 Smad2................................................96–100, 104–109, 125–136, 268, 358–363, 365, 366, 370, 373 Smad3.................................................... 96, 97, 99, 100, 125–130, 133, 135, 136 Smad4.......................................... 96, 98–100, 103–109, 125, 126, 268, 358 Smad phosphatases .................................................. 126 YFP-conjugated sMaf (small musculoaponeurotic fibrosarcoma) ............. 38, 41, 42, 50, 52, 53, 61, 62, 67. See also Musculoaponeurotic fibrosarcoma (Maf ) S-nitrosylation ..........................................19, 215, 217–218 SOD (superoxide dismutase) ..................................... 38, 40 SRC3 (Steroid receptor coactivator-3). See RAC3 Src kinases ....................................................................... 63 Stable-isotope-labeling by amino acid in cell culture (SILAC) .................................................... 255, 258 Staurosporine..........................................143, 145, 151, 152 Streptavidin-coated magnetic beads ...............294, 297, 298 Sulfhydryls......................................................44, 53, 54, 63 Sulforaphane (SUL) ..................... 42, 50, 53–55, 57, 58, 63 SUMOylation ......................................... 4, 6, 8–10, 12–14, 16, 19, 215, 218, 239, 260

T t-BHQ (tert-butylhydroquinone). See Butylated hydroxyanisole (BHA) Theaflavins ...................................................................... 39 Thearubigins ................................................................... 39 Thiol groups .........................................................50, 52, 53 Thioredoxin reductase (TR) .................................38, 40, 53 Threonine (T) .....................6, 34, 61, 62, 95, 136, 254, 258 TIP120A. See CAND1 T-lymphocytes CD4+ cells ........................ 378, 379, 382–384, 386–389 CD8+ cells ........................................382–384, 388, 389 lymph nodes .............................................379, 385–386 spleen ........................................................379, 384–386 Transactivation (TA) domain. See NESTA Transcription factors ..................................3–25, 31–35, 38, 77–92, 96, 113, 139–156, 162, 163, 171, 188, 199–211, 213, 237, 252, 257, 259, 267, 279–288, 291, 317, 339, 357–375, 377–389, 391–406 Transfection ............................................. 87, 116, 127, 142, 162–163, 172, 189–191, 201, 219, 241, 254, 269–270, 306–307, 329, 345 Transforming growth factor-b(TGF-b) TGF-b receptors ........................ 95, 125, 133, 135, 268 TGF-b signaling .............................................. 95–110, 125–136, 268, 357, 358, 366 Tyrosine (Y ) .............................................. 6, 44, 60–62, 66, 67, 139, 140, 142–145, 147–149, 152–154, 254, 258

TRANSCRIPTION FACTORS 412 Index U Ubc12... ........................................................................... 50 Ubiquitin (Ub) ..................................................6, 8, 10, 12, 32–35, 38, 46–52, 55, 56, 67, 215, 216, 218, 260, 261, 267–270, 272, 273, 275, 276, 305–314 Ubiquitin-activating enzyme ..................267, 306, 307, 310 Ubiquitination Nrf2 ubiquitination .......................................38, 51, 52, 54–59, 63, 65, 67 Ubiquitin-conjugating enzyme .................48, 267, 306, 308 Ubiquitin-proteasome proteolysis system .................. 48, 52 Ubiquitin-protein ligase .................................................. 48 UDP-glucuronosyltransferase (UDP-GT) ................ 38–40 Under basal condition..........................................14, 41, 46, 48, 51, 60, 66, 365, 366 Upstream stimulatory factor stress response .......................................................... 340 target genes ...................................................... 339–354

USF-1................339–354, 392, 393, 396, 400, 402–404 USF-2............................... 340, 392, 393, 396, 400–404 UV....... .......................................61, 86, 105, 148, 177, 247, 282, 286, 340, 341, 343, 346, 347, 349, 352, 353

V Vitamin E........................................................................ 39

W WWP1 .................................................................. 268–276

X Xanthohumol .................................................................. 54 Xenobiotic ......................................................38–40, 61, 62 Xenobiotic response element (XRE) ............................... 39

Z ZIP. See Leucine zipper domain