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
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Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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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
3
31
37
Part II 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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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
161
171
187
199
Part III 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
215
237 251 267
279
291 305
317
Part IV 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
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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
Members
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
PD
TAD
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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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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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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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
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. Ets Factor Deregulation and Post-translational Modification in Cancer 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
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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
HIF-1a
Ets-1
Cooperation
Hypoxic target genes
Ndrg1
HIF-2a Ets-1
Flk-1 Endosialin Vegfr1
Degradation
Elk-1
Net
Elk-1
Epo Pai-1 Cited-2 Wisp2 Igfbp3
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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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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45. Juang YT, Solomou EE, Rellahan B, Tsokos GC (2002) Phosphorylation and O-linked glycosylation of Elf-1 leads to its translocation to the nucleus and binding to the promoter of the TCRi-chain. J Immunol 168:2865–2871 46. Miyazaki Y, Boccuni P, Mao S, Zhang J, Erdjument-Bromage H, Tempst P, Kiyokawa H, Nimer SD (2001) Cyclin A-dependent phosphorylation of the ETS-related protein, MEF, restricts its activity to the G1 phase of the cell cycle. J Biol Chem 276: 40528–40536 47. Liu Y, Hedvat CV, Mao S, Zhu XH, Yao J, Nguyen H, Koff A, Nimer SD (2006) The ETS protein MEF is regulated by phosphorylation-dependent proteolysis via the protein-ubiquitin ligase SCFSkp2. Mol Cell Biol 26:3114–3123 48. Suico MA, Nakamura H, Lu Z, Saitoh H, Shuto T, Nakao M, Kai H (2006) SUMO down-regulates the activity of Elf4/myeloid Elf-1-like factor. Biochem Biophys Res Commun 348:880–888 49. Zhang XK, Watson DK (2005) The FLI-1 transcription factor is a short-lived phosphoprotein in T cells. J Biochem 137:297–302 50. van den Akker E, Ano S, Shih HM, Wang LC, Pironin M, Palvimo JJ, Kotaja N, Kirsh O, Dejean A, Ghysdael J (2005) FLI-1 functionally interacts with PIASxa a member of the PIAS E3 SUMO ligase family. J Biol Chem 280:38035–38046 51. Murakami K, Mavrothalassitis G, Bhat NK, Fisher RJ, Papas TS (1993) Human ERG-2 protein is a phosphorylated DNA-binding protein-a distinct member of the Ets family. Oncogene 8:1559–1566 52. Pongubala JM, Van Beveren C, Nagulapalli S, Klemsz MJ, McKercher SR, Maki RA, Atchison ML (1993) Effect of PU.1 phosphorylation on interaction with NF-EM5 and transcriptional activation. Science 259: 1622–1625 53. Wang JM, Lai MZ, Yang-Yen HF (2003) Interleukin-3 stimulation of mcl-1 gene transcription involves activation of the PU.1 transcription factor through a p38 mitogenactivated protein kinase-dependent pathway. Mol Cell Biol 23:1896–1909 54. Mao C, Ray-Gallet D, Tavitian A, MoreauGachelin F (1996) Differential phosphorylations of Spi-B and Spi-1 transcription factors. Oncogene 12:863–873 55. Ray-Gallet D, Moreau-Gachelin F (1999) Phosphorylation of the Spi-B transcription
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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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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).
antibody
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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:N<1) (Fig. 2a, c). This was not observed to be the case for Smad2,
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Fig. 2. (a) Immunofluorescent determination of Smad2/4 localization upon LMB treatment. HaCaT cells were treated with or without 100 pM TGF-b for 1 h prior to harvest in the presence or absence of a 30 min pretreatment of 20 ng/ml LMB. (b) Quantification of immunofluorescent images for Smad2, and Smad4
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C:N = 1.22 ± 0.15 C:N = 0.57 ± 0.08 C:N = 0.53 ± 0.16 C:N = 0.46 ± 0.08
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Fig. 2. (continued) (c) Black bars indicate cytoplasm fluorescence intensity, while white bars indicate nuclear fluorescent intensity. Error bars indicate one standard deviation for the two measurements made for each cell.
where the ratio of cytoplasmic Smad2 to nuclear Smad2 (C:N) is 0.78 ± 0.06 in the basal state, 0.54 ± 0.12 when treated with TGF-b, 0.73 ± 0.07 when treated with LMB, and 0.32 ± 0.08 when treated with both TGF-b and LMB (Fig. 2a, b). We hypothesize that cause for the C:N <1 in the absence of both TGF-b and LMB is due to high nuclear background of the anti-Smad2 antibody used for immunofluorescent staining. 2. 4× round coverslips are placed one in each well of 4 wells in a 12 well plate tissue culture plate. To each well is added 1 ml of room temperature 1 mg/ml poly-d-lysine hydrobromide solution, and incubated for 1 h in the dark. Poly-d-lysine Solution is removed and reused up to ten times or before 2 months if stored at −80°C. Each well is washed with 1 ml of distilled water, and allowed to air dry in a sterile cell culture hood, with the UV light on. After 1 h, 200,000 cells are added to each well in 1 ml of DMEM supplemented with 10% FBS, 1× penicillin G/streptomycin sulfate solution, 1× l-glutamine solution (see Subheading 2). Cells are allowed to adhere to the coverslips and grow for 24 h at 37°C in a 5% CO2 atmosphere. 3. One well is untreated, and existing media is exchanged with fresh prewarmed media. In two other wells, LMB is added at a final concentration of 20 ng/ml for 30 min prior to addition of TGF-b1 at a final concentration of 100 pM to one of these wells and the remaining fourth well. TGF-b1 treatment lasts for 1 h, at which time cells are harvested.
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4. All media are removed from each well by aspiration, and washed one time with 1 ml of PBS. 1 ml of room temperature 3.7% paraformaldehyde (freshly made from 16% stock, diluted with water), and incubated at room temperature for 30 min. Paraformaldehyde solution is removed and fixed cells are washed one time with 1 ml PBS. 5. Fixed cells are blocked with 1 ml of TBS-t supplemented with 5% (v/v) normal goat serum (NGS), and allowed to incubate while rocking at room temperature for 1 h. All liquid is removed prior to proceeding to the next step. 6. 1 ml of solution containing 1:400 dilution of rabbit antiSmad2 antibody in TBS-t supplemented with 5% NGS is added to each well, and incubated while rocking overnight (approximately 10 h) at 4°C. Primary anti-smad2 solution is removed and cells are washed one time for 10 min at room temperature with 1 ml of PBS before proceeding to the next step. 7. 1 ml of solution containing a 1:400 dilution of mouse antiSmad4 antibody in TBS-t supplemented with 5% NGS is added to each well, and incubated for 4 h while rocking at room temperature. Cells are washed one time with 1 ml PBS for 10 min at room temperature before proceeding to the next step. 8. 1 ml of solution containing a 1:400 dilution of Alexa488 conjugated goat anti-mouse and 1:400 dilution of Alexa555 conjugated goat anti-rabbit antibodies in TBS-t supplemented with 5% NGS, and incubated at 4°C while rocking for 4 h, wrapped in aluminum foil to completely prevent exposure to light. This secondary staining solution is removed, and cells are rinsed quickly with 1 ml of PBS, and then washed for 20 min with 1 ml PBS containing 10 µg/ml Hoescht 33258, while rocking at room temperature without exposure to light. 9. Cells are rinsed twice with 1 ml PBS, and mounted on slides with 8 µl of 30% glycerol/TBS-t solution and coverslips are secured and sealed to the slides with a perimeter application of clear nail polish. 10. Cells are imaged using a Nikon microscope with a camera and Metamorph software. Images were integrated over 5 s. 3.4. Image Analysis for Immunofluorescence and Live Cell Imaging
1. For each image a ratio of the average cytoplasmic fluorescence intensity to the average nuclear fluorescence intensity was calculated using measurements made using the ImageJ program (see Note 11). 2. Four 20 × 20 Pixel squares were used to measure four values of integrated density for each of the five cells for each condition.
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Two squares were used to obtain the integrated density of the cytoplasm, while two squares were used to do the same for the nucleus of the same cell. Integrated density was measured using the “Analyze/Measure” function in ImageJ. This process was repeated for five total representative cells for each condition tested in both immunofluorescence experiments and live cell imaging experiments. 3. Averages for each of the two measurements were used to calculate the average integrated density for both the cytoplasm and nucleus of each cell. Reported values for each Cytoplasmic Integrated Density: Nuclear Integrated Density (C:N) consisted of average C:N values from five cells. Error for C:N is reported as a single standard deviation of the 5 C:N values for the five cells. 3.5. Live Cell Imaging of GFP-Smad4 and YFP-Smad2
1. Using live cell imaging analysis, we find that the ratio of cytoplasmic Smad4 to nuclear Smad4 (C:N) is 1.28 ± 0.47 in the basal state, 0.60 ± 0.15 when treated with TGF-b, 0.43 ± 0.04 when treated with LMB, and 0.41 ± 0.11 when treated with both TGF-b and LMB. Thus, both LMB and TGF-b treatment is sufficient to drive the Smad4 from mostly cytoplasmic (C:N>1) to mostly nuclear (C:N<1) (Fig. 3a, c). This was not observed to be the case for Smad2, where the ratio of cytoplasmic Smad2 to nuclear Smad2 (C:N) is 1.29 ± 0.15 in the basal state, 0.66 ± 0.10 when treated with TGF-b, 1.68 ± 0.28 when treated with LMB, and 0.65 ± 0.16 when treated with
Fig. 3. (a) Live Cell Imaging of GFP-Smad4 and YFP-Smad2. HaCaT cells expressing either N-terminally tagged GFP Smad4 or N-terminally tagged YFP Smad2 were imaged to determine the relative localization of GFP-Smad4 or YFP Smad2.
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C:N = 0.66 ± 0.10 C:N = 01.68 ± 0.28 C:N = 0.65 ± 0.16
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Fig. 3. (continued) (b) Quantification of fluorescent images for YFP-Smad2, and GFPSmad4 (c) Black bars indicate cytoplasm fluorescence intensity, while white bars indicate nuclear fluorescent intensity. Error bars indicate one standard deviation for the two measurements made for each cell.
both TGF-b and LMB (Fig. 2a, b). Thus, our live cell imaging data supports our findings for endogenous Smad4 and Smad2 using immunofluorescence image analysis. 2. For both GFP-Smad4 and YFP-Smad2 HaCaT cells, 200,000 cells in 1 ml are added to 3.5 mm glass bottom Mat-tek Petri dishes and allowed to settle for 24 h. 3. For each of the two cell lines 4 × 3.5 mm plates were prepared, where one plate was used as a control and media was exchanged for 1 ml of prewarmed DMEM media lacking
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phenol red and supplemented with 10% FBS, 1% penicillin/ streptomycin, and 1% l-glutamine solution (referred to as media from here on). To another plate, media was exchanged with 1 ml prewarmed media supplemented with 100 pM TGF-b1. Two more plates were prepared the same way but both using media supplemented with leptomycin B at a final concentration of 2 ng/ml. Thus, we are testing two variables independently. Cells are treated for 1 h prior to imaging. 4. GFP-Smad4 cells were imaged using the same methods as for AlexaFluor488, stated above. YFP-Smad2 cells were imaged using a 492/18 band pass excitation filter and a 535/30 band pass emission filter. For both, images were integrations of 3 s. 5. Images were processed using ImageJ software, as stated above for immunofluorescence data.
4. Notes 1. Cells prior to treatment should be fully spread out and 80% confluent. If too many cells are seeded at too high a density, clumps of cells will form in subsequent steps and lead to uneven lysing. 2. The term “rinse” is used here to refer to brief addition, mixing, and removal of solutions. In contrast, the term “wash” refers to the addition, timed incubation while mixing on a rocker, and removal of solutions. 3. The rate of cell swelling is not constant over different cell types, and the time of this incubation will have to be adjusted accordingly. 4. If cells are clumped together after this point, the final fraction of nuclear lysate will be partially contaminated with cytoplasm, due to incomplete lysis of plasma membrane prior to separation of nuclei by centrifugation. 5. If there are significant amounts of clumps of unbroken cells at this point, one may raise the amount in the hypotonic lysis buffer by adding an additional 30 µl, but this will dilute cytoplasmic fraction protein concentration. Alternatively, one may trypsinize cells at harvest, separate cells from trypsin by centrifugation, wash cells 1 time with 1 ml D-PBS, separate cells again by centrifugation, and resuspend cells in hypotonic lysis buffer. This trypsinization step is in place of Steps 1 and 2. 6. When cells are scraped and transferred to an Eppendorf tube on ice, they will settle to the bottom of the tube within 5 min. However, when plasma membranes are disrupted and nuclei
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are released, nuclei will not settle in less than an hour, and no pellet will form by gravity. 7. If one desires to remove the plasma membrane component of this fraction, one may spin at 13,200 × g for 10 min, and separate the cytoplasmic supernatant from the plasma membrane pellet, but this is not done in our procedure. 8. If the nuclear pellet is washed with a greater volume or more times than once with hypotonic buffer, then nuclei will adsorb to the inner walls of the eppendorf tube, and no pellet will form during separation of nuclei by centrifugation. However, this will not influence the rest of the procedure, as all liquid can be removed and RIPA buffer added to elute nuclei from the walls at the same time as causing complete lysis. 9. From here on the “cytoplasm” fraction will refer to the fraction containing cytoplasmic proteins as well as plasma membrane proteins. 10. We have determined that under these conditions in HaCaT cells the total protein yield in the nuclear fraction is two-thirds that of the cytoplasmic fraction, allowing us to accurately load equal ratios of cytoplasm and nuclear fractions. Thus, a different cell line may have a different ratio of total protein in the cytoplasm/plasma membrane relative to the nuclei, and that ratio will have to be determined experimentally based on the total yield of protein in these fractions over several experiments. 11. Rasband, W.S., ImageJ, National Institutes of Health, Bethesda, Maryland, USA, http://rsb.info.nih.gov/ij/, 1997–2004. References 1. Massague J (1998) TGF-b signal transduction. Annu Rev Biochem 67:753–791 2. Shi Y, Massague J (2003) Mechanisms of TGF-b signaling from cell membrane to the nucleus. Cell 113(6):685–700 3. Heldin CH, Miyazono K, ten Dijke P (1997) TGF-b signalling from cell membrane to nucleus through SMAD proteins. Nature 390(6659):465–471 4. Lagna G, Hata A, Hemmati-Brivanlou A, Massague J (1996) Partnership between DPC4 and SMAD proteins in TGF-b signalling pathways. Nature 383(6603): 832–836 5. Zhang Y, Feng X, We R, Derynck R (1996) Receptor-associated Mad homologues synergize as effectors of the TGF-b response. Nature 383(6596):168–172
6. Feng XH, Derynck R (2005) Specificity and versatility in TGF-b signaling through Smads. Annu Rev Cell Dev Biol 21:659–693 7. Inman GJ, Nicolas FJ, Hill CS (2002) Nucleocytoplasmic shuttling of Smads 2, 3, and 4 permits sensing of TGF-b receptor activity. Mol Cell 10(2):283–294 8. Xu L, Kang Y, Col S, Massague J (2002) Smad2 nucleocytoplasmic shuttling by nucleoporins CAN/Nup214 and Nup153 feeds TGFb signaling complexes in the cytoplasm and nucleus. Mol Cell 10(2):271–282 9. Nicolas FJ, De Bosscher K, Schmierer B, Hill CS (2004) Analysis of Smad nucleocytoplasmic shuttling in living cells. J Cell Sci 117(Pt 18): 4113–4125 10. Schmierer B, Hill CS (2005) Kinetic analysis of Smad nucleocytoplasmic shuttling reveals a
Analysis of Ligand-Dependent Nuclear Accumulation of Smads in TGF-b Signaling mechanism for transforming growth factor b-dependent nuclear accumulation of Smads. Mol Cell Biol 25(22):9845–9858 11. Kurisaki A, Kose S, Yoneda Y, Heldin CH, Moustakas A (2001) Transforming growth factor-b induces nuclear import of Smad3 in an importin-beta1 and Ran-dependent manner. Molecular Biol Cell 12(4): 1079–1091 12. Xiao Z, Liu X, Henis YI, Lodish HF (2000) A distinct nuclear localization signal in the N terminus of Smad 3 determines its ligandinduced nuclear translocation. Proc Natl Acad Sci U S A 97(14):7853–7858 13. Xiao Z, Liu X, Lodish HF (2000) Importin beta mediates nuclear translocation of Smad 3. J Biol Chem 275(31):23425–23428 14. Kurisaki A, Kurisaki K, Kowanetz M et al (2006) The mechanism of nuclear export of Smad3 involves exportin 4 and Ran. Mol Cell Biol 26(4):1318–1332
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15. Yao X, Chen X, Cottonham C, Xu L (2008) Preferential utilization of Imp7/8 in nuclear import of Smads. J Biol Chem 283(33): 22867–22874 16. Pierreux CE, Nicolas FJ, Hill CS (2000) Transforming growth factor b-independent shuttling of Smad4 between the cytoplasm and nucleus. Mol Cell Biol 20(23):9041–9054 17. Watanabe M, Masuyama N, Fukuda M, Nishida E (2000) Regulation of intracellular dynamics of Smad4 by its leucine-rich nuclear export signal. EMBO Rep 1(2):176–182 18. Xiao Z, Latek R, Lodish HF (2003) An extended bipartite nuclear localization signal in Smad4 is required for its nuclear import and transcriptional activity. Oncogene 22(7): 1057–1069 19. Liu X, Constantinescu SN, Sun Y et al (2000) Generation of mammalian cells stably expressing multiple genes at predetermined levels. Anal Biochem 280(1):20–28
Chapter 6 Raf/MEK/MAPK Signaling Stimulates the Nuclear Translocation and Transactivating Activity of FOXM1 Richard Y. M. Ma, Tommy H. K. Tong, Wai Ying Leung, and Kwok-Ming Yao Abstract The proliferation-associated transcription factor FOXM1 is essential for cell cycle progression into mitosis. Using synchronized human fibroblasts we detected, by immunostaining, that FOXM1 is localized predominantly in the cytoplasm in cells at late-G1 and S phases. Nuclear translocation occurs just before progression into the G2/M phase of the cell cycle and requires activity of the Raf/MEK/MAPK signaling pathway. Using pharmacological modulators, we demonstrated that activity of the Raf/MEK/MAPK pathway is both necessary and sufficient for the nuclear translocation of FOXM1. Consistent with FoxM1c being the major isoform expressed in proliferating fibroblasts, constitutively active MEK1 enhances the transactivating effect of FOXM1c, but not FOXM1b, on the cyclin B1 promoter in transient reporter assays. Here, we describe in detail the methods involved in generating these findings, which support the notion that FOXM1 is an effector of Raf/MEK/MAPK signaling in G2/M regulation. Key words: FOXM1, hTERT-BJ1, Raf/MEK/MAPK signaling, Nuclear translocation, Cyclin B1 promoter, G2/M, U0126
1. Introduction The transcription factor Forkhead box (FOX) M1, known previously as WIN, HFH-11, and Trident, is ubiquitously expressed in proliferating cells (1–4). FOXM1 levels show cell cycle phasedependent changes; its expression starts just before entry into S phase and peaks at the G2/M phase of the cell cycle (3, 5). In line with the mainly G2/M expression of FOXM1, FoxM1depleted mouse embryonic fibroblasts and cancer cell lines exhibit mitotic defects and polyploidy (6–8).
Paul J. Higgins (ed.), Transcription Factors: Methods and Protocols, Methods in Molecular Biology, vol. 647, DOI 10.1007/978-1-60761-738-9_6, © Springer Science+Business Media, LLC 2010
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Studies in both transgenic mice and HeLa cells indicate a dependency of FOXM1 function on mitogenic signals (5, 9). To further explore the molecular basis of this mitogenic dependence, we employed the human foreskin fibroblastic cell line hTERTBJ1 (BJ1) which can be efficiently synchronized by serum deprivation/aphidicolin double block (10), due to the presence of an intact G1/S checkpoint. Immunostaining revealed that FOXM1 expression is mostly cytoplasmic at late-G1 and S phases of the cell cycle (11). Nuclear translocation of FOXM1 occurs just before entry into the G2/M phase and is associated with phosphorylation of FOXM1. FOXM1 nuclear entry is abrogated when Raf/MEK/MAPK signaling is suppressed using the specific MEK1/2 inhibitor U0126 (11). In contrast, stimulation of Raf/ MEK/MAPK pathway activity using aurintricarboxylic acid (ATA) is enough to trigger enhanced FOXM1 nuclear translocation, which can be abolished by U0126 pretreatment. This data led us to conclude that MAPK signaling is necessary and sufficient for the nuclear translocation of FOXM1. To test whether nuclear entry of FOXM1 enhanced by stimulation of Raf/MEK/MAPK signaling has any effect on the transactivating activity of FOXM1, a constitutively active form of MEK1 (caMEK1) (12) was coexpressed with FOXM1 and the cyclin B1 reporter in transient assays. caMEK1 coexpression enhances the transactivating activity of FOXM1c, but not of FOXM1b (11). This enhancing effect is lost when a dominant negative form of MEK1 (dnMEK1) (12) is coexpressed. Taken together, our findings indicate that Raf/MEK/MAPK signaling enhances the nuclear translocation and transactivating activity of FOXM1c in human fibroblasts.
2. Materials 2.1. Cell Culture, Synchronization, and Flow Analysis
1. The human neonatal foreskin fibroblastic cell line hTERTBJ1 (BJ1) (Clontech) and Swiss NIH 3T3 cells (American Type Culture Collection) (see Note 1). 2. For BJ1 cells, Dulbecco’s modified Eagle’s medium (DMEM)/medium 199 (4:1, v/v) (Sigma) is supplemented with 10% (v/v) fetal bovine serum (FBS, BD Biosciences), 1 mM sodium pyruvate (Sigma), and 4 mM l-glutamine (Sigma). Swiss NIH3T3 cells are cultured in DMEM and 10% (v/v) FBS. 3. Phosphate-buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 1.5 mM KH2PO4, and 8 mM Na2HPO4. 4. 1% trypsin–EDTA solution (supplied as 10× from Gibco).
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5. Aphidicolin (Calbiochem): blocks DNA replication by inhibition of DNA polymerases a and d; dissolved in dimethyl sulfoxide (DMSO) at 1 mg/ml and stored at 4°C. 6. Ribonuclease T1 (Calbiochem): 10 mg/ml aqueous solution, preheated to 100°C for 5 min and stored as aliquots at −20°C. 7. Propidium iodide (PI; Calbiochem): 250 mg/ml in PBS with 1% Triton™ X-100 and stored at 4°C. 8. poly-l-lysine (Mr ³ 300,000; Sigma): 5 mg/ml solution in PBS. 2.2. Immunostaining
1. Fixing solution: methanol/acetone (1:1, v/v). 2. Blocking buffer: 3% BSA in PBS. 3. Primary antibody: anti-FOXM1 (C-20 rabbit polyclonal from Santa Cruz, sc-502). 4. Secondary antibody: Alexa fluor 488-conjugated goat antirabbit IgG ( Molecular Probes). 5. PI solution for nuclear counterstaining: PI (Calbiochem) dissolved at 12.5 mg/ml in PBS with 0.5% Triton™ X-100 and stored at 4°C. 6. Mounting medium: Vectashield™ (Vector Lab, Burlingame). 7. Inhibitor of Raf/MEK/MAPK signaling: U0126 (Calbio chem) is a specific noncompetitive inhibitor of both MEK1 and MEK2. Dissolved in DMSO at 10 mM and stored at −20°C. U0124 (Calbiochem), an inactive analog of U0126, is used as negative control. 8. Aurintricarboxylic acid (ATA, Calbiochem) is used to activate Raf/MEK/MAPK pathway activity. Dissolved in DMSO at 200 mM and stored at 4°C.
2.3. Western Blotting to Monitor MAPK Activity
1. Lysis buffer: 10 mM Tris–HCl (pH 7.4) containing 1% SDS, 1 mM EGTA, 1 mM DTT, and phosphatase inhibitors (0.1 mM sodium orthovanadate, 50 mM b-glycerophosphate). 2. 5× sample buffer: 62.5 mM Tris–HCl (pH 6.8), 10% glycerol, 2% SDS, 5% (v/v) 2-mercaptoethanol, 0.05% bromophenol blue; running buffer: 25 mM Tris–HCl (pH 8.3), 197.5 mM glycine, 0.1% SDS. 3. Electroblotting buffer: 30 mM Tris–HCl (pH 8.3), 200 mM glycine, 20% methanol. 4. Hybond™-C Super nitrocellulose membrane (Amersham Pharmacia Biotech). 5. Tris-buffered saline with Tween (TBST): 50 mM Tris–HCl (pH 7.4), 150 mM NaCl, 0.05% (v/v) Tween-20.
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6. Blocking buffer: 5% (w/v) nonfat dry milk in TBST. 7. Primary antibodies: anti-ERK1/2 (C-14 rabbit polyclonal from Santa Cruz, sc-154); anti-phosphoERK1/2 (Thr202/ Tyr204) (rabbit polyclonal from Cell Signaling Technology, #9101). 8. Secondary antibody: anti-rabbit IgG conjugated to horseradish peroxidase (BD Biosciences). 9. SuperSignal ECL detection system from Pierce. 2.4. Transient Reporter Assays
1. Transfection reagent: FuGENE® 6 (Roche Diagnostics). 2. Plasmid Midi Kit for DNA purification (QIAGEN). 3. The pGL3-CycBpro reporter (cyclin B1 promoter cloned upstream of Firefly luciferase) and FOXM1 expression plasmids (pcDNA3-FOXM1b and pcDNA3-FOXM1c) were previously described (5, 11). The expression vectors a-HA-MAPKKSDSE (caMEK1) and pSRa-HA-MAPKK-SASA (dnMEK1) were kindly provided by Dr. E. Nishida (12). 4. Dual-Luciferase® Reporter Assay system (Promega); pRLSV40 control reporter (SV40 promoter cloned upstream of Renilla luciferase as internal control of transfection, Promega).
3. Methods 3.1. Synchronization of BJ1 Cells
1. BJ1 cells are synchronized by serum deprivation/aphidicolin double block. For DNA analysis to follow cell cycle progression using flow cytometry, ~1.5 × 105 cells are seeded in 100 mm plates. After 36 h, cells are washed three times with PBS and replenished with DMEM/M199 and 0.1% FBS for 48 h. Afterwards, cells are treated with DMEM/M199 and 10% FBS containing 1 mM aphidicolin for 20 h. To release from arrest at the G1/S boundary, cells are washed once with PBS and replenished with DMEM/M199 and 10% FBS. 2. At different time intervals, cells are trypsinized and resuspended in DMEM/M199 and 10% FBS. Cells are then fixed and permeabilized overnight in 70% (v/v) ethanol at −20°C (see Note 2). Prior to measurement, cells are resuspended in 1 ml serum-free medium and treated with 40 ml of RNase T1 (10 mg/ml) for 3 min at room temperature. After RNA removal, cells are incubated with 250 ml of PI staining solution for 15 min at 4°C in the dark. Cells are then measured for DNA content using a standard flow cytometer (see Note 3). Typical flow profiles of asynchronized BJ1 cells and cells
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released from synchronization by serum deprivation/aphidicolin double block are shown in Fig. 1a. 3. For study of FOXM1 subcellular localization by immunostaining, cells are seeded onto poly-l-lysine-coated coverslips placed in 6-well plates. 18 mm × 18 mm coverslips are sterilized
Fig. 1. FOXM1 is translocated to the nucleus before entry into the G2/M phase. (a) Flow diagrams of asynchronized and synchronized BJ1 cells. BJ1 cells were synchronized at the G1/S boundary by serum starvation/aphidicolin double block. Cells were released from arrest by removal of aphidicolin. At different time intervals after release, cells were harvested and stained with propidium iodide for DNA analysis using a flow cytometer. Synchronized cells gradually progressed through the cell cycle and G2/M cells could be detected at both 9 and 12 h after release. (b) Nuclear translocation of FOXM1. BJ1 cells grown on coverslips were synchronized by serum starvation/aphidicolin double block and fixed at various time points after removal of aphidicolin. Cells were immunostained with anti-FOXM1 antibody and counterstained with propidium iodide to detect nuclear DNA. a, d, g, j, m: Fluorescent images of FOXM1 staining; b, e, h, k, n: Fluorescent images of nuclear staining; c, f, i, l, and o: Merged images of FOXM1 and nuclear staining. FOXM1 was predominantly cytoplasmic (arrows) at 0, 3, and 6 h after release; FOXM1 became mainly nuclear (arrowheads) at 9 and 12 h after release (Reproduced from (11) with permission from The Company of Biologists Ltd).
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by briefly dipping into 70% ethanol (v/v) and flaming over a Bunsen burner. They are placed onto 6-well plates and incubated with 0.4 ml of poly-l-lysine solution for 20 min. After rinsing once with PBS, they are left to dry before coating with cells. For each well, ~2 to 5 × 104 of BJ1 cells are seeded and the cells are synchronized as described in step 1 of Subheading 3.1. 3.2. Study of FOXM1 Nuclear Translocation by Immunostaining
1. BJ1 cells synchronized on coverslips in 6-well plates are harvested at different time intervals by washing twice with PBS and fixing with ice-cold methanol/acetone (1:1, v/v) solution for 10 min. Cells are allowed to air-dry for 5 min, followed by rehydration in PBS for 10 min. 2. Coverslips containing cells are placed on parafilm and incubated with anti-FOXM1 antibody diluted 1:200 in blocking buffer. After 1 h, coverslips are placed back onto 6-well plates and washed with PBS for four times (5 min each). Afterwards, coverslips are placed on parafilm again and treated with fluorochrome-conjugated secondary antibody (diluted 1:200 in blocking buffer) for 30 min in the dark. This is followed by washing with PBS again for four times (5 min each) to get rid of the unbound antibodies. 3. Nuclear counterstaining is performed by incubating with the PI solution for 30 s, followed by brief washing with PBS for six times. Coverslips are mounted on slides using Vectashield™ mounting medium and sealed with nonfluorescent nail polish. Signals can be visualized using regular fluorescence microscope or captured by confocal microscopy for better contrast (see Note 4). Typical images to illustrate the nuclear translocation of FOXM1 before entry into the G2/M phase are shown in Fig. 1b.
3.3. Study of FOXM1 Nuclear Translocation Following Pharmacological Modulation of MAPK Signaling
1. To identify the time window during which FOXM1 translocation occurs, BJ1 cells on coverslips are arrested by serum deprivation/aphidicolin double block as described in step 1 of Subheading 3.1. After aphidicolin removal, cells are harvested at 1-h intervals and subjected to immunostaining as described in Subheading 3.2. This analysis reveals that nuclear translocation of FOXM1 happens between 7 and 8 h after release (Fig. 2c, d). 2. To investigate whether U0126 can block FOXM1 nuclear translocation, synchronized cells are treated 1 h with 10 mM U0126 (or U0124) at 7 h after aphidicolin removal. Cells are harvested at 8 h after release for immunostaining. Treatment with U0126, but not the inactive analog U0124, inhibits FOXM1 nuclear entry (Fig. 2e, f).
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Fig. 2. Activity of the Raf/MEK/MAPK pathway is required for nuclear translocation of FOXM1. (a) Immunoblot analysis of total ERK and phosphoERK expression to monitor the effect of U0126 and ATA treatment on Raf/MEK/MAPK activity. (b–f) Effect of inhibiting MEK1/2. (b) Scheme for drug treatment. Synchronized BJ1 cells were incubated with U0126 (10 mM) or the inactive analog U0124 (10 mM) from 7 to 8 h after aphidicolin release. Cells were harvested 1 h later (i.e., at 8 h after release) and immunostained for FOXM1. (c) Without drug treatment, FOXM1 was predominantly cytoplasmic at 7 h after release. (d) One hour later, FOXM1 became mainly nuclear. (e) Treatment with U0126 abolished FOXM1 nuclear translocation at 8 h after release. (f) FOXM1 nuclear translocation was not affected by U0124 treatment. (g–k) ATA promotes nuclear translocation of FOXM1. (g) Scheme for ATA treatment and U0126/U0124 pretreatment. (h) In cells treated with solvent (DMSO), FOXM1 was predominantly expressed in the cytoplasm. (i) After incubation with ATA (200 mM) for 1 h, FOXM1 became mainly nuclear-localized. (j) ATA stimulation of FOXM1 nuclear translocation was abrogated by preincubation with U0126 (10 mM, 1 h). (k) U0124 pretreatment could not counteract the stimulatory effect of ATA (Reproduced from (11) with permission from The Company of Biologists Ltd).
3. To study the effect of activation of Raf/MEK/MAPK signaling on FOXM1 localization, asynchronized BJ1 cells are incubated with ATA for 1 h before harvesting them for immunostaining. As controls, asynchronized cells are incubated with DMSO instead of ATA, or pretreated with 10 mM U0126 (or U0124) for 1 h before incubation with ATA. Without synchronization and drug treatment, most BJ1 cells are at G1 or S phase with FOXM1 expressed in the cytoplasm (Fig. 2h). Incubation of asychronized BJ1 cells with ATA for 1 h dramatically increases the number of nuclear-stained cells (Fig. 2i), suggesting that ATA promotes nuclear transloca tion of FOXM1. To confirm that this ATA-induced nuclear translocation is mediated by Raf/MEK/MAPK signaling, BJ1 cells are pretreated with U0126 or U0124 for 1 h before ATA addition. ATA-induced nuclear translocation is abolished in the U0126-treated but not the U0124-treated cells (Fig. 2j, k) (see Note 5).
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4. To monitor Raf/MEK/MAPK pathway activity following drug treatment, immunoblot analysis is performed. Cells in each well are harvested and lyzed in ~100 ml of lysis buffer. After protein quantitation by Bradford analysis, appropriate volumes of samples (~10 mg) are mixed with 5× sample buffer and running buffer to make up a volume of 40 ml. The samples are boiled for 10 min before gel loading. Protein samples are separated by SDS–PAGE in 8% regular Laemmli gels and electro-blotted onto Hybond C Super nitrocellulose membranes. Electroblotting is carried out at 25 V and 4°C overnight in electroblotting buffer. Blots are blocked by shaking in blocking buffer for 1 h at room temperature before incubation with anti-ERK (1:200) or anti-phosphoERK (1:1,000) antiserum diluted with blocking buffer for 1 h. After washing with TBST three times (15 min each), blots are incubated with secondary antibodies (1:2,000) for another hour before washing again, three times with TBST. Antigen–antibody complexes are visualized using the SuperSignal ECL detection system. 3.4. Transient Reporter Assays to Test the Effect of Raf/MEK/ MAPK Signaling on FOXM1 Transactivating Activity
1. The transactivating effect of FOXM1 on cyclin B1 promoter activity is assayed by transient transfection of expression and reporter plasmids into NIH3T3 cells using FuGENE® 6. ~9 × 103 NIH 3T3 cells are seeded onto each well of 24-well plates and allowed to settle for more than 24 h. For each reaction, a DNA mixture containing 200.4 ng of DNA (see Note 6) is added to 0.6 mg of FuGENE® 6 and 7.4 ml of DMEM (without FBS and antibiotics). Each DNA mixture typically contains two reporters (100 ng of pGL3-CycBpro and 0.4 ng of pRL-SV40) and different combinations of FOXM1 (i.e., pcDNA3-FOXM1c, pcDNA3-FOXM1b, or FOXM1cDCter) and MEK1 (i.e., caMEK1 or dnMEK1) expression plasmids (adding up to a total of 100 ng for each well, see Note 7). Each DNA–FuGENE® 6 mixture is left to sit at room temperature for 15 min and subsequently added to cells bathed in a well containing 0.5 ml DMEM (10% FBS). Cells are harvested after 48 h of incubation when they are about 90% confluent. 2. Luciferase activities are measured using the Promega DualLuciferase® Reporter Assay system. Cells in each well are washed with 1 ml of PBS and lyzed with 150 ml of 1× passive lysis buffer (PLB), which is prepared by mixing one part of 5× PLB (provided) with four parts of water. 20 ml of each sample is taken for measurement of luminescence (see Note 8). Luciferase Assay Reagent II (LAR II) is prepared by resuspending the provided Luciferase Assay Substrate in 10 ml of the supplied Luciferase Assay Buffer II. Stop and Glo® substrate is
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prepared by mixing the provided Stop & Glo® substrate powder with the provided solvent. One part of this substrate is mixed with 50 parts of Stop & Glo® Buffer to constitute the Stop & Glo® reagent prior to use. For each sample, 50 ml of LAR II reagent and the same volume of Stop & Glo® reagent are used for measurement. All experiments are conducted in triplicate using independent cultures and the results shown are in means ± s.e.m. (see Note 9). Typical data obtained for the transient reporter assays are shown in Fig. 3.
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Fig. 3. Constitutively active MEK1 enhances the transactivating activity of FOXM1c. (a) Schematic diagrams of FOXM1c, FOXM1b, and FOXM1cDCter. Positions of the DNA binding domain (DBD), exon Va, and transactivation domain (TAD) are shown. FOXM1b lacks exon Va. FOXM1cDCter was generated by deletion of the last 71 amino acids of FOXM1. (b–c) Transient reporter assays. NIH3T3 cells were cotransfected with the various expression plasmids and Cyclin B1 luciferase reporter. 48 h after transfection, cells were harvested for luciferase assay. (b) caMEK1 enhances the transactivating activity of FOXM1c. Cotransfection of caMEK1 (30 ng) with an increasing amount of FOXM1 strongly enhanced the transactivating activity of FOXM1c, but not FOXM1b. (c) The caMEK1 enhancing effect requires the presence of functional FOXM1 protein. Various amounts of FOXM1 and MEK1 expression plasmids, and empty vectors (pcDNA3 and pSRa), were cotransfected as indicated. Both caMEK1 and functional FOXM1c are required for the synergistic activation of Cyclin B1 promoter. (d) Western blot to demonstrate the activating and inhibitory effect of caMEK1 and dnMEK1, respectively, on Raf/MEK/MAPK signaling (Reproduced from (11) with permission from The Company of Biologists Ltd).
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4. Notes 1. BJ1 cells behave like primary cells that are contact-inhibited, and can be maintained as a monolayer at 100% confluency for more than 2 weeks. BJ1 cells can be synchronized at high efficiency (>95% synchronized) by serum-deprivation/aphidicolin double block although they are resistant to transient transfection using common transfection reagents. Swiss NIH3T3 cells which can be transfected efficiently are used as host for transient reporter assays. 2. This is a critical step as shorter treatment or treatment with ethanol at room temperature leads to suboptimal PI staining of cells. Cells can be stored at this step for up to a month before measurement. 3. We have been using an EPICS Elite ESP high performance cell sorter (Coulter Electronics) equipped with a red pass filter of 600 nm wavelength. Raw data is analyzed with EXPO™ for Windows 2.0 and Modfit™ LT version 2.0 (Verity software) to eliminate aggregated cells for determination of cell cycle distribution. The histograms are saved as TIFF images and are simplified using photo-editing software Corel© Draw™ Version 11. 4. We have been using a MRC1024 confocal imaging laser scanner (BioRad, Hercules, CA) equipped with Zeiss Axiovert 135 microscope (Carl Zeiss). Images are recorded and analyzed using Lasersharp version 3.2. For each sample, three images (red channel, green channel and overlay) are recorded through a 63× objective lens under oil immersion. Each image represents an area of approximately 175 mm × 180 mm. 5. Unlike U0126 which is specific against Raf/MEK/MAPK signaling by directly inhibiting MEK1/2, ATA acts more upstream by stimulating IGF-1 receptor signaling and it also activates phosphatidylinositol 3-kinase and phospholipase Cg. Therefore, the U0126/U0124 pretreatment controls are necessary to rule out possible nonspecific effects. Since ATAinduced nuclear translocation of FOXM1 is abolished by U0126 but not U0124, the ATA-dependent effect is mediated by Raf/MEK/MAPK signaling. 6. All plasmid DNAs are prepared using the Plasmid Midi Kit from Qiagen, according to the manufacturer’s instruction. 7. The parental cloning vectors pcDNA3 and pSRa are added to normalize the amount of plasmids across all transfections. 8. We have used the EG&G Berthold Microplate Luminometer LB96V to measure luminescence. The delay time is set at 2 s
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while the measure interval is 10 s for measurements of both Firefly and Renilla luciferase activities. 9. We have found that absolute luciferase values may vary from one experiment to another, probably due to variations among different bottles of LAR II and Stop & Glo reagents. However, the fold induction values are generally reproducible across experiments performed under similar conditions.
Acknowledgments We thank Dr. E. Nishida (Kyoto University) for providing the caMEK1 and dnMEK1 expression plasmids. This work was supported by a Hong Kong Research Grants Council Grant 7650/05M awarded to K.-M. Yao. References 1. Yao K-M, Sha M, Lu Z, Wong GG (1997) Molecular analysis of a novel winged helix protein, WIN: expression pattern, DNA binding property and alternative splicing within the DNA binding domain. J Biol Chem 272:19827–19836 2. Ye H, Kelly TF, Samadani U, Lim L, Rubio S, Overdier DG, Roebuck KA, Costa RH (1997) Hepatocyte nuclear factor 3/fork head homolog 11 is expressed in proliferating epithelial and mesenchymal cells of embryonic and adult tissues. Mol Cell Biol 17:1626–1641 3. Korver W, Roose J, Clevers H (1997) The winged-helix transcription factor Trident is expressed in cycling cells. Nucleic Acids Res 25:1715–1719 4. Kaestner KH, Knochel W, Martinez DE (2000) Unified nomenclature for the winged helix/forkhead transcription factors. Genes Dev 14:142–146 5. Leung TWC, Lin SSW, Tsang ACC, Tong CSW, Ching JCY, Leung WY, Gimlich R, Wong GG, Yao K-M (2001) Over-expression of FoxM1 stimulates cyclin B1 expression. FEBS Lett 507:59–66 6. Laoukili J, Kooistra MRH, Bras A, Kauw J, Kerkhoven RM, Morrison A, Clevers H, Medema RH (2005) FoxM1 is required for execution of the mitotic programme and chromosome stability. Nature Cell Biol 7:126–136
7. Wang I-C, Chen Y-J, Hughes D, Petrovic V, Major ML, Park HJ, Tan Y, Ackerson T, Costa RH (2005) Forkhead box M1 regulates the transcriptional network of genes essential for mitotic progression and genes encoding the SCF (Skp2-Cks1) ubiquitin ligase. Mol Cell Biol 25:10875–10894 8. Wonsey DR, Follettie MT (2005) Loss of the forkhead transcription factor FoxM1 causes centrosome amplification and mitotic catastrophe. Cancer Res 65:5181–5189 9. Ye H, Holterman AX, Yoo KW, Franks RR, Costa RH (1999) Premature expression of the winged helix transcription factor HFH11B in regenerating mouse liver accelerates hepatocyte entry into S phase. Mol Cell Biol 19:8570–8580 10. Chen MS, Hurov J, White LS, WoodfordThomas T, Piwnica-Worms H (2001) Absence of apparent phenotype in mice lacking Cdc25C protein phosphatase. Mol Cell Biol 21:3853–3861 11. Ma RYM, Tong THK, Cheung AMS, Tsang ACC, Leung WY, Yao K-M (2005) Raf/ MEK/MAPK signaling stimulates the nuclear translocation and transactivating activity of FOXM1c. J Cell Sci 118:795–806 12. Fukuda M, Gotoh I, Adachi M, Gotoh Y, Nishida E (1997) A novel regulatory mechanism in the mitogen-activated protein (MAP) kinase cascade. Role of nuclear export signal of MAP kinase kinase. J Biol Chem 272:32642–32648
Chapter 7 Coupling of Dephosphorylation and Nuclear Export of Smads in TGF-b Signaling Fangyan Dai, Xueyan Duan, Yao-Yun Liang, Xia Lin, and Xin-Hua Feng Abstract In eukaryotes, regulation of signaling mediators/effectors in the nucleus is one of the principal mechanisms that govern duration and strength of signaling. Smads are a family of structurally related intracellular proteins that serve as signaling effectors for transforming growth factor beta (TGF-b) and TGF-b-related proteins. Accumulating evidence demonstrates that Smads possess intrinsic nucleocytoplasmic shuttling capacity, which enables them to transmit TGF-b signals from cell membrane to nucleus. We recently identified two important steps in the termination of nuclear Smad signaling. The first step is initiated by a serine/threonine phosphatase PPM1A that dephosphorylates Smad2/3 in the nucleus, thereby shutting down signaling capacity of phosphorylated Smad2/3. The second step involves nuclear export of dephosphorylated Smad2/3 with the aid of nuclear protein RanBP3 to terminate Smad signaling. This chapter introduces methods for examining nuclear export of Smad2/3 in TGF-b signaling. Key words: Smad, PPM1A, RanBP3, Nuclear phosphatase, Nuclear export, Signal transduction
1. Introduction Smad proteins are critical intracellular signaling mediators for the transforming growth factor beta (TGF-b) superfamily. Eight Smads have been identified in mammals, including five receptoractivated Smads (R-Smads), one common mediator Smad (Co-Smad, i.e., Smad4) and two inhibitory Smads (I-Smads). Within R-Smads, Smad2 and Smad3 transduce signals from TGF-b and activin, while Smad1, Smad5, and Smad8 transduce signals from members of the Bone Morphogenetic Proteins/ Growth Differentiation Factors (BMP/GDF) family. At the resting state, Smad2/3 are generally diffused within cells. Upon ligand-induced activation of TGF-b receptors, Smad2/3 become phosphorylated by the TGF-b type I receptor (TbRI), dissociate Paul J. Higgins (ed.), Transcription Factors: Methods and Protocols, Methods in Molecular Biology, vol. 647, DOI 10.1007/978-1-60761-738-9_7, © Springer Science+Business Media, LLC 2010
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from the receptor, oligomerize with Co-Smad4, and translocate to the nucleus, where the Smad complex regulates transcription of TGF-b target genes (1, 2). At the end of active signaling, phospho-Smad2/3 are dephosphorylated (3) and exported from nucleus to cytoplasm. During the activation–inactivation cycles of TGF-b signaling, TGF-b-induced Smad nuclear accumulation has been the subject of particular scrutiny. The subcellular distribution of Smads depends on their association with nuclear import or export factors and retention proteins (4–8). Although it has been reported that TGF-b-induced phosphorylation favors the nuclear import of phosphorylated R-Smads by enhancing its association with importin-b (9, 10) and/or disassociation with cytoplasmic retention factors such as SARA (11), in-depth analysis suggests a more pivotal role of export process in Smad nuclear accumulation. It appears that TGF-b does not affect the nuclear import rate of Smad2 (8), instead it decreases Smad2 nuclear export and therefore retains more Smad2 in the nucleus (8, 12). Our recent studies investigate the functional roles of Smad2/3 export in TGF-b signaling. Examination of Smad2/3 export with the methods introduced in this chapter has revealed that Ranbinding protein 3 (RanBP3) negatively regulates TGF-b signaling by facilitating Smad2/3 nuclear export. In the nucleus, RanBP3 recognizes dephosphorylated Smad2/3, which results from the activity of nuclear Smad phosphatases and exports Smad2/3 in a Ran-dependent manner. As a result, increased expression of RanBP3 inhibits TGF-b-induced Smad2/3 nuclear accumulation. Conversely, depletion of RanBP3 expression or interference of RanBP3–Smad2/3 interaction retains more Smad2/3 in the nucleus. The methods for analyzing Smad2/3 nuclear export are presented here to illustrate the identification and characterization of RanBP3 as the Smad2/3 nuclear export mediator. We hope these methods would provide researchers with necessary concepts and tools for studying subcellular distribution, in particular nuclear export of proteins.
2. Materials 2.1. Cell Culture Reagents (Hyclone)
1. Minimum Essential Medium (MEM). 2. Dulbecco’s Modified Eagle’s Medium (DMEM). 3. Phosphate buffered saline (PBS), calcium and magnesium free. 4. Fetal bovine serum (FBS).
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5. Penicillin/streptomycin sulfate. 6. Nonessential amino acids (NEAA). 7. 0.25% Trypsin/EDTA. 8. Human keratinocytes (HaCaT cells). 9. Human embryonic kidney (HEK293T) cells. 2.2. Chemicals and Buffers
1. TGF-b1 (R&D systems). 2. TbRI inhibitor SB431542 (CalBiochem). 3. Protease inhibitor cocktail (Roche). 4. ATP, creatine phosphate, and creatine phosphokinase (Sigma). 5. Fractionation buffer: 10 mM HEPES (pH 7.9), 1.5 mM MgCl2, 10 mM KCl, 0.5 mM DTT, 0.5% NP-40. 6. Lysis buffer: 25 mM Tris (pH 7.5), 300 mM NaCl, 1% Triton X-100. 7. Western blot Sample buffer: 2× SDS Sample buffer (BioRad). 8. Transporting buffer: 20 mM HEPES (pH 7.3), 110 mM KAc, 5 mM NaAc, 2 mM DTT, 1 mM GTP, 1.0 mM EGTA. 9. ATP regeneration system (Sigma): 1 mM ATP, 5 mM Creatine phosphate, and 20 U/ml Creatine phosphokinase. 10. In vitro phosphatase reaction buffer: 50 mM Tris–HCl (pH 7.5), 30 mM MgCl2, 5 mM DTT, and 1 mg/ml of BSA.
2.3. Plasmids and Reporter Reagents
1. MS2 coat protein-fused Smad2 plasmid (gift of Joan Massagué) (12). 2. pDM128/8xMS2 export reporter system (gift of Bryan Cullen) (14). 3. pSVbgal (Promega). 4. ELISA-based Chloramphenicol acetyltransferase (CAT) assay kit (Roche). 5. ELISA-based b-galactosidase assay kit (BD Pharmingen). 6. LipofectAmine transfection reagent (Invitrogen).
2.4. Antibodies
1. Anti-PPM1A antibody (Abcam). 2. Anti-Flag affinity gel (Sigma). 3. Anti-GADPH antibody (Research Diagnostics). 4. Anti-Lamin A/C (E-20) (Santa Cruz Biotechnology). 5. Anti-Smad2 (Zymed), anti-Smad2/3 (Santa Cruz Biotech nology), anti-phospho-Smad2 and phospho-Smad3 antibodies (Cell Signaling Technology).
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3. Methods The nucleocytoplasmic shuttling of Smad2/3 is governed by several mechanisms involving nuclear import/export mediators and/or retention factors that tend to retain Smad2/3 in the cytoplasm or nucleus (4–8). Often, retention factors interact with Smad2/3 and function through interfering with the recognition of Smad2/3 by import/export mediators (11, 12). Thus the nuclear Smad2/3 level at a specific time is determined by the net effect of its import and export. Three assays introduced here, including cell fractionation assay, in vitro export assay, and quantitative Smad2 export assay, are all aimed at examination of Smad2/3 nuclear export. The first two assays determine the nuclear Smad2/3 level under manipulated experimental conditions so that Smad2/3 import could be neglected (cell fractionation assay) or is disrupted (in vitro export assay). The quantitative Smad2 export assay utilizes a reporter system that produces CAT activity proportional to the nuclear export of Smad2. In the end of this chapter, we will also introduce the in vitro phosphatase assay, which we used to determine whether phosphorylated Smad2/3 are direct substrates of phosphatase PPM1A. This chapter assumes the familiarity of the readers with several common techniques including production of recombinant protein, cell culture, SDS–PAGE, Western blot analysis, and RNA interference. 3.1. Mammalian Cell Culture
1. HaCaT cells are propagated in MEM supplemented with 10% FBS, 100 µM NEAA, 100 µg/ml streptomycin sulfate, and 100 U/ml penicillin (10% FBS MEM). To maintain HaCaT cells in the resting state, cells are cultured in MEM with 0.2% FBS for 24 h before the experiments (0.2% FBS MEM). 2. Human embryonic kidney (HEK293T) cells are propagated in DMEM with 10% FBS, 100 µg/ml streptomycin sulfate, and 100 U/ml penicillin (10% FBS DMEM).
3.2. Cell Fractionation Assay
Subcellular fractionation is an extremely useful method to produce extracts enriched for proteins from the specific cellular compartments. The separation of nuclear and cytoplasmic fractions via this method allows the determination of Smad2/3 levels in each fraction by Western blot analysis. As the dysfunction of Smad2/3 nuclear export would alter its cellular distribution, here we compare the redistribution of nuclear accumulated Smad2/3 in RanBP3-depleted HaCaT cells and its parental HaCaT cells. 1. HaCaT cells, and HaCaT cells stably expressing short hairpin RNA (shRNA) against human RanBP3 at 75–85% confluency are cultured in 0.2% FBS MEM for 24 h to maintain cells in the resting state. HaCaT cells stably expressing shRNA against
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human RanBP3 are established beforehand by using standard method (see Note 1). 2. Treat cells (1 × 105) with TGF-b (2 ng/µl) in 0.2% FBS MEM for 1 h to induce nuclear accumulation of Smad2/3 (see Note 2). 3. Wash cells with PBS three times to remove TGF-b, treat with SB431542 (5 µM) in 0.2% FBS MEM to inhibit TGF-b type I receptor kinase activity for up to 4 h (see Note 3). 4. At each chosen time point (e.g., 0, 2, 4 h after SB431542 treatment), wash cells with PBS once, detach cells with 0.25% Trypsin/ EDTA. After cells are detached, inactivate Trypsin activity by the addition of 1 ml regular culture medium (i.e., 10% MEM). 5. Resuspend cells in 1 ml of 10% FBS MEM and split cell suspension into Eppendorf tubes with 200 µl in one tube and 800 µl in the other. Collect cell pellets by centrifugation at 500 × g, 4°C, for 5 min. Pellets from 200 µl cells are lysed directly with 100 µl 1× SDS sample buffer (50 µl H2O and 50 µl 2× SDS sample buffer) as whole cell lysates (see Note 4). 6. For subcellular fractionation, pellets from 800 µl cells were washed with cold PBS once, cell pellets were gently resuspended in 100 µl fractionation buffer and kept on ice for 20 min. 7. Collect cells by centrifugation at 500 × g, 4°C, for 5 min. Transfer the supernatant (cytoplasmic fraction) in a new tube. 8. Wash the pellets (nuclei) with 500 µl fraction buffer twice to eliminate cytoplasmic material contamination and then lyse the samples with 100 µl lysis buffer. 9. Add 100 µl 2× SDS Sample buffer into the subcellular fractions for sample preparation. 10. To evaluate the separation quality of subcellular fractions, examine GADPH (cytoplasmic protein marker) and Lamin A/C (nuclear protein marker) protein levels in each sample by Western blot analysis (see Note 5). These protein markers also serve as loading control for each fraction. 11. After successful separation of the cytoplasmic and nuclear fractions is confirmed, determine the Smad2/3 level in nuclear and cytoplasmic fraction as well as in whole cell lysates by Western blot analysis. An example of the result produced is shown in Fig. 1. 3.3. In Vitro Export Assay
In vitro export assay utilizes digitonin to permeabilize cell plasma membranes for macromolecules, yet leave the nuclei structurally and functionally intact. Washing the permeabilized cells removes majority of the cytoplasmic proteins, which prevents the cytoplasmic Smad2/3 being imported into the nucleus again. Thus, the level of Smad2 in samples represents nuclear Smad2 that has not been exported. We use this assay to compare the effect of RanBP3 and its Ran-binding mutant RanBP3-wv (see Note 6) on Smad2 export.
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Fig. 1. Depletion of RanBP3 blocks nuclear export of Smad2/3 in HaCaT cells. RanBP3-KD1 and parental HaCaT cells were treated with TGF-b for 1 h, cells were washed three times to remove TGF-b and treated with SB431542 up to 4 h. Cells were then harvested at indicated time, and the nuclear and cytoplasmic fractions were collected. Total Smad2/3 levels as well as phosphorylated Smad2 levels were examined by Western blot analysis. Detection of nucleus-localized Lamin A/C and cytoplasm-localized GADPH demonstrates separation of the fractions and proper sample loading. Quantitation of image intensity was done using Image J software. The arbitrary unit for cytoplasmic Smad2/3 level in lane 4 was set to 1. N nuclear fraction, C cytoplasmic fraction. As shown here, at time point 0 (immediately after 60 min TGF-b treatment), a very low level of Smad2/3 in the cytoplasmic fractions of both wild-type and knockdown cell lines is detected, indicating most of the Smad2/3 resided in the nucleus. Although SB431542 treatment promotes the dephosphorylation of Smad2 in the nucleus and the cycling back of Smad2/3 in the cytoplasm in parental HaCaT cells, it exhibits a weaker effect on cytoplasmic accumulation of Smad2/3 in RanBP3-KD1 cells. The total Smad2/3 levels are comparable in RanBP3-KD1 and parental HaCaT cells.
1. HaCaT cells stably expressing GFP-Smad2 (see Note 7) at 75–85% confluency are cultured in 0.2% FBS MEM for 24 h to maintain cells in the resting state. 2. Treat cells (1 × 104 per well in a 12-well plate) with TGF-b (2 ng/µl) in 0.2% FBS MEM for 1 h to induce nuclear accumulation of GFP-Smad2 (see Note 2).
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3. Wash cells with cold PBS once, incubate cells with 250 µl cold Transporting buffer containing digitonin (30 ng/µl) on ice for 5 min (see Note 8; Fig. 2). 4. Wash cells with cold Transporting buffer three times, remove excess buffer after last wash. 5. Immediately incubate the permeabilized cells with 200 µl Transporting buffer (prewarmed at 30°C) supplemented with ATP regeneration system and 500 ng of recombinant protein RanBP3, RanBP3-wv, or BSA as control (see Note 9). Perform the assay at 30°C for up to 60 min. 6. At each chosen time points (0, 30, and 60 min after incubation), quickly wash cells with the cold Transporting buffer three times, remove excess buffers after last wash. 7. Lyse cells in 100 µl 1× SDS sample buffer (50 µl H2O and 50 µl 2× SDS-sample buffer) for sample preparation. 8. Determine GFP-Smad2, GADPH (cytoplasmic protein marker), and Lamin A/C (nuclear protein marker) by Western blot analysis. These protein markers also serve as loading control for each sample. An example of the result is shown in Fig. 3. 3.4. Quantitative Smad2 Export Assay
This assay utilizes a CAT reporter system based on the observation that intron-containing mRNAs are exported out of nucleus only after splicing is completed (13). As shown in Fig. 4, the system consists of two components: (1) CAT reporter plasmid pDM128-8xMS2 (13). The reporter expresses an mRNA hybrid
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Fig. 3. RanBP3-mediated Smad2 export depends on its Ran-binding ability. The ability of RanBP3 or RanBP3-wv protein on Smad2 export is examined by in vitro export assay. In vitro export of GFP-Smad2 is progressive over time as indicated by a decreased level of GFP-Smad2 (lanes 1–3 ) in the permeabilized cells. RanBP3 (lanes 4 and 5 ) but not RanBP3-wv (lanes 6 and 7 ) promotes nuclear export of Smad2.
Fig. 4. Schematic diagram for MS2-based Nuclear Export Reporter Assay. In this cartoon, under normal conditions, the SD-CAT-MS2-SA RNA is only exported after splicing. In the presence of MS2–Smad2 fusion protein that can bind to the MS2 RNA, SD-CAT-MS2-SA RNA is exported and translated in the cytoplasm and CAT is then produced. SD splicing donor site, SA splicing acceptor site.
between CAT-encoding RNA and MS2 translation operator RNA, inserted between a pair of splicing donor/acceptor sites. Export of this hybrid mRNA depends on the specific recruitment of RNA-binding export factor to the unspliced CAT RNA. In the absence of export factor, CAT-encoding RNA is spliced out and no CAT is produced. (2) The second component is the MS2– Smad2 fusion protein, which contains a bacteriophage MS2 coat protein fused to Smad2 (12). MS2 coat protein can bind to MS2
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RNA. If the MS2 fusion partner undergoes nuclear export, the fusion protein binds to the unspliced CAT-MS2 RNA and transports the hybrid RNA into the cytoplasm, where the CAT RNA can be translated. Thus, production of CAT relies on the ability of Smad2, the fusion partner of MS2 coat protein, to undergo nuclear export. The activity of CAT quantitatively reflects the amount of Smad2 molecules exported. We use this assay to quantitatively determine the effect of RanBP3 and PPM1A on Smad2 nuclear export. 1. HEK293T cells at 30% confluency are cultured in 12-well plates. Two or three wells (duplicate/triplicate) are required for each experimental data point. 2. Transfect HEK293T cells with plasmids encoding MS2– Smad2, pDM128/8xMS2, pSVbgal (Promega) (see Note 10) together with plasmids encoding RanBP3 or PPM1A by using LipofectAmine transfection reagent (Invitrogen). 3. Forty-five hours after transfection, wash cells with PBS once, scrape cells into 1 ml PBS, harvest cells by centrifugation at 500 × g, 4°C, for 5 min. 4. Lyse the cell pellets with 200 µl lysis buffer on ice for 10 min. 5. Cell lysates are ready to be analyzed for CAT activity (150 µl of cell lysates)/b-galactosidase activity (20 µl of cell lysates) by using ELISA-based assay on a 96-well microreader according to manufacturer’s instructions. 6. Relative MS2–Smad2 export activity is calculated by normalizing the CAT activity with b-galactosidase activity in each sample. An example of the results produced is shown in Fig. 5. 3.5. In Vitro Phosphatase Assay
We found RanBP3 preferentially recognizes dephosphorylated Smad2/3, and PPM1A facilitates the interaction between RanBP3 and Smad2/3 in the nucleus (15). These data indicate that dephosphorylated Smad2/3 serve as a proper cargo for RanBP3. Although both PPM1A and RanBP3 promote nuclear export of Smad2 (Fig. 5), PPM1A dephosphorylates Smad2/3 (2) and appears to act upstream of RanBP3 in the process of Smad2 export. To rule out the possibility that decreased P-Smad2/3 levels may be attributed to PPM1A phosphatase activity toward upstream activators of Smad2/3 (e.g., TGF-b receptor kinases) instead of Smad2/3 themselves, it is necessary to examine PPM1A phosphatase activity in a cell-free system where only purified recombinant proteins are used. We use in vitro phosphatase assay to determine whether PPM1A directly dephosphorylates Smad2/3.
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Fig. 6. PPM1A dephosphorylates the phospho-SXS motif in the Smad2 MH2 domain (P-S2MH2). The phosphatase activity of PPM1A toward recombinant phosphorylated Smad2 MH2 domain (a) or immunoprecipitated phosphorylated Smad2 (b) is examined by in vitro phosphatase assay. The levels of PPM1A, Smad2-MH2, Smad2 as well as the phosphorylated Smad2-MH2 or Smad2 were examined by Western blot analysis. The results in panel (a) indicate that equal amounts of semi-synthetic recombinant P-S2MH2 (lanes 1–5 ) and recombinant PPM1A (lanes 2–5 ) are loaded. Dephosphorylation of P-Smad2MH2 by PPM1A requires metal ion (lanes 3 and 4 ) and is abolished by 40 mM EDTA (lane 5 ). Similar results were also observed by using immunoprecipitated full-length Smad2 as shown in (b).
1. To setup cell-free phosphatase reactions (see Note 11), add 100 ng of Escherichia coli-expressed, purified recombinant His-tagged PPM1A protein and 100 ng of semi-synthetic recombinant phospho-Smad2 MH2 (P-S2MH2) peptide (see Notes 12 and 13) in a 50 µl phosphatase reaction buffer. 2. Perform the phosphatase reaction at 30°C for 30 min. Stop the reactions by addition of 50 µl of 2× SDS loading buffer. 3. Examine the levels of total Smad2; P-Smad2 and PPM1A by Western blot analysis. Examples of the result are shown in Fig. 6.
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4. Notes 1. To evaluate the function of RanBP3 in Smad2/3 export and subsequent TGF-b signaling, stable HaCaT cell lines with depleted expression of RanBP3 are established. Several shRNAs against RanBP3, whose expression is controlled by RNA polymerase III promoter H1, are tested for their knockdown efficiency on RanBP3 expression. Three highly effective shRNAs are then selected to stably transfect HaCaT cells. Pools or multiple clones of each shRNA-transfected stable cells can be selected for confirmation of knockdown of RanBP3 expression and for subsequent analyses. 2. To allow the examination of Smad2/3 nuclear export, steps 1 and 2 induce the Smad2/3 nuclear accumulation in the cultured cells in a “synchronized” manner by stimulating the cells in the resting state with TGF-b. 3. SB431542 almost instantaneously inhibits TGF-b receptor I kinase activity. It effectively blocks TGF-b-induced Smad2/3 phosphorylation in the cytoplasm and nuclear accumulation thereafter (6). The redistribution of nuclear accumulated Smad2/3 during this period is mainly determined by Smad2/3 nuclear export. 4. The absolute nuclear or cytoplasmic Smad2/3 level is also determined by the total cellular Smad2/3 level. It is thus necessary to examine whether the altered experimental condition (in this case, the depletion of RanBP3 in HaCaT cells) affects the total Smad2/3 levels. 5. The assay relies on successful separation of the nuclear and cytoplasmic fractions, which could be confirmed by examination of proteins with known subcellular localization through Western blot analysis. The existence of GADPH in the nuclear fractions or Lamin A/C in the cytoplasmic fractions all indicates the potential problems with the cell fractionation. It is critical to evaluate the quality of cell fractionation before any subsequent analyses. 6. RanBP3-wv mutant (W352A/V353A) binds RanGTP 200fold less than wild-type RanBP3 in RanGAP protection assay (16) and is utilized here as RanBP3 Ran-binding mutant in comparison with wild-type RanBP3. 7. This assay could also be used to examine the endogenous Smad2 export in parental HaCaT cells. GFP-Smad2 allows direct visualization of nuclear GFP-Smad2 export process under fluorescence microscope, which is extremely convenient for optimizing the experimental conditions.
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8. Digitonin effectively solubilizes membrane proteins and permeabilizes the plasma membrane. It is thus very critical in this assay to avoid over-permeabilization that may disrupt the nuclear membrane and hence the functionality of nuclei. Proper cell membrane permeabilization could be evaluated by: (1) lack of GAPDH in the total lysates of permeabilized cells, indicating a clearance of all cytoplasmic proteins in permeabilized cells; (2) a comparable level of nuclear envelope marker Lamin A/C between digitonin treated and nontreated cells, indicating integrity of the nuclear envelope; (3) a comparable level of Smad2 before and immediately after the digitonin permeabilization (defined as time point 0 in the assay), as disruption of nuclear envelope would result in the loss of nuclear Smad2. For example, as shown in Fig. 2, treatment with digitonin at 30 or 60 ng/ml (but not 10 ng/ml) for 5 min is able to permeabilize the HaCaT cell membrane, as both dosages resulted in the substantial loss of cytoplasmic GADPH after washes. 9. RanBP3 and RanBP3-wv are prepared from recombinant GST-RanBP3 or GST-RanBP3-wv proteins by removing GST with Precission proteinase (Amersham) according to the manufacturer’s instruction. 10. pSVbgal expresses b-galactosidase under the control of the SV40 early promoter. Measurement of b-galactosidase activity from the samples allows normalization of transfection efficiency. 11. PPM1A is a member of PPM family serine/threonine phosphatase. Its activity depends on metal irons Mg2+ or Mn2+ and could be inhibited by chelating agents (e.g., EDTA). Avoid using chelating agents in buffer and/or recombinant protein elutes in this assay. 12. Please refer to a previous study (17) for the generation of phospho-Smad2 MH2 (241–462) in vitro. Of note, PPM1A can also dephosphorylate the C-terminal SSXS motif of Smad1/5/8, which is highly conserved in all R-Smads (18). 13. Alternatively, readers could use immunoprecipitated fulllength Smad2/3 proteins as the substrate for this assay (see Fig. 6b). To prepare the phosphorylated Smad2/3 proteins, transfect Flag-tagged Smad2/3 into HEK293T cells together with constitutively activated TbRI kinase. Phosphorylated Smad2/3 proteins could be immunoprecipitated with antiFlag antibodies and eluted with Flag-peptide (Sigma) according to the manufacturer’s instructions. Our lab uses rat TbRI-T202D (19). The mutation of threonine to aspartic acid at the position of amino acid 202 in rat TbRI renders it constitutively active and able to signal in a ligand-independent manner, which is similar to the analogous mutation in the human TbRI-T204D (20).
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Acknowledgments We thank members of Feng and Lin labs for their contributions to the original research and helpful discussion. The described research is supported by NIH grants (R01AR053591 and R01CA108454 to X.-H.F., R01DK073932 to X.L.) and a Leukemia and Lymphoma Society Scholar Award (X.-H.F.). References 1. Feng XH, Derynck R (2005) Specificity and versatility in tgf-beta signaling through Smads. Annu Rev Cell Dev Biol 21:659–693 2. Massague J, Seoane J, Wotton D (2005) Smad transcription factors. Genes Dev 19:2783–2810 3. Lin X, Duan X, Liang YY et al (2006) PPM1A functions as a Smad phosphatase to terminate TGFbeta signaling. Cell 125:915–928 4. Watanabe M, Masuyama N, Fukuda M, Nishida E (2000) Regulation of intracellular dynamics of Smad4 by its leucine-rich nuclear export signal. EMBO Rep 1:176–182 5. Pierreux CE, Nicolas FJ, Hill CS (2000) Transforming growth factor beta-independent shuttling of Smad4 between the cytoplasm and nucleus. Mol Cell Biol 20:9041–9054 6. Inman GJ, Nicolas FJ, Hill CS (2002) Nucleocytoplasmic shuttling of Smads 2, 3, and 4 permits sensing of TGF-beta receptor activity. Mol Cell 10:283–294 7. Nicolas FJ, De Bosscher K, Schmierer B, Hill CS (2004) Analysis of Smad nucleocytoplasmic shuttling in living cells. J Cell Sci 117:4113–4125 8. Schmierer B, Hill CS (2005) Kinetic analysis of Smad nucleocytoplasmic shuttling reveals a mechanism for transforming growth factor beta-dependent nuclear accumulation of Smads. Mol Cell Biol 25:9845–9858 9. Xiao Z, Brownawell AM, Macara IG, Lodish HF (2003) A novel nuclear export signal in Smad1 is essential for its signaling activity. J Biol Chem 278:34245–34252 10. Kurisaki A, Kose S, Yoneda Y, Heldin CH, Moustakas A (2001) Transforming growth factor-beta induces nuclear import of Smad3 in an importin-beta1 and Ran-dependent manner. Mol Biol Cell 12:1079–1091 11. Xu L, Chen YG, Massague J (2000) The nuclear import function of Smad2 is masked by SARA and unmasked by TGFbeta-dependent phosphorylation. Nat Cell Biol 2:559–562
12. Xu L, Kang Y, Col S, Massague J (2002) Smad2 nucleocytoplasmic shuttling by nucleoporins CAN/Nup214 and Nup153 feeds TGFbeta signaling complexes in the cytoplasm and nucleus. Mol Cell 10:271–282 13. Coburn GA, Wiegand HL, Kang Y, Ho DN, Georgiadis MM, Cullen BR (2001) Using viral species specificity to define a critical protein/RNA interaction surface. Genes Dev 15:1194–1205 14. Cullen BR (2004) Assaying nuclear messenger RNA export in human cells. Methods Mol Biol 257:85–92 15. Dai F, Lin X, Chang C, Feng X-H (2009) Nuclear export of Smad2 and Smad3 by RanBP3. Dev Cell 16:345–357 16. Englmeier L, Fornerod M, Bischoff FR, Petosa C, Mattaj IW, Kutay U (2001) RanBP3 influences interactions between CRM1 and its nuclear protein export substrates. EMBO Rep 2:926–932 17. Wu JW, Hu M, Chai J et al (2001) Crystal structure of a phosphorylated Smad2. Recognition of phosphoserine by the MH2 domain and insights on Smad function in TGF-beta signaling. Mol Cell 8:1277–1289 18. Duan X, Liang YY, Feng XH, Lin X (2006) Protein serine/threonine phosphatase PPM1A dephosphorylates Smad1 in the bone morphogenetic protein signaling pathway. J Biol Chem 281:36526–36532 19. Feng XH, Derynck R (1996) Ligandindependent activation of transforming growth factor (TGF) beta signaling pathways by heteromeric cytoplasmic domains of TGFbeta receptors. J Biol Chem 271:13123–13129 20. Wieser R, Wrana JL, Massague J (1995) GS domain mutations that constitutively activate T beta R-I, the downstream signaling component in the TGF-beta receptor complex. EMBO J 14:2199–2208
Chapter 8 Assessing Sequence-Specific DNA Binding and Transcriptional Activity of STAT1 Transcription Factor Thomas Meyer and Uwe Vinkemeier Abstract Continuous nucleocytoplasmic shuttling of signal transducer and activator of transcription (STAT) proteins is a key to understand their function as cytokine-responsive transcription factors. STATs enter the nucleus both by carrier-dependent and carrier-independent transport pathways, and it was previously shown that STAT1 exits the nucleus only after its prior enzymatic dephosphorylation by nuclear phosphatases. The identification of different transport pathways for unphosphorylated and tyrosine-phosphorylated STAT dimers was made possible by a combination of a diverse set of experimental approaches in the field of molecular biology. In the following, we will summarize some of the techniques that have been successfully used to decipher molecular mechanisms engaged in STAT1 dynamics. Key words: STAT proteins, Nucleocytoplasmic transport, Shuttling, DNA binding
1. Introduction Cytokines regulate the function of a variety of cells mainly by modifying transcriptional profiles. Transcription factors of the signal transducer and activator of transcription (STAT) family are the key downstream effectors of cytokine signaling that execute different biological responses (1–3). STAT1 and probably other family members as well form high-affinity dimers both before and after tyrosine phosphorylation (4). In the absence of cytokine stimulation the STATs are nucleocytoplasmic shuttling proteins that enter and leave the nucleus through direct interactions with nucleoporins in the nuclear pore complex, independent of transport factors (5). Cytokine signaling begins at the plasma membrane with the engagement of cytokines with their cognate receptors, which triggers autophosphorylation on tyrosines of
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noncovalently attached Janus kinases (JAKs). The activated JAK kinases then phosphorylate signature tyrosine residues in the intracellular receptor tails thus providing docking sites for the STAT Src homology 2 (SH2) domain. Phosphorylation on a single tyrosine residue at the carboxy-terminus of STATs causes a gross conformational change within the STAT dimers that shift from an “antiparallel” conformation to the “parallel” conformation that confers DNA binding (4, 6, 7). The phosphorylated STAT dimers then translocate via an importin-mediated import pathway in the nucleus, where they bind to high-affinity DNA recognition sites in the promoter region of target genes (8–11). These elements are termed gamma-activated sites (GAS) and contain the consensus sequence 5¢-TTC(N)3–4GAA¢-3. In the nucleus, the STATs are inactivated by dephosphorylation, for example, the nuclear tyrosine phosphatase Tc45 was identified to confer this inhibitory role for STAT1 (12–15). As dephosphorylation is required for subsequent nuclear export (16), and the enzymatic reaction is inhibited by DNA binding, cytokine stimulation of cells results in the transient accumulation of tyrosinephosphorylated STATs in the nucleus due to the slight delay in nuclear export (17). In humans seven different members of the STAT family are expressed (STAT1, STAT2, STAT3, STAT4, STAT5a, STAT5b, and STAT6), all of which are organized in the same modular domain arrangement. The STATs consist of three proteolytically separable structural subunits, an amino-terminal domain, a central core fragment, and a carboxy-terminal transactivation domain (18–20). The amino-terminal domain of about 130 residues consists of a unique hook-shaped architecture. This domain is required for dimerization of unphosphorylated STAT1 and facilitates oligomerization of phosphorylated STATs on DNA (16, 21–23). The large core domain encompasses several distinct functional domains beginning amino-terminally with a four-helix bundle, which is engaged in interactions with other proteins (24). The DNA-binding domain exhibits an immunoglobulin fold and is required for binding of tyrosine-phosphorylated STAT dimers to DNA (25). The consecutive linker region consists of a unique all-alpha helical structure and assists in DNA binding and nucleocytoplasmic shuttling (26, 27). The SH2 domain mediates the recruitment to phosphorylated tyrosine residues in the intracellular receptor molecules and allows the formation of tyrosinephosphorylated STAT dimers (21, 28). The carboxy-terminus contains the transactivation domain, which is most divergent in size and sequence between the different STAT family members. It is frequently subject to alterative splicing (29). Best described is the complex role of STAT1 in regulating the transcription of numerous genes that are instrumental in controlling cell growth, proliferation, and cellular responses to microbial
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and viral infections (30). The pathway leading to activation by type II interferons begins when interferon-g binds to its receptor IFNGR that catalyzes activation of JAK1 and JAK2 and results in the formation of STAT1 homodimers (31). Activation of STAT1 by type I interferon such as interferon-a is initiated by ligand binding to the IFNAR receptor, resulting in the auto- and transphosphorylation of JAK1 and TYK2. Subsequently, heterodimers of STAT1–STAT2 are formed that associate with interferon regulatory factor 9 (IRF-9, also called p48) to form the transcriptionally active interferon-stimulated gene factor 3 (ISGF3) (24). In line with the role of STAT1 as a key player in antimicrobial defense, STAT1-knockout mice exhibit a severe immune defect with impaired interferon signaling resulting in a high susceptibility to viral infections (32, 33). Much of what has been learned in the past decade about the highly dynamic STAT1 signaling resulted from experiments with recombinant STAT1 fusion proteins with green fluorescent protein (GFP). Although we have recently shown that nucleocytoplasmic shuttling of unphosphorylated STAT1 is significantly reduced after fusion to GFP (34), GFP-tagging nevertheless remains a valuable tool to investigate the biological effects of mutant STAT1 in terms of intracellular mobility and localization. In the following, we present a selection of readily applicable protocols that have been successfully used to study interferon signaling. In particular, we focus on cell biological methods that have offered mechanistic insight into the dynamic regulation of JAK– STAT signal transduction and describe them in more detail here.
2. Materials 1. U3A cells, derived from the human fibrosarcoma cell line 2fTGH. 2. U3A cells engineered to express STAT1 allow for the study of mutant STAT1 in the absence of an endogenous STAT1 background. 3. HeLa cells. 4. Cytoplasmic extraction buffer: 10 mM KCl, 0.2% Nonidet P40, 1 mM EDTA, 10% glycerol, 20 mM Hepes, 50 mM NaF, 1 mM vanadate, 1 mM DTT, 0.1 mM PMSF, and complete protease inhibitors from Roche, pH 7.4. 5. Nuclear extraction buffer: 420 mM KCl, 1 mM EDTA, 20% glycerol, 20 mM Hepes, 50 mM NaF, 1 mM vanadate, 1 mM DTT, 0.1 mM PMSF, and complete protease inhibitors, pH 7.4.
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6. Extraction buffer: 20 mM MES, 100 mM KCl, 10 mM NaF, 10 mM Na2HPO4/NaH2PO4, pH 7.0, 10 mM NaPPi, 0.02% NaN3, 4 mM EDTA, 1 mM EGTA, 20 mM DTT, and complete protease inhibitors. 7. Transport buffer: 20 mM Hepes, pH 7.3, 110 mM KOAC, 2 mM Mg(OAC)2, 1 mM EGTA, 2 mM DTT, and complete protease inhibitors. 8. Digitonin transport buffer: 10 mM KCl, 1.5 mM MgCl2, 10 mM Hepes, pH 7.4, 0.2% Triton X-100, 1 mM DTT, and complete protease inhibitors. 9. Lysis buffer A: 50 mM Tris–HCl, pH 8.0, 1% NP-40, 5 mM EDTA, 2 mM EGTA, 10% glycerol, 50 mM NaF, 0.1 mM vanadate, 400 mM NaCl, 3 mM DTT, 0.1 mM PMSF, and complete protease inhibitors.
3. Methods 3.1. Cell Culture
Elucidating biological fundamentals in the JAK–STAT pathway has benefited much from complementation experiments in the cell line U3A, which was shown to be defective in interferon signaling due to the loss of STAT1 expression (35). 1. U3A cells as well as HeLa cells are cultured at 37°C in a humidified 5% CO2 atmosphere in 10% Dulbecco’s modified Eagle’s medium supplemented with 10% fetal calf serum (FCS) (Gibco), 1% penicillin, and 1% streptomycin. 2. Transfection of cells with STAT1 expression vectors is routinely done with Lipofectamine plus (Gibco) according to the manufacturer’s recommendation. 3. Twenty-four hours after transfection, cells are either left unstimulated or stimulated with 5 ng/ml human interferon-g (Biomol) for 30 min to induce tyrosine phosphorylation and nuclear accumulation of STAT1 (see Note 1).
3.2. Plasmid Construction
1. The mammalian expression plasmid pSTAT1–GFP coding for full-length human STAT1 fused carboxy-terminally to GFP has been widely used to study nuclear retention of STAT1. This plasmid was originally created by PCR amplification of a STAT1 cDNA coding for the amino acid residues 1–746 with the primer pair 5¢-ATATATGAATTCATGTCTCAGTGGTA CGAACTTCAG-3¢ and 5¢-ATATATGGATCCATCATACT GTCGAATTCTAC-3¢ (36). 2. After cleavage with EcoRI and the internal SmaI site (EcoRI and BamHI restriction sites underlined) of the resulting PCR
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product, the DNA fragment comprising the amino-terminal part of STAT1 was introduced into the corresponding sites of pEGFPNI. 3. The vector that resulted from this cloning step was ligated into its BamHI/SmaI sites with a DNA-fragment representing the residual carboxy-terminal part of STAT1 to generate pSTAT1–GFP. 4. Wild-type and mutant STAT1 gene constructs cloned in the expression vector pcDNA3 (Invitrogen) are used for expression of untagged STAT1. Mutations in these expression vectors are introduced by site-directed point mutagenesis using the QuikChange kit from Stratagene, as recommended by the manufacturer. Mutations are verified by standard didesoxy termination DNA sequencing. 3.3. Pharmacological Suppression of STAT Signaling
In cultured cells, staurosporine (Sigma) functions as a highly potent inhibitor of JAK kinases, when used at a concentration of 500 nM (37). 1. For staurosporine treatment, activation of STAT1 is induced by stimulating cells with interferon-g for at least 30 min, and subsequently replacing the medium with medium containing staurosporine to block further tyrosine kinase activity. Exposure of interferon-prestimulated cells to staurosporine results in the rapid breakdown of STAT1 tyrosine phosphorylation and nuclear accumulation (see below). Contrarily, inactivating tyrosine phosphatases by treatment of cells with pervanadate (38) suppresses STAT1 dephosphorylation and hence extends the duration of nuclear accumulation. 2. In order to test for the contribution of carrier-mediated nuclear export to STAT signaling, cells must be treated with 10 ng/ml leptomycin B (Sigma) or ratjadone A (Alexis). These actinobacterial cytotoxins efficaciously inhibit the export receptor chromosomal region maintenance 1 (CRM1) and thus prolong STAT1 nuclear accumulation (36, 39, 40). However, treatment with these compounds merely delays the cessation of nuclear accumulation, demonstrating the presence of CRM1-independent export.
3.4. Western Blotting
1. Cells grown on six-well dishes are lysed in 30 ml/well of cytoplasmic extraction buffer for 5 min on ice. The lysates are centrifuged at 16,000 × g for 10 s at 4°C. 2. The supernatants are recentrifuged for 5 min at 16,000 × g and the pellets resuspended in 30 ml of nuclear extraction buffer. 3. After 30 min on ice the nuclear lysates are centrifuged for 15 min at 16,000 × g and the combined cytoplasmic and
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nuclear extracts are then boiled in sodium dodecyl sulfate (SDS) sample buffer. 4. Proteins are resolved by 10% SDS-polyacrylamide gel electrophoresis and subsequently transferred to nitrocellulose membranes according to standard techniques. 5. Phospho-STAT1 is detected by incubating the membranes with a rabbit polyclonal antibody specifically recognizing tyrosine-phosphorylated STAT1 (Cell Signaling) followed by a horseradish peroxidase-conjugated secondary antibody (Dako) with three washes in between. 6. Blots are stripped by incubation for 60 min at 60°C in 62.5 mM Tris–HCl, pH 6.8, 2% SDS, and 0.7% b-mercaptoethanol and reprobed with polyclonal STAT1-specific antibody C-24 (Santa Cruz) followed by incubation with a horseradish peroxidase-conjugated secondary antibody. Bound immunoreactivity is again detected using the enhanced chemiluminescence reaction (NEN) (Fig. 1). 3.5. Fluorescence Microscopy
Fluorescence microscopy is an indispensable technique in the study of STAT1 nucleocytoplasmic shuttling. The intracellular distribution of STAT1 in interferon-stimulated cells can best be monitored in cells expressing GFP fusions of STAT1 (34, 41–43). 1. For fluorescence microscopical detection of GFP-tagged STAT1 proteins, transiently transfected cells are treated with interferon-g as described and subsequently followed by either live-cell imaging techniques including as laser scanning microscopy or direct epifluorescence microscopy after fixation of cells for 15 min at room temperature (RT) in 3.7% paraformaldehyde diluted in phosphate-buffered saline (PBS). 2. The nuclei of fixed cells are additionally stained for 3 min with 5 mg/ml Hoechst 33258 (Sigma) and the samples are
Fig. 1. Kinetics of tyrosine phosphorylation in interferon-g-stimulated U3A cells expressing recombinant STAT1 proteins. U3A cells were transiently transfected with STAT1 expression plasmids coding for either wild-type (WT) or a DNA-binding mutant with two glutamyl residues at positions 428 and 429 mutated to alanines (EE/AA) (25). Twentyfour hours after transfection equal numbers of cells were stimulated with 5 ng/ml human interferon-g for 0, 30, 60, and 150 min, respectively. Western blots from whole cells extracts incubated with a phosphotyrosine-specific anti-STAT1 antibody (top panels) and their corresponding reblots after stripping off bound immunoreactivity and reincubation with a pan-STAT1 antibody (bottom panels) are shown.
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Fig. 2. Kinetics of STAT1 nucleocytoplasmic redistribution in interferon-g-prestimulated cells as determined by wide-field fluorescence microscopy. HeLa cells expressing green-fluorescent proteins fused amino-terminally to wild-type (WT) STAT1 (top panel ), the DNA-binding mutant EE/AA (middle panel ) or a mutant with a truncated amino-terminus (bottom panel ) were stimulated for 30 min with 5 ng/ml interferon-g. Cells were then immediately fixed (0.5 h) or the medium was replaced before incubation for additional 4 and 8 h in the absence of cytokines (4.5 and 8.5 h, respectively). Note that STAT1–WT is still accumulated 4 h after termination of interferon treatment, while the EE/AA mutant already showed a pancellular resting distribution at that time due to decreased DNA-binding affinity (17). The amino-terminal deletion mutant completely failed to accumulate upon cytokine stimulation, but conversely showed a depletion of nuclear localization. Thus, an intact amino-terminal domain of STAT1 is required for cytokine-induced nuclear import of tyrosinephosphorylated STAT1 (44).
mounted in fluorescence mounting medium (Southern Biotech). Fluorescence microscopy is performed on a confocal or conventional microscope equipped with appropriated fluorescence filters. Images are taken with a CCD camera and further processed with imaging software for visualization and quantification of fluorescence signals (Figs. 2 and 3). 3.6. Immunocytochemistry
1. Immunocytochemical detection of recombinant STAT1 is done in U3A cells expressing either wild-type or mutant STAT1. 2. Intracellular localization of native STAT1 in HeLa cells is determined by means of indirect immunocytochemistry using the following protocol. Adherent cells grown on chamber slides are stimulated with interferon-g for 30 min and either immediately fixed or additionally incubated in the presence of 500 nM staurosporine for additional 60 or 90 min. The cells are then fixed with chilled methanol for 20 min at −20°C. 3. After two washes in PBS the cells are permeabilized with 1.0% Triton X-100 in PBS and nonspecific binding is blocked by incubation with 25% FCS in PBS for 30 min at RT. 4. The samples are then incubated for 45 min with anti-STAT1 antibody C-24 (Santa Cruz) diluted 1:1,000 in 25% FCS/PBS. After three washes in PBS bound immunoreactivity is detected with Cy3-conjugated secondary antibodies (Dianova) for 45 min at RT followed by nuclear staining with Hoechst dye.
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Fig. 3. Time series of interferon-g-treated cells expressing a GFP-tagged fusion of a STAT1 deletion mutant lacking the STAT1 amino-terminus. The images show the redistribution of STAT1-DN-GFP before (0) and up to 90 min (90) after the addition of 5 ng/ml interferon-g to the cells as observed by confocal laser scanning microscopy. Note that amino-terminally deleted STAT1 showed a nuclear import defect upon cytokine stimulation and, conversely to the wild-type, accumulated in the cytosol.
5. The samples are mounted in fluorescence mounting medium (Southern Biotech) and images are taken by fluorescence microscopy (Fig. 4). For quantification STAT1 immunofluorescence densities are measured separately in the nucleus and in the cytoplasm using confocal microscopy. Correction for background fluorescence should be routinely done. For a number of cells (usually n = 20) the ratio of [cytoplasmic fluorescence density]/[nuclear fluorescence density] is calculated and values are depicted as means and standard deviations. Student’s t-tests are used to compare different STAT1 mutants or stimulation conditions. 3.7. Antibody Microinjection Assay
Antibody microinjection assays have been used to reveal the constitutive nucleocytoplasmic shuttling of unphosphorylated STAT1 (26, 45). 1. HeLa cells grown on glass coverslips are injected with injection solution containing 0.1 mg/ml anti-STAT1 antibody C-136 and 0.5 mg/ml fluorescein isothiocyanate-labeled
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Fig. 4. Vanadate-induced hyperphosphorylation did not restore the defective nuclear accumulation of an amino-terminal point mutant (W37A) of STAT1. U3A cells expressing wild-type (WT) STAT1 or the W37A mutant that results in aminoterminal deletion were stimulated with 5 ng/ml interferon-g for 0, 0.5, and 3.5 h, respectively. Where indicated, 0.8 mM vanadate and 0.2 mM H2O2 were added to the cells. (a) Shown is a Western blot with tyrosine-phosphorylated STAT1 and the corresponding reblot after stripping and reprobing with a pan-STAT1 antibody. Note that treatment with the phosphatase inhibitor vanadate led to hyperphosphorylation of both STAT1 variants and that the W37A mutant displayed a structural defect with a decreased molecular weight. (b) Gelshift experiments from the same extracts as used in (a) demonstrating increased DNA-binding activity of both STAT1 proteins in cells treated with vanadate/H 2O 2. (c) Immunocytochemical staining of STAT1 in U3A cells expressing STAT1–WT or the W37A mutant that were treated as described above. The results show that the amino-terminally deleted W37A mutant exhibited a nuclear import defect that cannot be overcome by inhibiting phosphatase activity.
bovine serum albumin (FITC-BSA, Sigma) to mark the injection site (see Note 2). 2. Antibodies are administered in the cell by using preformed capillaries (Femtotips, Eppendorf) attached to the Transjector 5246 and Micromanipulator 5171 (Eppendorf). Typically, 20 cells are injected within 10 min using a maximal pressure of 40 hPa. 3. After injection the cells are incubated at 37°C, before fixation and immunocytochemical antigen detection using Cy3conjugated anti-mouse immunoglobulins.
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Fig. 5. Mutation of a critical serine residue at position 727 of the STAT1 molecule reduces its nuclear export rate as demonstrated by antibody microinjection. HeLa cells expressing green fluorescent protein (GFP) fused to wild-type STAT1 or the Ser727Ala mutant were prestimulated for 30 min with 5 ng/ml interferon-g to induce nuclear accumulation of STAT1– GFP. Anti-STAT antibody C-136 (0.2 mg/ml) was injected into the cytosol of STAT1–GFP-expressing cells together with the injection site marker fluorescein isothiocyanate-labeled bovine serum albumin (FITC–BSA, 0.5 mg/ml). Ninety minutes after injection cells were fixed and stained with Hoechst dye. Note that the injected STAT1-specific antibody induced the collapse of nuclear accumulation of wild-type STAT1–GFP, while the predominant nuclear distribution of STAT1-Ser727AlaGFP was unchanged. For additional information on the phenotype of this nuclear retention mutant see (44).
4. A Zeiss inverted microscope (Axiovert 25) equipped with UV light emission can be used for detection of immunofluorescence pattern (Fig. 5). 3.8. Purification of Recombinant STAT1
Purified recombinant phospho-STAT1 has been used in microinjection assays to reveal the nuclear export block of tyrosinephosphorylated STAT1 (17). Additionally, purified STAT1 is required in import assays and in vitro dephosphorylation assays (5, 17). Our purification protocol of full-length and truncated STAT1 is based on the Bac-to-Bac system in baculovirus-infected Sf9 insect cells as specified by the manufacturer (Gibco) using Baculovirus-expressing STAT1 variants. Protein purification and in vitro tyrosine phosphorylation of truncated STAT1 are done as described (22, 46). In the following, we briefly report the purification strategy for full-length STAT1 (see Note 3). 1. Cells are lysed in ice cold extraction buffer and the lysates are cleared by centrifugation at 20,000 × g for 30 min. 2. The supernatant is brought to pH 6.2 with 1 M MES and after the addition of 0.5 vol of buffer 1 (20 mM MES, 0.02% NaN3, 20 mM DTT, pH 6.0) is again centrifuged for 20 min at 25,000 × g. Affinity chromatography on S-Sepharose and Q-Sepharose columns and the subsequent alkylation reaction are done as described (22). Alternatively, purification is done via the eight amino acids long Strep-affinity tag, which was engineered to the STAT1 C terminus (4).
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3. For tyrosine phosphorylation the purified STAT1 is incubated with a preparation containing epidermal growth factor receptor (EGFR) kinase that was isolated from A431 cells by immunoprecipitation. Fluorescence labeling is done with a succinimidyl ester of Oregon green 488 (Molecular Probes) used at a protein/dye molar ratio of 1:5 as described by the manufacturer; for additional information see (5). 4. Purified STAT1 proteins are microinjected into mammalian cells at a concentration of 1 mg/ml (17, 26). 3.9. Import Assays with Permeabilized Cells
Import assays have been successfully employed to study molecular mechanisms in nuclear import of STAT1 (5). This cell transport assay requires purified recombinant STAT1 and digitonin-permeated cells. Treatment with digitonin solubilizes the plasma membrane thereby releasing cytosolic proteins, but leaves the integrity of the nuclear envelope intact (47). Using import assays we have shown that nuclear import of unphosphorylated STAT1 and STAT3 does not require metabolic energy or added soluble transport factors, but occurs through direct contacts of STAT1 with nucleoporins in the nuclear pore complex (5). In contrast, tyrosine-phosphorylated STAT1 is incapable to enter the nucleus in the absence of added cytosol and adenosine 5¢-triphosphate. The import assay is performed as follows: 1. Adherent HeLa-S3 cells are grown on coverslips and permeabilized with 40 mg/ml digitonin (Roche) in transport buffer for 6 min on ice (47). 2. Cells are washed twice in transport buffer followed by incubation with 20 ml import mix at RT for 1 h. The cytosol-free import mix contains transport buffer supplemented with 10 mg/ml BSA and 1 mM of purified STAT1 protein labeled with Oregon green 488 (5). 3. For ATP depletion, import mix is preincubated for 15 min on ice in the presence of 0.8 U/ml apyrase (Sigma) before addition to the permeabilized cells. 4. To demonstrate translocation through the nuclear pore, cells are preincubated with 250 mg/ml wheat germ agglutinin (WGA). Treatment with this lectin inactivates proteins in the nuclear pore and prevents passage through the nuclear envelope (48). 5. When the import reaction is accomplished, cells are washed with ice-cold transport buffer and fixed for 15 min at RT with 3.7% paraformaldehyde in PBS, before permeabilization with 0.2% Triton X-100 in PBS (2 min at RT). 6. Finally, samples are mounted in TB containing 50% glycerol, 1% (w/v) NaN3, and 1% (w/v) diazobicyclo (2, 2, 2)-odone and STAT proteins imported into the nucleus are detected by fluorescence microscopy (Fig. 6).
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Fig. 6. Import assay demonstrating energy-independent nuclear import of purified unphosphorylated STAT1 through the nuclear pore complex. HeLa cells were treated with 40 mg/ml digitonin to permeate the plasma membrane, subsequently washed twice in transport buffer, and incubated with 20 ml of transport buffer containing 10 mg/ml bovine serum albumin and 1 mM truncated Oregon green 488-conjugated STAT1 (STAT1-tc). Depletion of metabolic energy was done by incubating digitonin-permeabilized cells with 0.8 U/ml apyrase. To demonstrate passage through the nuclear pore, permeabilized cells were incubated with 250 mg/ml of wheat germ agglutinin (WGA) for 15 min on ice. For additional experimental details see ref. 5.
3.10. Digitonin Assay for Assessing Increased DNABinding Activity
The influence of STAT DNA-binding activity on nuclear retention can be assessed by digitonization of interferon-g-prestimulated HeLa cells that expressed GFP-tagged STAT1 mutants. Treatment with digitonin permeates the plasma membrane and washes away cytosolic components, but preserves the structural integrity of the nuclear envelope. STAT1 mutants with increased DNA-binding affinity resist digitonin treatment while in a DNAbound state that precludes nuclear export, while the majority of the wild-type STAT1–GFP had already exited the nuclear compartment after its dephosphorylation. The digitonization assay is performed in the following way: 1. Nuclear accumulation of STAT1–GFP is achieved by stimulation of adherent HeLa cells with interferon-g. 2. Cells are either left untreated or permeabilized in the presence of 50 mg/ml digitonin (Roche) in digitonin transport buffer for 6 min on ice. 3. After two washes in transport buffer, the cells are fixed for 15 min at RT with 3.7% paraformaldehyde in PBS followed
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Fig. 7. Digitonin assay demonstrating increased nuclear retention of the STAT1 mutant dnaplus, which contains three amino acid substitutions: Thr327Arg, Val426His, and Thr427His. HeLa cells expressing STAT1–WT–GFP (WT) and STAT1– dnaplus–GFP (DNA+) were stimulated with interferon-g to induce nuclear accumulation of STAT1. In nonpermeabilized cells (left panels), both WT and mutant STAT1 are localized in the nucleus. All images show an overlay of the GFP fluorescence (green), indicating the localization of STAT1, and the nuclear staining with Hoechst dye (blue). Note that approximately only 50% of the cells were expressing STAT1–GFP. Treatment with 50 mg/ml digitonin to permeate the plasma membrane (middle panel) resulted in the loss of WT from the nucleus, whereas the dnaplus mutant remained nuclear. Preincubation of the living cells with vanadate/H2O2 before permeabilization to preserve the phosphorylation of STAT1 (right panel), however, blocked the loss of WT STAT1 from the nucleus. The latter experiment confirmed the inability of activated STAT1 to exit the nucleus.
by staining with Hoechst dye. The presence of STAT1–GFP in the nuclei is probed by means of direct fluorescence microscopy (Fig. 7). 3.11. Dephosphorylation Assays
1. In vivo dephosphorylation assays are performed with U3A cells transiently expressing wild-type or mutant STAT1. 2. Cells grown on 6-cm dishes are transferred to six-well plates 24 h posttransfection and cultivated for another 24 h, before stimulation with interferon-g for 30 min. 3. The medium is replaced by medium without or with 500 nM staurosporine; treatment with the kinase inhibitor is for 30–90 min. 4. Cells are washed in PBS and lysed in lysis buffer A before the extracts are analyzed by Western blotting (Fig. 8). 5. In vitro dephosphorylation assays have been used to demonstrate that DNA-bound STAT1 is protected from enzymatic dephosphorylation (17). The reactions are performed at 30°C with 1 nM of purified STAT1 dimers and 1.5 U of the truncated human Tc45 phosphatase (residues 1–352, Sigma) or 15 U of the full-length Tc45 phosphatase. The total reaction volume is 20 ml in a reaction buffer containing 25 mM
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Fig. 8. In vivo dephosphorylation assay using staurosporine to suppress STAT1 tyrosine phosphorylation and to trigger the concomitant collapse of STAT1 nuclear accumulation. Cells expressing STAT1-WT-GFP or the DNA-binding mutant EE/AA, which is devoid of sequence-specific DNA binding (25), were prestimulated with 5 ng/ml interferon-g and subsequently exposed to 500 nM staurosporine for the indicated times. Shown are Western blot results (a) with U3A cell extracts incubated with anti-phospho-STAT1 and reprobed with a pan-STAT1-antibody; and immunofluorescence micrographs (b) from transfected HeLa cells demonstrating the breakdown of nuclear accumulation within 30 min of staurosporine exposure (compare to Fig. 2). Note the indiscriminate behavior of WT and mutant STAT1, indicating the minor contribution of sequence-specific DNA binding to nuclear retention.
Tris–HCl, pH 7.5, 0.5 mg/ml BSA, 10 mM DTT, 50 mM KCl, 5 mM EDTA, and complete protease inhibitors (Roche). 6. Recombinant full-length Tc45 is isolated from BL21pLysS bacteria and purified by virtue of a carboxy-terminal Strep-tag as has been described (17). Increasing concentrations of a duplex 37-mer DNA containing either a single GAS site (GAS) or an unrelated sequence (Mut) are included in the reaction (GAS: 5¢-AAGTCGTTTCCCGGAAATAGAAGATTATTATCA TTAT and its antisense; Mut: 5¢-AAGTCGAGGTACAGGT AAAGAAGAACCTCGTTGTCAC and its antisense) with DNA concentrations varying from 0.5 to 25 nM. 7. The in vitro reactions are stopped after 60 min by boiling in SDS sample buffer and analyzed by Western blotting. 3.12. Fluorescence Recovery After Photobleaching
Fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP) have been used to measure the intracellular mobility of fluorescently tagged molecules (43, 49, 50). 1. In a typical FRAP experiment, a GFP-tagged protein within a small target region is subjected to intense illumination, applied by a short pulse of high-intensity laser, which leads to a nearly
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complete photobleaching of all GFP-fusion molecules in this region of interest (ROI). 2. Following the photobleaching pulse, the initially low fluorescence intensity increases owing to the repopulation by unbleached molecules from regions outside the bleached spot. 3. The rate and extent of fluorescence intensity recovery in the bleached region is monitored as a function of time to estimate the mobile fraction of fluorescently labeled protein. We have performed FRAP experiments in STAT1–GFP-expressing HeLa cells before or 60 min after interferon-g addition using an LSM510 inverted confocal laser scanning microscope (Carl Zeiss Jena) equipped with a ×100/1.3 objective. The photobleaching pulse is applied by an argon laser (l = 488 nm), and fluorescence images are usually scanned as a time series with 250 ms intervals. 4. The relative fluorescence intensities in a bleached ROI (10 mm2) are measured over time. Fluorescence intensity differences (Fu − Fb) between unbleached ROI (Fu) and adjoining bleached ROI (Fb) are then calculated and used for an exponential curve fitting (prism program) to determine recovery times. Usually, recovery is considered complete if Fu − Fb £ 0.01 Fu (Fig. 9).
Fig. 9. Inhibition of tyrosine phosphatase activity dramatically decreased the intranuclear mobility of STAT1 as measured by fluorescence recovery after photobleaching (FRAP). HeLa cells expressing GFP-tagged wild-type STAT1 were either left untreated (black curve, composite of four independent experiments) or treated with 5 ng/ml interferon-g in the absence (red curve, composite of five independent experiments) or presence of 0.8 mM vanadate/0.2 mM H2O2 (green curve, composite of five independent experiments). Fluorescence intensity differences between unbleached regions of interest (ROIs) and adjoining bleached ROIs were calculated and used to determine recovery times as described in the text. The means and standard deviations of the recovery times are given below the plotted curves.
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3.13. Pull-Down Assay with Biotinylated Oligos
Pull-down assays as described here have been developed to investigate the differential sequence requirements of STAT1 DNA-binding mutants (17). 1. Duplex oligonucleotides (0.5 ml, 28 pmol/µl, with a 5¢-biotin on one strand) containing either two optimal GAS sites [ 5 ¢- G A G A C T C A G T T T C C C G TA A AT C G T C C A G T TTCCCGTAAAGACTATGC-3¢ and its antisense] or a single GAS-like site [5¢-(TTC)4TAC(TTC)15-3¢ and its antisense] were conjugated to streptavidin agarose (0.5 ml packed vol, Pierce) for 1 h at 4°C. 2. Confluent U3A cells (one 10-cm dish) transiently expressing STAT1–GFP variant were stimulated for 30 min with interferong and for another 15 min in the additional presence of vanadate/ H2O2 before the cells were lysed in 400 ml lysis buffer A (17). 3. After preclearing with streptavidin agarose, 250 ml of the normalized extracts are rotated with 50 µl (packed vol) of DNAconjugated beads for 90 min at 4°C. 4. The beads are washed three times with 500 ml lysis buffer. 5. DNA-bound proteins are eluted by boiling in SDS sample buffer and analyzed by Western blotting (Fig. 10).
3.14. Electrophoretic Mobility Shift Assay
1. U3A cells are transfected with expression plasmids coding for wild-type and mutant STAT1, respectively, and allowed to recover for 24 h after transfection. 2. Cells are either left untreated or treated with 5 ng/ml interferon-g for different time periods.
Fig. 10. Pull-down assay for assessing sequence-specific and sequence-unspecific DNA binding of WT and mutant STAT1. U3A cells were transfected with expression plasmids coding for either wild-type STAT1 (WT) or the DNA-binding mutants STAT1-dnaplus (DNA+) and STAT1-dnaminus (DNA−), respectively. Twenty-four hours later cells were stimulated with 5 ng/ml interferon-g for 35 min followed by the additional presence of vanadate/H2O2 for 15 min. Increasing amounts of lysates (5, 10, and 15 ml, respectively) were loaded on a gel and detected by immunoblotting using a phosphotyrosine-specific STAT1 antibody to confirm comparable loading (top panel). Equal amounts of lysates were incubated with agarose-bound oligonucleotides comprising tandem M67 binding sites (GAS) or an unrelated sequence (Mut). Unmodified agarose beads served as a control (w/o). The precipitates were analyzed by Western blotting with anti-STAT1 antibody C-24 (17).
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Fig. 11. Competition gel shift assay used to demonstrate defective tetramerization of an amino-terminally deleted STAT1 mutant (DN) and the impaired DNA-binding affinity of the double glutamine to alanine mutant EE/AA. Shown are cell extracts from untransfected HeLa cells (lane 2 ) and U3A cells (lanes 1, 3–8 ) expressing GFP fusions of wildtype STAT1, DN, and EE/AA, respectively. The extracts were incubated with [32P]-labeled oligonucleotides containing a tandem GAS site. Where indicated, a 750-fold molar excess of an unlabeled single GAS site was added to the reaction (lanes 4, 6, and 8 ) before they were loaded on the gel. Note that tetrameric STAT1–WT (marked with T at the right margin), but not STAT1-W37A, resisted displacement when challenged by excess unlabeled oligonucleotides, which is indicative for an intact cooperative DNA binding. The EE/AA mutant had completely lost its ability to bind to GAS sites.
3. Cytoplasmic and nuclear cell extracts are prepared and combined as described above. 4. Four microliters of each extract are incubated with 1 ng of duplex oligonucleotides, which have been labeled with [32P] by end-filling reaction using Klenow fragment (New England Biolabs). 5. The EMSA reactions are equilibrated for 15 min at RT before loading on a 4% 29:1 acrylamide:bisacrylamide gel at 4°C (36). In competition experiments used to determine the off-rate from DNA, a 750-fold molar excess of an unlabeled high-affinity single GAS site, termed M67, is added to the reaction. 6. Reactions are incubated at RT for 15 min before loading on the gel. In supershift assays, 20 ng of STAT1-specific antibody C-24 is preincubated with the shift reaction for 15 min at RT. DNA-binding activity is visualized with a phosphoimaging system on vacuum-dried gels (Fig. 11). 3.15. Reporter Gene Assay
1. U3A cells grown on 48-well plates are transiently transfected with the following amounts of cDNAs added into a single well: 250 ng of the respective STAT1 expression plasmid, 70 ng of a constitutively expressed b-galactosidase reporter (Stratagene), and 200 ng of an interferon-g-sensitive reporter gene containing a triple Ly6E STAT-binding site in front of a luciferase reporter (4).
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Fig. 12. The STAT1-W37A mutant fails to induce a luciferase reporter gene upon stimulation of cells with interferon-g. U3A cells expressing wild-type STAT1 or the W37A mutant thereof were either left untreated (grey columns) or stimulated for 6 h in the continuous presence of 5 ng/ml interferon-g (black columns) before cell extracts were obtained. Expression of luciferase, which is under the control of a promoter with triple Ly6E STAT-binding sites, was normalized to the expression of a constitutively expressed b-galactosidase reporter. Shown are means and standard deviations of six independent experiments.
2. Twenty-four hours posttransfection, cells are either left untreated or treated for 6 h with interferon-g. 3. Whole cell extracts are prepared with a lysis buffer containing 25 mM glycylglycine, 1% Triton X-100, 15 mM MgSO4, 4 mM EGTA, 1 mM DTT, pH 7.8, and complete protease inhibitors. Luciferase (Promega) and b-galactosidase activities are measured spectroscopically. The data are normalized for the expression of b-galactosidase. Routinely, for each STAT1 variant and stimulation mode six independent samples have to be tested and the experiment is usually repeated at least three times (Fig. 12).
4. Notes 1. Unless stated otherwise, all solutions should be prepared in deionized water. 2. Antibodies used in antibody-microinjection assays should be tested in immunoprecipitation assays to confirm reactivity with their native antigens. Injection and detection antibodies have to be from differed species to avoid cross-reactivity. 3. For storage, solutions containing recombinant STAT1 or lysates from STAT1-expressing cells should be snap-frozen in liquid nitrogen or on dry ice and kept at −80°C. During thawing the temperature should not rise significantly above 0°C to avoid STAT1 denaturation.
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Acknowledgments We thank Petra Lemke, Leibniz-Forschungsinstitut für Molekulare Pharmacology, Berlin, for providing us with Fig. 10. Financial support was provided by the Leibniz-Institut für Molekulare Pharmakologie, and grants VI 218/4 and ME 1648/2-1 from Deutsche Forschungsgemeinschaft to U.V. and T.M., respectively. References 1. Darnell JE Jr (1997) STATs and gene regulation. Science 277:1630–1635 2. Brivanlou AH, Darnell JE Jr (2002) Signal transduction and the control of gene expression. Science 295:813–818 3. Sehgal PB (2008) Paradigm shifts in the cell biology of STAT signaling. Semin Cell Dev Biol 19:329–340 4. Wen Z, Zhong Z, Darnell JE Jr (1995) Maximal activation of transcription by Stat1 and Stat3 requires both tyrosine and serine phosphorylation. Cell 82:241–250 5. Marg A, Shan Y, Meyer T, Meissner T, Brandenburg M, Vinkemeier U (2004) Nucleocytoplasmic shuttling by nucleoporins Nup153 and Nup214 and CRM1-dependent nuclear export control the subcellular distribution of latent Stat1. J Cell Biol 165:823–833 6. Mertens C, Zhong M, Krishnaraj R, Zou W, Chen X, Darnell JE Jr (2006) Dephosphorylation of phosphotyrosine on STAT1 dimers requires extensive spatial reorientation of the monomers facilitated by the N-terminal domain. Genes Dev 20:3372–3381 7. Zhong M, Henriksen MA, Takeuchi K, Schaefer O, Liu B, ten Hoeve J, Ren Z, Mao X, Chen X, Shuai K, Darnell JE Jr (2005) Implications of an antiparallel dimeric structure of nonphosphorylated STAT1 for the activation-inactivation cycle. Proc Natl Acad Sci USA 102:3966–3971 8. Decker T, Kovarik P, Meinke A (1997) GAS elements: a few nucleotides with a major impact on cytokine-induced gene expression. J Interferon Cytokine Res 17:121–134 9. Ehret GB, Reichenbach P, Schindler U, Horvath CM, Fritz S, Nabholz M, Bucher P (2001) DNA binding specificity of different STAT proteins. Comparison of in vitro specificity with natural target sites. J Biol Chem 276:6675–6688
10. Fagerlund R, Melen K, Kinnunen L, Julkunen I (2002) Arginine/lysine-rich nuclear localization signals mediate interactions between dimeric STATs and importin alpha 5. J Biol Chem 277:30072–30078 11. McBride KM, Banninger G, McDonald C, Reich NC (2002) Regulated nuclear import of the STAT1 transcription factor by direct binding of importin-alpha. EMBO J 21:1754–1763 12. Haspel RL, Darnell JE Jr (1999) A nuclear protein tyrosine phosphatase is required for the inactivation of Stat1. Proc Natl Acad Sci USA 96:10188–10193 13. Haspel RL, Salditt-Georgieff M, Darnell JE Jr (1996) The rapid inactivation of nuclear tyrosine phosphorylated Stat1 depends upon a protein tyrosine phosphatase. EMBO J 15:6262–6268 14. Ibarra-Sanchez MJ, Simoncic PD, Nestel FR, Duplay P, Lapp WS, Tremblay ML (2000) The T-cell protein tyrosine phosphatase. Semin Immunol 12:379–386 15. ten Hoeve J, de Jesus Ibarra-Sanchez M, Fu Y, Zhu W, Tremblay M, David M, Shuai K (2002) Identification of a nuclear Stat1 protein tyrosine phosphatase. Mol Cell Biol 22:5662–5668 16. Meyer T, Hendry L, Begitt A, John S, Vinkemeier U (2004) A single residue modulates tyrosine dephosphorylation, oligomerization, and nuclear accumulation of Stat transcription factors. J Biol Chem 279:18998–19007 17. Meyer T, Marg A, Lemke P, Wiesner B, Vinkemeier U (2003) DNA binding controls inactivation and nuclear accumulation of the transcription factor Stat1. Genes Dev 17:1992–2005 18. Chen X, Vinkemeier U, Zhao Y, Jeruzalmi D, Darnell JE Jr, Kuriyan J (1998) Crystal structure of a tyrosine phosphorylated STAT-1 dimer bound to DNA. Cell 93:827–839
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19. Ihle JN (2001) The Stat family in cytokine signaling. Curr Opin Cell Biol 3:211–217 20. Levy DE, Darnell JE Jr (2002) Stats: transcriptional control and biological impact. Nat Rev Mol Cell Biol 3:651–662 21. Shuai K, Horvath CM, Huang LH, Qureshi SA, Cowburn D, Darnell JE Jr (1994) Interferon activation of the transcription factor Stat91 involves dimerization through SH2-phosphotyrosyl peptide interactions. Cell 76:821–828 22. Vinkemeier U, Cohen SL, Moarefi I, Chait BT, Kuriyan J, Darnell JE Jr (1996) DNA binding of in vitro activated Stat1 alpha, Stat1 beta and truncated Stat1: interaction between NH2-terminal domains stabilizes binding of two dimers to tandem DNA sites. EMBO J 15:5616–5626 23. Xu X, Sun YL, Hoey T (1996) Cooperative DNA binding and sequence-selective recognition conferred by the STAT amino-terminal domain. Science 273:794–797 24. Horvath CM, Stark GR, Kerr IM, Darnell JE Jr (1996) Interactions between STAT and non-STAT proteins in the interferon-stimulated gene factor 3 transcription complex. Mol Cell Biol 16:6957–6964 25. Horvath CM, Wen Z, Darnell JE Jr (1995) A STAT protein domain that determines DNA sequence recognition suggests a novel DNAbinding domain. Genes Dev 9:984–994 26. Marg A, Meyer T, Vigneron M, Vinkemeier U (2008) Microinjected antibodies interfere with protein nucleocytoplasmic shuttling by distinct molecular mechanisms. Cytometry 73A:1128–1144 27. Yang E, Henriksen MA, Schaefer O, Zakharova N, Darnell JE Jr (2002) Dissociation time from DNA determines transcriptional function in a STAT1 linker mutant. J Biol Chem 277:13455–13462 28. Shuai K, Stark GR, Kerr IM, Darnell JE Jr (1993) A single phosphotyrosine residue of Stat91 required for gene activation by interferon-gamma. Science 261:1744–1746 29. Hendry L, John S (2004) Regulation of STAT signaling by proteolytic processing. Eur J Biochem 271:4613–4620 30. Darnell JE Jr, Kerr IM, Stark GR (1994) JakSTAT pathways and transcriptional activation in response to IFNs and other extracellular signaling proteins. Science 264:1415–1421 31. Greenlund AC, Morales MO, Viviano BL, Yan H, Krolewski J, Schreiber RD (1995) Stat recruitment by tyrosine-phosphorylated cytokine receptors: an ordered reversible affinity-driven process. Immunity 2:677–687
32. Durbin JE, Hackenmiller R, Simon MC, Levy DE (1996) Targeted disruption of the mouse Stat1 gene results in compromised innate immunity to viral disease. Cell 84:443–450 33. Meraz MA, White JM, Sheehan KC, Bach EA, Rodig SJ, Dighe AS, Kaplan DH, Riley JK, Greenlund AC, Campbell D, CarverMoore K, DuBois RN, Clark R, Aguet M, Schreiber RD (1996) Targeted disruption of the Stat1 gene in mice reveals unexpected physiologic specificity in the JAK-STAT signaling pathway. Cell 84:431–442 34. Meyer T, Begitt A, Vinkemeier U (2007) Green fluorescent protein-tagging reduces the nucleocytoplasmic shuttling specifically of unphosphorylated STAT1. FEBS J 274:815–826 35. Müller M, Laxton C, Briscoe J, Schindler C, Improta T, Darnell JE Jr, Stark GR, Kerr IM (1993) Complementation of a mutant cell line: central role of the 91 kDa polypeptide of ISGF3 in the interferon-alpha and -gamma signal transduction pathways. EMBO J 12:4221–4228 36. Begitt A, Meyer T, van Rossum M, Vinkemeier U (2000) Nucleocytoplasmic translocation of Stat1 is regulated by a leucine-rich export signal in the coiled-coil domain. Proc Natl Acad Sci USA 97:10418–10423 37. Shuai K, Schindler C, Prezioso VR, Darnell JE Jr (1992) Activation of transcription by IFN-gamma: tyrosine phosphorylation of a 91-kD DNA binding protein. Science 258:1808–1812 38. Gordon JA (1991) Use of vanadate as proteinphosphotyrosine phosphatase inhibitor. Meth ods Enzymol 201:477–482 39. Kudo N, Wolff B, Sekimoto T, Schreiner EP, Yoneda Y, Yanagida M, Horinouchi S, Yoshida M (1998) Leptomycin B inhibition of signalmediated nuclear export by direct binding to CRM1. Exp Cell Res 242:540–547 40. Meissner T, Krause E, Vinkemeier U (2004) Ratjadone and leptomycin B block CRM1dependent nuclear export by identical mechanisms. FEBS Lett 576:27–30 41. Köster M, Hauser H (1999) Dynamic redistribution of STAT1 protein in IFN signaling visualized by GFP fusion proteins. Eur J Biochem 260:137–144 42. Lillemeier BF, Köster M, Kerr IM (2001) STAT1 from the cell membrane to the DNA. EMBO J 20:2508–2517 43. Köster M, Frahm T, Hauser H (2005) Nucleocytoplasmic shuttling revealed by FRAP and FLIP technologies. Curr Opin Biotechnol 16:28–34
Assessing Sequence-Specific DNA Binding and Transcriptional Activity 44. Lödige I, Marg A, Wiesner B, Malecová B, Oelgeschläger T, Vinkemeier U (2005) Nuclear export determines the cytokine sensitivity of STAT transcription factors. J Biol Chem 280:43087–43099 45. Meyer T, Begitt A, Lödige I, van Rossum M, Vinkemeier U (2002) Constitutive and IFNg-induced nuclear import of STAT1 proceed through independent pathways. EMBO J 21:344–354 46. Bromberg J, Chen X (2001) STAT proteins: signal transducers and activators of transcription. Methods Enzymol 333:138–151 47. Adam SA, Marr RS, Gerace L (1990) Nuclear protein import in permeabilized mammalian
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cells requires soluble cytoplasmic factors. J Cell Biol 111:807–816 48. Finlay DR, Newmeyer DD, Price TM, Forbes DJ (1987) Inhibition of in vitro nuclear transport by a lectin that binds to nuclear pores. J Cell Biol 104:189–200 49. Axelrod D, Koppel DE, Schlessinger J, Elson E, Webb WW (1976) Mobility measurements by analysis of fluorescence photobleaching recovery kinetics. Biophy J 16:1055–1069 50. Herrmann A, Vogt M, Mönnigmann M, Clahsen T, Sommer U, Haan S, Poli V, Heinrich PC, Müller-Newen G (2007) Nucleocytoplasmic shuttling of persistently activated STAT3. J Cell Sci 120:3249–3261
Chapter 9 Analysis of Nuclear Export Using Photoactivatable GFP Fusion Proteins and Interspecies Heterokaryons Kerry-Ann Nakrieko, Iordanka A. Ivanova, and Lina Dagnino Abstract In this chapter, we review protocols for the analysis of nucleocytoplasmic shuttling of transcription factors and nuclear proteins, using two different approaches. The first involves the use of photoactivatable forms of the protein of interest by fusion to photoactivatable green fluorescent protein to follow its movement out of the nucleus by live-cell confocal microscopy. This methodology allows for the kinetic characterization of protein movements as well as measurement of steady-state levels. In a second procedure to assess the ability of a nuclear protein to move into and out of the nucleus, we describe the use of interspecies heterokaryon assays, which provide a measurement of steady-state distribution. These technologies are directly applicable to the analysis of nucleocytoplasmic movements not only of transcription factors, but also other nuclear proteins. Key words: Nuclear export, Nucleocytoplasmic shuttling, Photoactivatable green fluorescent protein, Heterokaryon
1. Introduction The subcellular distribution of many proteins is key to the regulation of their function. An important challenge in this field of study has been the development of approaches that provide insight into the dynamic behavior of proteins in intact cells. A major advance in the characterization of protein localization patterns and movements in cells has been achieved with the introduction of photoactivatable green fluorescent protein (PA-GFP) (1). Photoactivatable fluorescent proteins are easily observed over darker backgrounds and, thus, they allow analysis of specific protein pools, organelles, or cells. Further, the dynamics of these proteins can be monitored spatiotemporally in the population of
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interest. In this chapter, protocols for activation and analysis of PA-GFP fusion proteins in living cells by confocal microscopy are described. The methods we describe were developed for primary cultures of epithelial cells (2); however, details regarding transfection and subsequent analyses are applicable to other cell types. We also describe complementary assays using interspecies heterokaryons, which are adaptable to visualization of nucleocytoplasmic shuttling of transcription factors and nuclear proteins.
2. Materials 2.1. Keratinocyte Culture
1. Bacterial grade Petri dishes. 2. Dissecting scissors (Fine Science Tools), scalpel, and two forceps (Fine Science Tools). 3. Sterile gauze (5 × 5 cm pads). 4. 50-ml conical tubes. 5. 70-mm nylon cell strainer (BD Falcon, BD Biosciences). 6. 0.25% (w/v) Trypsin, prepared by dilution of GIBCO 2.5% Trypsin (Invitrogen) in sterile Ca2-free phosphate-buffered saline (PBS). 7. Chelex-treated fetal bovine serum (FBS): Stir the FBS with Chelex 100 chelating resin (200–400 mesh, sodium salt, BioRad) to remove Ca2+ ions (40 g resin/500 ml serum) at room temperature for 1 h. Filter sequentially through filter paper and 0.45-mm sterile filters. Store at −20°C in 40-ml aliquots. 8. Keratinocyte culture medium, prepared with Eagle’s Minimum essential medium without CaCl2 (EMEM without CaCl2, BioWhittaker Lonza), supplemented with 8% (v/v) FBS pretreated with chelex resin, 100 units/ml penicillin and 0.1 mg/ml streptomycin (GIBCO; Invitrogen), hydrocortisone (74 ng/ml, Sigma-ALDRICH), insulin (5 mg/ml, Sigma-ALDRICH), cholera toxin (9.5 ng/ml, List Biological Laboratories), triiodothyronine (6.7 ng/ml, SigmaALDRICH), and epidermal growth factor (5 ng/ml, SigmaALDRICH). 9. Sterile glass-bottom culture dishes (MatTek, see Note 1). 10. Collagen Type I matrix, prepared by dissolving Collagen Type I (BD Biosciences) in sterile 0.02 N acetic acid, to a final concentration of 50 mg/ml.
2.2. PEI Transfection Mix
1. PEI transfection reagent stock solution (1 mg/ml), prepared by dissolving 20 mg polyethyleneimine (PEI), 25 kDa linear (Polysciences) in 18 ml distilled water, adjusting the solution
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to pH 7.0 with HCl (see Note 2). Add distilled water to a final volume of 20 ml, filter sterilize through 0.22-mm membrane. Store in sterile microfuge tubes in 500-ml aliquots at −20°C (see Note 3). 2. Sterile 150 mM NaCl solution. 3. Sterile 1.5-ml microfuge tubes. 2.3. Microscopy Equipment
2.4. Expression Vectors and Pharmacological Inhibitors
We conducted activation of PA-GFP and micrograph acquisition on a Zeiss LSM 510 confocal microscope equipped with a META system, Zen 2007 software, and live imaging capabilities (Zeiss). This system included a PlanApochromat 63×/1/4 Oil Ph3 objective lens, a mercury lamp, green and red fluorescence filter cubes, 488 nm Argon 2 laser, 543 nm HeNe1 laser, 720–930 nm multiphoton chameleon laser (see Note 4), a Tempcontrol 37-2 digital heated stage set to 37°C, a Tempcontrol miniobjective warmer, and a CTI-Controller 3700 digital heated air flow-through chamber with humidifier set to 5% CO2 with both ventilation speed and heat intensity set to 2.0. The equipment has Zeiss LSM Image Browser software for image processing. 1. Photoactivatable fusion proteins were generated using a mammalian expression vector encoding PA-GFP (1) (gift from Dr. J. Lippincott-Schwartz). 2. Mammalian expression vector encoding a monomeric red fluorescent protein (RFP) containing the SV40 nuclear localization signal (SV40NLS-RFP (2), see Note 5), and for V5-tagged E2F1 transcription factors (3, 4). 3. Leptomycin B (Sigma-ALDRICH). Prepare stock of 5 mg/ ml stock in dimethylsulfoxide and store in aliquots at −20°C. 4. Cycloheximide (Sigma-ALDRICH). Prepare stock of 10 mg/ ml in ethanol and store in aliquots at −20°C.
2.5. Microscopy Experiments and Heterokaryon Formation Assays
1. Triton X-100 (EMD). 2. Fraction V bovine serum albumin (BSA; Calbiochem). 3. GIBCO Dulbecco’s modified minimum essential medium (DMEM) without phenol red (Invitrogen, see Note 6). 4. Solution of 0.25% (w/v) trypsin and 1 mM ethylenediaminetetraacetic acid (EDTA; Invitrogen). 5. 20% (w/v) paraformaldehyde (PFA): Add 20 g PFA (Fisher Scientific) to 80 ml PBS. With constant stirring, adjust the pH to 7.0 by adding 10 N NaOH dropwise. Once the PFA has dissolved, adjust the final volume of the solution to 100 ml. Store at −20°C. 6. 4% (w/v) PFA, prepared by diluting the 20% (w/v) PFA solution with PBS (see Note 7).
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7. Glycine quench solution: 100 mM glycine (Bioshop) dissolved in PBS. 8. Permeabilizing solution: 0.1% (v/v) Triton X-100 and 0.1% (w/v) BSA dissolved in PBS. 9. Blocking solution: PBS containing 1% (w/v) nonfat dry milk. 10. Hoescht 33258 solution. Dissolve 12 mg Hoescht 33258 (Sigma-ALDRICH) in 10 ml water. Store at 4°C protected from the light. 11. 50% (w/v) Polyethylene glycol solution for cell fusion (HybriMax, Sigma-ALDRICH).
3. Methods 3.1. Preparation of Collagen-Coated Culture Substrates
3.2. Preparation of Primary Murine Keratinocyte Cultures
Coating of the glass-bottom culture dishes must be conducted 24 h prior to seeding the cells. To coat, add in a laminar flow cell culture hood 300 ml of sterile Collagen Type I matrix (50 g/ml) to the coverglass portion of each 35-mm glass-bottom culture dish to be used. Cover the dish, and incubate overnight at room temperature. Remove the collagen matrix by aspiration, wash thrice with sterile PBS. Add enough sterile PBS to cover the coverglass, and remove by aspiration immediately prior to use. 1. Newborn mice (0–3 days old) are sacrificed by CO2 inhalation, and immersed in 70% ethanol. 2. Dispense 6 ml of freshly thawed 0.25% trypsin into a 10-cm bacterial grade Petri dish. 3. In a laminar flow cell culture hood, remove the trunk skin using scissors, forceps, and a scalpel. 4. Place the skin dermis side down in the Petri dishes with 0.25% trypsin, and incubate overnight at 4°C. 5. Replace the trypsin with fresh 0.25% trypsin, and incubate at 37°C for 0.5 h (see Note 8). 6. Place the trypsinized skin on a clean Petri dish with the epidermis side down, and gently separate the epidermis from the dermis with a pair of sterile forceps. 7. Place the epidermis in a bacterial culture dish containing keratinocyte growth medium, mince with sterile scissors, and transfer to a conical tube. 8. Incubate at 37°C with gentle rocking for 20 min, and filter through a 70-mm nylon cell strainer into a clean, sterile conical tube.
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9. Seed the keratinocytes on the collagen-coated 35-mm glass-bottom culture dishes at a density of 2.5 × 105 cells/cm2 and culture overnight. 10. The day after plating, replace the medium with fresh keratinocyte growth medium. Replace the culture medium every other day until transfection, to a maximum of 4 days from the time of initial plating. 3.3. Transient Transfection and Inhibitor Treatments
1. Prior to transfection, ensure that the cells have reached 70–80% confluence. Replace culture medium with 1.9 ml fresh medium. 2. Thaw PEI transfection reagent on ice. 3. Mix 2 mg of vector DNA encoding SV40NLS-RFP (1 mg/ml stock) and 2 mg of vector DNA encoding the PA-GFP-labeled protein of interest with 188 ml of 150 mM NaCl in a sterile microfuge tube (see Note 9). 4. Add 8 ml of PEI transfection reagent (1 mg/ml), and vortex (see Note 10). 5. Incubate for 10 min at room temperature. 6. Add dropwise the DNA/PEI mix to the cultures and incubate the cells for 4–16 h at 37°C. 7. Replace the DNA/PEI-containing medium with warm fresh growth medium and culture the cells at 37°C for 24 h. 8. To inhibit CRM1-mediated nuclear export, add leptomycin B to the culture medium to a final concentration of 5 ng/ml. Incubate 2–4 h. To inhibit de novo protein synthesis (see Note 11), add cycloheximide to a final concentration of 100 mg/ml, and incubate 2–4 h. Incubation of cell cultures under these conditions with both leptomycin B and cycloheximide is possible, as it does not affect cell viability. 9. Proceed with optimization of PA-GFP fusion protein photoactivation and live-cell confocal microscopy.
3.4. Optimization of PA-GFP Fusion Protein Photoactivation
1. Place culture dish with transfected cells on ice. Aspirate the culture medium. 2. Rinse the cultures with ice-cold PBS. 3. Fix the cells by incubation with freshly diluted 4% PFA in icecold PBS for 15–30 min. 4. Rinse the cells thrice with ice-cold PBS and maintain them in PBS for subsequent imaging. 5. Turn on the confocal imaging system, as well as the chameleon 720–939 nm, the 488 nm, and the 543 nm lasers. 6. Transfer the glass-bottom culture dish with the transfected cells to the microscope stage.
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7. Identify and focus on the nucleus of a transfected cell by mRFP fluorescence. 8. Observe the selected cell using green fluorescence. If this cell also expresses PA-GFP, it will generally exhibit a very faint green fluorescence. If it does not, it will appear completely dark. In that case, select another RFP-positive cell. 9. Focus on the nucleus by scanning with the 543 nm laser, to identify the Z-position with the strongest signal. 10. Under the Regions function, use the image acquired in the 543 nm scan to select a region of interest (ROI) within the nucleus as target for photoactivation. Select a circular ROI with 3–5 mm diameter. 11. Systematically determine the optimal excitation wavelength for the PA-GFP fusion protein analyzed, by increasingly tuning the chameleon laser in 10-nm increments, beginning at 720 nm. For each increment, use the Bleach function to select activation for 25 iterations (5–10 s) at a laser capacity of 95%. Use a larger or smaller number of iterations (15–250) to optimize photoactivation and PA-GFP detection capabilities, with the aim of producing the strongest PA-GFP signal possible (see Note 12). 3.5. Photoactivation and Confocal Microscopy
1. Remove growth medium, rinse with PBS, and add warm DMEM without serum or phenol red. 2. Turn on the confocal imaging system, as well as the chameleon 720–930 nm, the 488 nm, and the 543 nm lasers. Tune the chameleon laser to 750 nm, or to the wavelength determined during the optimization steps of Subheading 3.4. Set the 488 nm laser to 40–50% output capacity. 3. Set the objective warmer to 37°C and place around the 63× lens. 4. Ensure that the heated stage and heated air flow-through chamber with humidifier are equilibrated, respectively, to 37°C and 5% CO2. 5. Place the culture dish with the transfected cells onto the heated stage inside the heated air chamber, and allow equilibration for 5–10 min. 6. Identify, select, and focus on a transfected cell as described in steps 7–9, Subheading 3.4. 7. Under the Regions menu, use the image acquired in the 543 nm scan to select a circular ROI (3–5 mm diameter) within the nucleus targeted for photoactivation. 8. Acquire an image of the selected cell prior to activation, by selecting one scan at 488 nm excitation.
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9. Using the Bleach function, select the nuclear ROI (3–5 mm diameter) to photobleach. Set the output capacity of the chameleon laser to 95%. Activate the PA-GFP moiety using 50–250 iterations, with a bleaching time range of 15–45 s (see Note 13). 10. Acquire time-lapse images of the cell following activation, using the same settings used for the image acquired prior to activation. 3.6. Interspecies Heterokaryon Assays
1. Prepare cultures of NIH 3T3 mouse fibroblasts in 75-cm2 culture flasks, using DMEM supplemented with 8% (v/v) FBS as growth medium. 2. Seed HeLa cells suspended in DMEM supplemented with 8% (v/v) FBS at a density of 20,000 cells/cm2 on collagen-coated glass-bottom culture dishes and allow them to attach for 4 h. 3. Transfect the HeLa cells with the vector of interest, as described in Subheading 3.3, culturing the cells in the presence of the DNA/transfection mix for 24 h. 4. At this time, prepare NIH 3T3 and HeLa cells for heterokaryon formation. 5. Remove the culture medium of the NIH 3T3 cells and rinse twice with Ca2-free PBS. 6. Detach the NIH 3T3 cells from the culture flask with 3 ml trypsin/EDTA mix. Once they have detached, transfer to a conical centrifuge tube containing 3 ml growth medium and centrifuge at 500 × g for 5 min. 7. Resuspend the NIH 3T3 cells in growth medium and determine the cell density with a hemocytometer or a cell counter. 8. Rinse the HeLa cell cultures four times with PBS to thoroughly remove the DNA/transfection mix. 9. Add NIH 3T3 cells at a density of 30,000 cells/cm2. 10. Add growth medium to obtain a final volume of 2 ml. Gently swirl the culture dish to ensure even distribution of the NIH 3T3 cells. 11. Incubate the cocultures at 37°C for 1 h to allow the mouse cells to attach. 12. Add cycloheximide to the cocultures, to a final concentration of 100 mg/ml, and continue culture for 2 h. If CRM1mediated inhibition of nuclear export is desired, add leptomycin B at a final concentration of 5 ng/ml. 13. Rinse the cells twice with warm PBS. Fuse the two cell types by incubation in 1 ml of 50% Hybri-Max polyethylene glycol for exactly 3 min at 37°C.
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14. Remove the polyethylene glycol and rinse thrice with warm PBS. 15. Add growth medium supplemented with 100 mg/ml cycloheximide, and culture at 37°C for 2 h. 16. Process the cultures for indirect immunofluorescence microscopy. 3.7. Indirect Immunofluorescence Microscopy Analysis of Heterokaryons
1. Place culture dish with fused cells on ice. Aspirate the culture medium. 2. Rinse the cultures with ice-cold PBS. 3. Fix the cells by incubation with freshly diluted 4% PFA in ice-cold PBS for 15–30 min. 4. Rinse the cells thrice with ice-cold PBS. 5. Incubate with 100 mM glycine at 22°C for 15 min to quench the PFA. 6. Permeabilize the cells by incubation in Triton X-100 permeabilizing solution, 10 min at 22°C, with gentle rocking. Remove the permeabilizing solution. 7. Rinse the cells thrice with PBS. Remove the PBS. 8. To block nonspecific antibody binding, add Blocking solution containing PBS with 5% (v/v) serum obtained from the same species in which the secondary antibody was raised. Rock 1 h at 22°C. Remove the blocking solution. 9. Repeat step 7. 10. Incubate in a solution of primary antibody diluted in PBS supplemented with 5% (v/v) serum obtained from the same species in which the secondary antibody was raised. Rock the samples for 1 h at 22°C, or 16 h at 4°C. 11. Remove the primary antibody solution. Wash with four changes of PBS, rocking for 10–15 min each. 12. Remove the PBS and add a solution containing an Alexa fluorochrome-labeled secondary antibody diluted in PBS containing 5% serum from the same species in which this secondary antibody was raised. Incubate with rocking for 1 h at 22°C protected from the light. 13. Remove secondary antibody solution. Wash as in step 11, keeping the samples protected from the light. 14. Add Hoescht 33258 (0.1–0.3 mg/ml, final) in PBS to stain the DNA. Rock samples for 5–15 min at 22°C, protected from the light. 15. Remove the Hoescht-containing PBS. Wash with PBS as in step 13. 16. Mount with mounting medium. Allow samples to dry for 16 h at 22°C in the dark. Store mounted specimens at −20°C.
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4. Notes 1. The choice of coverglass in the culture dish will depend on the type of microscope objectives to be used. Many lenses have optimal performance with No. 1.5 coverslips. 2. The pH of the solution is critical to ensure optimal transfection efficiency. 3. The PEI stock solution will store well at −20°C for about 6 months. 4. An ultraviolet 405 or 413 nm laser line may be used instead of the multiphoton chameleon laser if the latter is not available. 5. Cotransfection with an mRFP marker facilitates visualization of the transfected cells. 6. The use of medium with phenol red or serum creates background fluorescence, generating light that is collected in both green and red spectra, and interferes with the detection of PA-GFP. 7. 4% PFA is prepared by diluting the appropriate amount of 20% PFA with PBS immediately prior to use. This is a single-use solution. 8. Check periodically to verify extent of tissue digestion. When the epidermis appears as a thin translucent sheet overlaying the thick, pink dermis, it is ready for isolation. Do not continue the incubation with trypsin longer than necessary, as overdigestion of the tissue substantially reduces keratinocyte viability. 9. Due to the very low background fluorescence of the PA-GFP moiety, we cotransfect the cells with a monomeric RFP protein, which also contains a nuclear localization signal, such as SV40NLS-RFP. This allows visualization of transfected cells by epifluorescence microscopy. Depending on the cell type transfected, the proportion of cells expressing both RFP and PA-GFP will be 30–70%. 10. It is critical to add the PEI to the diluted DNA mix. Do not mix concentrated DNA with either dilute or concentrated PEI. 11. A major advantage of photoactivation is that, unlike other approaches, the fluorescence of newly synthesized proteins will not contribute to the signal. Therefore, inhibition of protein synthesis is not required if the experiment exclusively aims at detection of the photoactivated protein. However, note that inhibition of protein synthesis is necessary for interspecies heterokaryon assays. 12. Tuning the chameleon laser to 826 nm would approximate the optimal excitation wavelength reported for PA-GFP (413 nm). However, PA-GFP can be activated by multiphoton lasers
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tuned in the 720–840 nm range (5). The activation of some of the PA-GFP fusion proteins we have analyzed was optimized with a multiphoton laser line tuned to 750 nm. 13. Select the smallest ROI that gives efficient activation to minimize photodamage to the cell. Save the selected ROI for use on additional samples or in future experiments.
Acknowledgments This work was supported by grants from the Canadian Institutes of Health Research, the Natural Sciences and Engineering Research Council, and the National Cancer Institute of Canada. References 1. Patterson GH, Lippincott-Schwartz J (2002) A photoactivatable GFP for selective photolabeling of proteins and cells. Science 297:1873–1877 2. Nakrieko KA, Vespa A, Mason D, Irvine T, D’Souza SJA, Dagnino L (2008) Modulation of integrin-linked kinase cytoplasmic shuttling by ILKAP and CRM1. Cell Cycle 7:2157–2166 3. Ivanova IA, Vespa A, Dagnino L (2007) A novel mechanism of E2F1 regulation via
nucleocytoplasmic shuttling: determinants of nuclear import and export. Cell Cycle 6:2186–2195 4. Ivanova IA, Nakrieko KA, Dagnino L (2009) Phosphorylation by p38 MAP kinase is required for E2F1 degradation and keratinocyte differentiation. Oncogene 28(1):52–62 5. Schneider M, Barozzi S, Testa I, Faretta M, Diaspro A (2005) Two-photon activation and excitation properties of PA-GFP in the 720920-nm region. Biophys J 89:1346–1352
Chapter 10 Determination of Nuclear Localization Signal Sequences for Krüppel-Like Factor 8 Tina S. Mehta*, Farah Monzur*, and Jihe Zhao Abstract Transcription factor proteins function in the nucleus to regulate gene expression. Many transcription factors are critical regulators of tumor progression. Conversely, many oncogenic and tumor suppressor proteins are transcription factors or other types of nuclear proteins. Because of their critical physiological and pathological roles, these tumor regulators are tightly regulated not only in the protein expression but also in their subcellular localization. This chapter is focused on experimental strategies and method details for the identification and characterization of nuclear localization signal sequences for nuclear proteins using the Krüppel-like transcription factor 8 as an example. Key words: Krüppel-like factor, KLF8, Nuclear localization, Site-directed mutagenesis, Overlapping PCR, Transfection, Western blotting, Fluorescent microscopy
1. Introduction KLF8 belongs to the Krüppel-like C2H2 zinc-finger transcription factor family proteins. Several KLF family members, including KLF4, KLF5, and KLF6, have been identified as either oncogenes or tumor suppressors (1–3). KLF8 is a relatively new member of this family and is emerging as a critical regulator of cancer progression. KLF8 was initially identified as a transcriptional repressor (4). Later we found KLF8 to be a dual transcriptional factor that can both repress and activate transcription of target genes including cyclin D1, KLF4, and E-cadherin (5–7). We and others identified KLF8 as a downstream effector of FAK (7) and a crucial regulator of oncogenic transformation, epithelial to mesenchymal transition and tumor cell invasion (7–11). Nuclear oncogenic or tumor suppressor proteins are tightly regulated in
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nuclear localization in addition to protein expression. Nuclear localization signal (NLS) sequences have been identified for a few KLFs (12–14). Importantly, we have now identified the NLS sequences for KLF8. While the results of this study are submitted to elsewhere, we describe here in details about the experimental design strategies and methods which hopefully apply to nuclear localization studies for other nuclear proteins.
2. Materials 2.1. PCR and DNA Agarose Gel Electrophoresis (DAGE)
1. Deep Vent DNA polymerase (New England Biolabs). 2. T4 DNA ligase (New England Biolabs). 3. dNTP (New England Biolabs). 4. Ethidium bromide (Bio-Rad). 5. Certified Molecular Biology Agarose (Bio-Rad). 6. Aurum Plasmid Mini Kit (Bio-Rad). 7. Powerpac Basic Power Supply (Bio-Rad). 8. Powerpac HC Power Supply (Bio-Rad). 9. Qiaquick Gel Extraction Kit (Qiagen). 10. QIAprep Spin Mini-Prep Kit (Qiagen). 11. iCycler Thermal Cycler (Bio-Rad). 12. Mini Sub-Cell GT System (Bio-Rad). 13. Kodak GL440 Imaging Documentation System (Kodak). 14. 1 kb Plus DNA Ladder (Invitrogen). 15. Ampicillin (Fisher). 16. Microcentrifuge (Eppendorf).
2.2. Cell Culture and Transfection
1. Lipofectamine 2000 Transfection Reagent (Invitrogen). 2. DMEM (Invitrogen). 3. Falcon 12-well tissue culture plate (Fisher).
2.3. SDSPolyacrylamide Gel Electrophoresis (PAGE) and Western Blotting
1. 40% Acrylamide:Bisacrylamide Solution (Bio-Rad). 2. TEMED (Bio-Rad). 3. Precision Plus Standard Dual Color protein molecular weight markers (Bio-Rad). 4. Mini-PROTEAN 3 CELL (Bio-Rad). 5. HRP-Donkey Anti-mouse IgG (H + L) (Jackson Immuno Research Lab). 6. HRP-Donkey Anti-rabbit IgG (H + L) (Jackson Immuno Research Lab).
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7. Normal goat serum (Jackson ImmunoResearch Lab). 8. Protran BA83 Nitrocellulose Membrane (Whatman). 9. Nestle Carnation Instant Nonfat Dry Milk (Nestle USA). 10. Microcentrifuge (Eppendorf). 11. Isotemp Heating Block (Fisher). 12. Pierce* Enhanced Chemiluminescent (ECL) Substrate (Fisher). 13. Automatic X-ray Film Processor (Hisapdebu JP33). 14. Fuji Film LAS-3000 Imager (Fuji Film Global). 2.4. Immunofluorescent Staining and Microscopy
1. FITC-Donkey Anti-Mouse IgG (H + L) (Jackson Immuno Research Lab). 2. FITC-Donkey Anti-rabbit IgG (H + L) (Jackson Immuno Research Lab). 3. TexasRed-Donkey Anti-Mouse IgG (H + L) (Jackson Immuno Research Lab). 4. TexasRed-Donkey Anti-Rabbit IgG (H + L) (Jackson Immuno Research Lab). 5. SlowFad Gold Antifade Reagent (Invitrogen). 6. Microscope Slides (Fisher). 7. Microscope Cover Glass (coverslip) (Fisher). 8. Microscope (Olympus BMX-60 microscope equipped with a cooled charge-coupled device (CCD) sensi-camera (Cooke, Auburn Hills, MI) and Slidebook software (Intelligent Imaging Innovations, Denver, CO)).
3. Methods 3.1. Overall Research Flow (Fig. 1)
As shown in Fig. 1, the general procedure involves the identification of suspected NLS sequences in KLF8 and determination of their requirement and sufficiency for the nuclear localization of KLF8 by disrupting the sequences and fusing the sequences to GFP, respectively, followed by transfection and fluorescent microscopy.
3.2. NLS Sequence Search (Fig. 2)
There are in general two strategies to search for suspected NLS sequences: computer programs and amino acid sequence homology search (see Note 1). There are a number of free internet computer programs that are widely used for this purpose. Essentially, these programs perform the automatic search for potential NLS sequences, particularly the classical type NLSs including monopartite NLS (mNLS, a single stretch of basic aa
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Fig. 1. Flowchart of the procedure. Shown are general strategic sequences of the experiments to be done.
mNLS? N
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Fig. 2. NLS prediction. The diagram of KLF8 protein structure is shown. The PSORT computer program predicts two classical monopartite nuclear localization signal (mNLS) sequences, the first one is located at the amino terminus of the first zinc finger and the second one is located within the carboxyl terminus of the third zinc finger. Both the first mNLS and the zinc fingers are conserved among the KLF family proteins and have been suggested to be the functional NLS sequences for KLF1 and KLF4. Neither the computer program nor the homology search has predicted any NLS sequences within the amino terminal regulatory region marked with the question mark.
with a consensus sequence of (K/R)4–6) and bipartite NLS (two small stretches of basic aa linked by 10–12 aa in between with a consensus sequence of (K/R)2x10–12(K/R)3) (15). We used the PSORT program (http://psort.ims.u-tokyo.ac.jp/form2.html) to predict potential NLSs in KLF8. With this program, all that needs to be done was to enter the amino acid sequence or access number of SWISS-PROT and then click the “Submit” button. The program would then immediately predict the probability of nuclear localization and the potential NLS sequences and positions. This program identified two mNLSs (269KRRR and 352HRRR or 353RRRH) in KLF8 flanking the zinc-finger region (see Fig. 2). Homology search predicted the zinc-finger region as suspected NLS sequence as well, given that previous studies demonstrated that the conserved zinc-finger regions (as well as the leftmost mNLS) are functional NLSs for both KLF1 (12, 13) and KLF4 (14).
Determination of Nuclear Localization Signal Sequences for Krüppel-Like Factor 8
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Fig. 3. Construction of NLS mutants by PCR. Shown are Schematic illustrations of the PCR strategies to be used. The double strand cDNA template and primers are shown in lines and the PCR products are illustrated as protein peptide diagrams shown as white and black boxes. Terminal truncation mutations are generated by a one-step PCR. Point mutations and internal deletion mutations involve two-step overlapping PCR.
3.3. PCR-Based Mutagenesis and DAGE to Disrupt the Predicted NLS Sequences (Fig. 3)
Disruption of the suspected NLS sequences can be achieved by site-directed mutagenesis using PCR to generate either point mutations (see Note 2) or deletion mutations. Because it is relatively easy to make terminal truncation mutants by a simple onestep PCR, no method details are discussed here. Overlapping PCR is one of the many convenient techniques used to generate point or internal deletion mutations. The main steps of the procedure are described below:
3.3.1. Mutant Primer Design
The pair of the mutant primers must match the cDNA template except for the nucleotides encoding the residues to be mutated or deleted. The nonmutated nucleotides in the primers should be at least 8-base loner at both ends. G/C content in the primers should be 50% or greater with a preferable 5-prime A/T and a 3-prime C/G placement. Master primers are usually 16–20-base long (see Note 3) and are immediately upstream (forward primer) or downstream (reverse primer) to the multiple cloning sites in the plasmid vector. Many companies can provide good quality service for primer synthesis and we usually use the services provided by Invitrogen.
3.3.2. Overlapping PCR
This procedure consists of two rounds of PCR. The first-round PCR uses plasmid cDNA as template and the forward master primer (Master-F) paired with the reverse mutant primer (Mutant-R) or the forward mutant primer (Mutant-F) paired
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with the reverse master primer (Master-R). The products are two fragments with the targeted site either mutated or deleted. The second-round PCR uses a mixture of these two fragments purified from gel (see below) as template (see Note 4) and the master primer pair. The PCR reaction in 50 ml (see Note 5) are prepared in the following order: 39.5 ml of nanopure water, 5 ml of 10× buffer, 1 ml of 10 mM dNTP, 1 ml of 50 mM MgCl2, 1 ml of 1 mg/ml primer-F, 1 ml of 1 mg/ml primer-R, 1 ml of 0.2 mg/ml template DNA, and 1 ml of DNA polymerase (see Note 6). The PCR program consists of 1 cycle of 94°C for 5 min, 30 cycles of 94°C for 30 s, 55–65°C (determined by the primers used) for 45 s and 72°C for 1 min per kb in length of product, and 1 cycle of 72°C for 15 min. 3.4. DAGE and DNA Purification
1. Prepare a small tray for agarose pouring. Make sure the tray is level. Insert comb.
3.4.1. Making and Running Agarose Gel
2. Pour 50 ml of the agarose gel TAE solution pre-warmed in the 55°C water bath into a 50 ml centrifuge tube. Add 5 ml ethidium bromide (10 mg/ml) into the solution. Invert a few times to mix. 3. Pour the gel solution slowly into the tray (see Note 7). Wait for about 30 min at room temperature. 4. Pour about 220 ml of 1× TAE running buffer (for a 500 ml stock solution, combine 93.05 g EDTA-Na2 and 400 ml water and adjust the pH to 8 with NaOH, then top up with water to a final volume of 500 ml) into the running chamber. 5. When the gel is solid, release by untwisting the gel from the big white chamber slowly, take comb out by slowly pulling up the comb up while pushing the glass tray down. 6. Place the glass tray into the running chamber. The side with the wells should be facing the negative (black) side. 7. Add 22 ml of the ethidium bromide stock to the running buffer on the positive (red) side of the running chamber. 8. Take 10 ml of 6× loading buffer and aliquot it into eight dots on a piece of parafilm (1 ml/dot). 9. Write down in notebook the positions of where each DNA sample is going to be loaded. 10. Load the gel (see Note 8) starting with the marker. For each DNA sample, mix 5 ml of DNA solution with the dye dot by slowly pipetting up and down a couple of times. 11. Put the cover with the electrodes red-to-red and black-toblack.
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12. Run at 120 V for 30 min (see Note 9). When done, carefully remove the gel with the tray. 13. Take a picture of the gel using the Kodak Image Documentation System. 3.4.2. Purification of DNA Fragments from DNA Agarose Gels
1. Place the gel on top of the long-beam UV light box with plastic shield in the chamber of the Kodak Imaging Documentation System. 2. Cut desired DNA bands from gel (see Note 10). 3. Transfer the gel slices into a 1.5-ml Eppendorf tube by using pipette tips. 4. Spin down the gel slice to the bottom of the tube for 15 s for easy estimation of the gel volume. Add 3 volumes of Buffer QG (see Note 11). 5. Put the Epp tube(s) in the 55°C water bath for 3–10 min until the gel is melted. 6. Add 1 volume of isopropanol to the solution. After pipetting up and down a few times to mix the solution, transfer 700 ml of the solution to a Qiaquick spin column. Spin at 13,000 rpm for 30 s (see Note 12). Discard flow-through. 7. Add 0.5 ml Buffer QG to each spin column. Spin for 30 s at 13,000 rpm. Discard flow-through. 8. Add 0.75 ml Buffer PE. Spin for 30 s at 13,000 rpm. Discard flow-through. 9. Spin at 13 krpm for 1 min without adding any solution. Discard flow-through. 10. Transfer the top part of the spin column into a 1.5 ml Eppendorf tube. 11. Add 35 ml sterile Buffer EB pre-warmed in the 55°C water bath. Let the solution sit at room temperature for 1 min. Spin down at 13,000 rpm for 1 min. Repeat once to collect a final volume of 70 ml of DNA solution. 12. Run an agarose gel with a little (1–5 ml) of the DNA to confirm the recovery. 13. After determining the DNA concentrations by spectrometry, keep the remaining at –20°C for future use.
3.4.3. Restriction Digestion
1. Prepare 20 ml of reaction mixture in an Eppendorf tube containing 0.5–2 mg of DNA, 2 ml of 10× Buffer, 0.2 ml of 100× BSA, 0.5 ml of enzyme, and nanopure water that adds up to 20 ml. 2. Mix well by stirring with the pipette tip. Centrifuge briefly at 13,000 rpm to spin down all the contents in the tube.
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3. Start reaction at the temperature and duration specific for the enzyme (see Note 13). 4. Add 4 ml of 6× loading dye to the reacted mixture to stop the reaction. Run an agarose gel to confirm the presence and size of the digested DNA. 3.4.4. Ligation
1. Prepare 20 ml of reaction mixture in a Eppendorf tube containing 1:1 molecular ratio of gel purified vector and insert DNA fragments (up to 2 mg in total), 5 ml of 5× Buffer and nanopure water. Use the vector mixture alone as a control. 2. Mix well by stirring with the pipette tip. Centrifuge briefly at 13,000 rpm to spin down all the contents in the tube. 3. Put the tube in 16°C water bath (in 4°C fridge or room) and let the reaction take place overnight.
3.4.5. Transformation
1. Set the temperature on the heating block (Isotemp) to 42°C (see Note 14). 2. Take new Eppendorfendorf tube and place on ice. 3. Thaw DH5a competent E. coli taken from the –80°C fridge on ice for a few minutes (see Note 15). 4. Pipette 100 ml DH5a into new Eppendorf tubes containing the 20 ml of ligation mixture pre-cooled on ice. Mix well by stirring with the pipette tip. Leave on ice for 20 min. 5. Heat the tubes in 42°C for 60 s. Immediately put on ice. 6. Add 900 ml of 4°C-stored sterile liquid broth (LB) (see Note 16). 7. Incubate in a 37°C shaker for 1 h. 8. In the meantime, prepare for streaking. Take out 2 clean LB + ampicillin (see Note 17) agar plates and let them dry under the fume hood for ~30 min. 9. Spin down the transformed E. coli at 5,000 rpm for 2 min. Take out ~900 ml of supernatant. Resuspend the bacteria in the remaining supernatant by tapping the tube. 10. Transfer the bacteria suspension with pipette and streak it evenly to the LB + Amp agar plate. 11. Incubate at 37°C overnight with the plates inverted. During the time the bacterial colonies containing the plasmid DNA should form on the agar surface.
3.4.6. Plasmid DNA Preparation
1. Pick up a single colony of transformed E. coli by scraping the single spot with an autoclaved culture tip. 2. Drop the colony along with the tip into a 15 ml bacteria culture tube containing 2 ml of LB with 50 mg/ml of ampicillin.
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3. Incubate in a 37°C shaker overnight. 4. Place 1.5 ml of the bacteria growth in a 1.5 ml Eppendorf tube. Centrifuge for 1 min at 13 krpm. Decant supernatant. 5. Add 250 ml of resuspension solution and vortex. 6. Add 250 ml of lysis solution and invert 5–6 times. 7. Add 350 ml of neutralization solution and invert 5–6 times. 8. Centrifuge for 5 min at 13 krpm. 9. Transfer supernatant to a spin column in another 1.5 ml tube with. 10. Centrifuge for 5 min at 13 krpm. Discard filtrate. 11. Add 750 ml of wash solution and flash-spin for 1 min. 12. Discard filtrate and spin again without adding any solution. 13. Transfer column to a clean 1.5 ml tube. 14. Elute, quantify, and keep the DNA for future use as described above in c.2. 15. Pick two clones for each mutant plasmid and send for sequencing to confirm the mutation by following the instructions from the service providers. We use Fisher’s sequencing service. 3.5. Transfection and Western Blotting to Confirm the Expression of the Mutant Proteins 3.5.1. Transfection
Correct DNA sequencing results do not always guarantee that the plasmids are capable of driving the protein expression. 1. Split the subconfluent 293 cells (see Note 18) by 1:4 or 1:5 into a 12-well dish and incubate overnight. 2. When the cells are 70–80% confluent, continue with the transfection. 3. Everything is to be done under the tissue culture hood under sterile condition. 4. Prepare transfection reaction mixture in the order of diluting 1 mg DNA in 50 ml of serum-free medium, diluting 3 ml of Lipofectamine 2000 in 50 ml of serum-free medium, and combine these two dilutions by gentle pipetting. Incubate at room temperature for 20 min. 5. Transfer the mixture to a tube containing 0.4 ml of serum-free medium, gently pipette and add to the cells pre-washed with serum-free medium. Incubate for 3–5 h. 6. Replace the medium with growth medium (see Note 19) and incubate overnight. 7. Carefully decant the medium and wash the cells with ice–cold PBS (see Note 20). 8. Add 100 ml of 1× SDS Sample Buffer to the cells. Swirl the plate on the rocker for 5–10 min.
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9. Transfer the lysates to 1.5 ml Eppendorf tubes and label these tubes. 10. Boil the tubes for 3–5 min in the Isotemp heat block. The lysates are now ready for western blotting or can be kept at −20°C for future use. 3.5.2. SDS-PAGE
1. Place 2 glass plates on napkins. Wipe the inner sides with 70% ethanol and Kimwipes. Wait a few seconds for the ethanol to evaporate. 2. Set up the gel cassette and make sure the glass plates are in level with the ground and there is no leakage. 3. Make a 1.5-mm 10% separating gel: mix 4 ml of nanopure water, 2 ml of 4× separating buffer (combine 91 g of Tris with 450 ml of nanopure water, adjust pH to 8.8 with 10–15 ml of concentrated HCl, add 10 ml of 20% SDS and nanopure water to 500 ml), 2 ml of 40% acrylamide:bisacrylamide solution, 40 ml of APS and 8 ml of TEMED. Pour the gel mix into the cassette. Gently add 0.5 ml of water on the top of the gel solution. Wait 20–30 min until the gel polymerizes (when a sharp line between the gel and water is visible). 4. While waiting, make the stacking gel: Mix 2.6 ml of nanopure water, 1 ml of 4× stacking buffer (combine 30.25 g of Tris with 450 ml of nanopure water, adjust pH to 6.8 with 10–15 ml of concentrated HCl, add 10 ml of 20% SDS and nanopure water to 500 ml), 0.4 ml of 40% acrylamide: bisacrylamide solution, 20 ml of APS and 4 ml of TEMED. Pour the gel mix into the cassette after removing the 0.5 ml water from it. Gently insert the comb into the gel solution (see Note 21). Wait for 20–30 min for the gel to polymerize. 5. Move the gel cassette into the running tank with the short glass plate facing the center. 6. Pour 800 ml of 1× SDS running buffer diluted from the 5× stock (combine 320 g of NaCl, 8 g of KCl, 57.6 g of Na2HPO4, 9.6 g of KH2PO4, and 330 ml of water, adjust pH to 7.4 with 5 N NaOH or HCl, and add water to 4 L) into the tank, starting from the middle of the apparatus. Let it spill into the outside compartment of the tank until it is under the white part of the apparatus that juts out. Carefully remove the comb (see Note 22) and bubbles from the wells on the top and from the bottom of the gel using a syringe. 7. Pipette up to 20 ml of boiled lysates into each well (see Note 23). For the molecular size marker, combine 5 ml of the marker and 15 ml of the 1× SDS sample buffer to make a final volume of 20 ml.
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8. Put the tank lid on and run for 10–20 min at 50 V and then for 1 h at 150 V. 9. Prepare the transfer sandwich while waiting for the running to complete: per gel, get 2 sponges, 2 filter papers, and 1 nitrocellulose membrane. Cut a small top-right corner off the membrane and write on it: initials, date, name of antibody (across the top), and the name of the samples on the gel (on the side). Put the sponges on the pan. Submerge them with 1× transfer buffer diluted from 5× stock (combine 58 g of glysine and 116 g of Trizma base, and add water to 4 L; to make 1× buffer, mix 1 volume of the 5× stock, 3 volumes of water, and 1 volume of methanol). Remove bubbles from the sponges by rubbing them out through the sides. Put one piece of 3MM filter paper on top of one of the sponges. Put the membrane on top of the filter. Put the second filter paper on top and then the second sponge. Let the sandwich soak in the transfer buffer while the gel is running. 10. Once the gel has completed running, take out the gel and cut the top right corner of the gel. Rinse the gel a few times in the transfer buffer. Place the gel facing down on one of the filter paper in the sandwich. Place the membrane on top of the gel with the cut corner aligned together. Assemble the sandwich in the black/white holder (see Note 24) and slide it into the transfer tank. 11. Put an ice block from the −20°C fridge in the transfer tank next to the black side. 12. Pour 1× transfer buffer into the tank until the tank juts out (~800 mL). Run at 100 V for 1 h (see Note 25). 3.5.3. Western Blotting
1. After the transfer is complete, use tweezers to remove the membrane and place it facing up in 20 ml of blocking buffer (5% carnation skim milk in PBS-T. To make PBS-T, mix 100 ml of 10× PBS, 900 ml of nanopure water and 1 ml of Tween-20). Rock slowly for 1 h. 2. Dump the blocking buffer into the sink. Rinse the membrane twice with 20 ml of PBS-T. 3. Apply 10 ml of the primary antibody solution in PBS-T (see Note 26). Rock at 4°C overnight (see Note 27). 4. Recover the antibody solution and keep at 4°C for future reuse. 5. Wash the membrane with 20 ml of PBS-T for five times (rinse twice, rock once for 10 min and twice for 5 min). 6. Apply 10 ml of the second antibody solution in PBS-T. Repeat incubating and washing as in steps 3 and 5 above.
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7. Treat the membrane with 3 ml of ECL Reagents for 1 min. Put saran wrap on top of the membrane. Use Kimwipes to smooth out the membrane and rub out bubbles. 8. Expose the membrane to an X-ray film and develop the film: Fold the top right corner of the X-ray film to match the cut corner of the membrane. Make four exposures of one membrane on a single film for 1 s, 15 s, 30 s, and 1 min. Develop the film. 9. Retrieve the X-ray film and record the results. 3.6. IF Staining and Microscopy to Determine the Requirement of the Predicted NLS Sequences for the Nuclear Localization (Fig. 4)
1. Using tweezers put a sterile coverslip into the bottom of each well of a 12-well plate. Rinse the coverslips with 1 ml of 1× PBS. 2. Re-plate NIH3T3 cells to the coverslips at a density of 1/4 or 1/5 confluence. Incubate overnight. 3. Transfect the cells as described above in 3.5.1.
3.6.1. Set up Cells for Transfection 3.6.2. Immunostaining for Fluorescent Microscopy
1. Rinse cells twice with ice–cold 1× PBS plus 100 mM CaCl2 and 100 mM MgCl2. 2. Add 1 ml of the fixative solution (i.e., 4% paraformaldehye in PBS: in fume hood, combine 36 g of paraformaldehyde, 600 ml of water, and 2 ml of 10 N NaOH and stir on 45°C hot plate until dissolved. Add 90 ml of 10× PBS and water to 900 ml. Store 50–100 ml aliquots at −20°C. Thaw at 37°C water bath until turbidity disappears before use) to each well. Incubate for 15 min at room temperature (see Note 28). 3. Rinse the cells twice with PBS. 4. Apply 30–60 ml of primary antibody solution (1:100–1:300 in 1× PBS). Incubate at 37°C for 30 min. Rinse as in step 3.
Fig. 4. Immunofluorescent microscopic examination of protein localization. Shown is immunofluorescent microscopic images for KLF8 (top row, stained with anti-Myc tag antibody) and the nuclei (middle row, stained with Hoechst 33258).
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5. Apply 30–60 ml of the FITC-conjugated secondary antibody and repeat steps 3 and 4. 6. Add 0.2 ml of 0.5 mg/ml Hoechst 33258 in PBS to the cells. Incubate at room temperature for 10 min. Rinse twice with 1× PBS and once with water. 7. Place a drop of mounting solution on a slide. Carefully transfer the coverslip upside down to the drop with curved edge tweezers (see Note 29). 8. Gently press down on the coverslip to get rid of any excess mounting solution. Use a pipette tip and the vacuum to remove excess mounting solution from the coverslip edges (see Note 30). 9. Glue the edge of the coverslip to the microscope slide with clear nail polish. 10. Take cell images under a fluorescent microscope (green image for the protein stained and blue images for the nuclei in the cells) (see Fig. 4). 3.6.3. GFP-based Fluorescent Microscopy to Determine the Sufficiency of the Predicted NLS Sequences for the Nuclear Localization
A sequence required for the nuclear localization may be only a part of the intact NLS sequence (see Note 31). Therefore, such a fusion approach is essential to determine the entire NLS sequence that can sufficiently target a recombinant protein to the nucleus. The suspected NLS sequence can be fused to either the amino (using the pEGFP-N vector series) or carboxyl (using the pEGFPC vector series) terminus of GFP. Inclusion of both types of the fusions can enhance the result of the interpretation. Vector construction, transfection, and fluorescent microscopy are the same as described above except that the antibody staining is not necessary and the protein localization can be observed in live cells.
4. Notes 1. Although the classical NLS sequences dominate the nuclear localization of nuclear proteins, nonclassical NLS sequences also play an important role in regulating many nuclear proteins and these types of NLS sequences may not be identified by the computer programs or sequence homology analysis. Therefore, step-wise terminal truncation in combination of internal deletion strategy is necessary to identify nonclassical, in most cases novel, NLS sequences from the amino terminal region of KLF8 as one example (Fig. 2). 2. To disrupt the mNLS sequences, point mutation of K or R to A or M is usually performed to disrupt the basic feature of the key residues. To disrupt the zinc fingers, the zinc-binding
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residues and/or DNA contact residues (16, 17) are often point-mutated. 3. Longer primers may be designed to meet the requirement for the G/C content or the 5¢-A/T or 3¢-G/C criteria. 4. Up to a half of the total amount of the purified fragments can be included in the second PCR reaction. 5. PCR reaction can be scaled up to 100 ml if necessary. 6. Add DNA polymerase to the reaction mixture last. 7. Pour slowly to avoid bubbles but quickly enough to avoid gel polymerization in the tube. Do not move the gel until it is solid. 8. Load the sample all the way down to the bottom of the well. Try to be stable. Put the pipette tip into the well closely to, but do not puncture, the bottom. 9. Check if the gel starts to run by looking at the electrophoresis-generated bubbles from the wires in the tank. 10. Be sure to cut the gel into little pieces so that it is easier to remove for purification. A little piece on the gel should be left for further reference by imaging. Make sure not to touch neighbor bands in order to avoid cross contamination. Thoroughly wash and clean the blade between different band cuttings. 11. The QG Buffer can be added a little more than 3 volumes to get both tubes to the same level so that they are balanced for centrifugation. 12. If gel slices from the same DNA band had to be transferred into more than one Eppendorf tubes in step 1, combine them now into the same column by repeating the centrifugation. 13. Most restriction enzymes digest DNA well at 37°C for 2 h. 14. Never trust the mechanical or electronic temperature indicator. Be sure to check for the accurate temperature using a mercury or ethanol thermometer. 15. Never return thawed competent E. coli back into freezer because the thawing/freezing operation dramatically reduces transformation efficiency. 16. Make sure sterile measures are taken during transformation experiments to avoid cross contamination by other bacteria and contamination of the culture medium. 17. Depending on vector’s feature, an antibiotic other than ampicillin (e.g., tetracycline) may be used. 18. 293 and Cos7 cells are the most frequently used cells to quickly test for expression due to their high transfection efficiency and protein expression capacity.
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19. Before moving on, make sure that lipofecamine particles are present in the medium by microscopy. 20. Vacuum remove the PBS residue from the wells to prevent the buffer to be added from over-diluting. 21. Avoid bubbles during comb inserting. 22. Do not take out the comb too early before the gel becomes solid to avoid formation of tortured wells. 23. Put the pipette tip against the tall glass plate and slide into the well and stop right before the tip reaches the bottom of the well. Apply slowly to prevent leak and bubbles. Load same volume of the 1× sample buffer alone into unused wells to avoid asymmetrical protein resolution such as “smiling lanes.” 24. It is critical to rub out all bubbles from the sandwich. 25. Alternatively, the transfer can be done overnight at 15 V. 26. For most primary and secondary antibodies, 1:500–1:2,000 and 1:5,000–1:10,000 dilution are recommended, respectively. 27. Pros and cons: the overnight strategy makes the antibody solution more reusable (more than five times on average) but takes longer, whereas the room temperature strategy is quick but makes the antibody less reusable. It is the matter of choice between saving time and saving reagent. Reuse of secondary antibodies is not recommended. 28. Fixed cells can be kept overnight or longer at 4°C. 29. Make sure not to drop or break the coverslips while transferring. 30. No bubbles should be present between the coverslips and the microscopic slide. 31. A sequence that is required but not sufficient for nuclear localization may represent just a part of the NLS sequence, a non-NLS sequence that plays a critical role in maintaining the NLS function or inhibiting the nuclear export of the protein. Additional experiments are helpful to test the effect of the sequence on the protein interaction with nuclear importin and exportin proteins and on the exportin dependent (e.g., Leptomycin B-sensitive) nuclear export.
Acknowledgments This research was supported by grants from American Cancer Society (#RSG CCG-111381) to JZ.
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References 1. Ghaleb AM, Nandan MO, Chanchevalap S, Dalton WB, Hisamuddin IM, Yang VW (2005) Kruppel-like factors 4 and 5: the yin and yang regulators of cellular proliferation. Cell Res 15:92–96 2. Narla G, DiFeo A, Fernandez Y et al (2008) KLF6-SV1 overexpression accelerates human and mouse prostate cancer progression and metastasis. J Clin Invest 118:2711–2721 3. Narla G, Heath KE, Reeves HL et al (2001) KLF6, a candidate tumor suppressor gene mutated in prostate cancer. Science 294:2563–2566 4. van Vliet J, Turner J, Crossley M (2000) Human Kruppel-like factor 8: a CACCC-box binding protein that associates with CtBP and represses transcription. Nucleic Acids Res 28:1955–1962 5. Wang X, Zheng M, Liu G et al (2007) Kruppel-like factor 8 induces epithelial to mesenchymal transition and epithelial cell invasion. Cancer Res 67:7184–7193 6. Wei H, Wang X, Gan B et al (2006) Sumoylation delimits KLF8 transcriptional activity associated with the cell cycle regulation. J Biol Chem 281:16664–16671 7. Zhao J, Bian ZC, Yee K, Chen BP, Chien S, Guan JL (2003) Identification of transcription factor KLF8 as a downstream target of focal adhesion kinase in its regulation of cyclin D1 and cell cycle progression. Mol Cell 11:1503–1515 8. Wang Z, Spittau B, Behrendt M, Peters B, Krieglstein K (2007) Human TIEG2/KLF11 induces oligodendroglial cell death by downregulation of Bcl-X(L) expression. J Neural Transm 114:867–875 9. Wang X, Urvalek AM, Liu J, Zhao J (2008) Activation of KLF8 transcription by focal
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adhesion kinase in human ovarian epithelial and cancer cells. J Biol Chem 283: 13934–13942 Wang X, Zhao J (2007) KLF8 transcription factor participates in oncogenic transformation. Oncogene 26:456–461 Ding Q, Grammer JR, Nelson MA, Guan JL, Stewart JE Jr, Gladson CL (2005) p27Kip1 and cyclin D1 are necessary for focal adhesion kinase regulation of cell cycle progression in glioblastoma cells propagated in vitro and in vivo in the scid mouse brain. J Biol Chem 280:6802–6815 Pandya K, Townes TM (2002) Basic residues within the Kruppel zinc finger DNA binding domains are the critical nuclear localization determinants of EKLF/KLF-1. J Biol Chem 277:16304–16312 Quadrini KJ, Bieker JJ (2002) Kruppel-like zinc fingers bind to nuclear import proteins and are required for efficient nuclear localization of erythroid Kruppel-like factor. J Biol Chem 277:32243–32252 Shields JM, Yang VW (1997) Two potent nuclear localization signals in the gut-enriched Kruppel-like factor define a subfamily of closely related Kruppel proteins. J Biol Chem 272:18504–18507 LaCasse EC, Lefebvre YA (1995) Nuclear localization signals overlap DNA- or RNAbinding domains in nucleic acid-binding proteins. Nucleic Acids Res 23:1647–1656 Corbi N, Libri V, Onori A, Passananti C (2004) Synthetic zinc finger peptides: old and novel applications. Biochem Cell Biol 82:428–436 Pabo CO, Peisach E, Grant RA (2001) Design and selection of novel Cys2His2 zinc finger proteins. Annu Rev Biochem 70:313–340
Chapter 11 Methods to Measure Nuclear Export of b-Catenin Using Fixed and Live Cell Assays Manisha Sharma and Beric R. Henderson Abstract b-Catenin is a multifunctional protein which is overexpressed in several types of cancers. The subcellular location of b-catenin at the membrane junctions or in the nucleus determines its function in cell adhesion and transcription activation, respectively. The aberrant localization of b-catenin in the nucleus activates transcription of a number of genes involved in cancer progression. Our laboratory studies intracellular trafficking of b-catenin with particular emphasis on its movement and regulation in and out of the nucleus. In this chapter, we outline a number of assays to measure the nuclear export of b-catenin in fixed and live cells. Key words: b-catenin, FRAP, Immunofluorescence, Leptomycin B, Nuclear export, Rev export assay
1. Introduction b-Catenin is an oncogene and is overexpressed in several types of cancers, in particular those that afflict the colon, breast, skin, liver and pancreas (1, 2). b-catenin was originally identified as a component of cellular adherens junctions where it binds the transmembrane protein, E-cadherin, and by indirectly regulating a-catenin dimerization modulates local actin filament bundling and dynamics at cell:cell junctions (3, 4). It is therefore a crucial regulator of cell membrane adhesion and the actin cytoskeleton. In primary tumors b-catenin is found in the nucleus of cells at the invasive front, whereas it is localized mainly at peripheral membrane in more central parts of a tumor (5). The loss of b-catenin from the membrane and its subsequent increase in the nucleus are recognized as a hallmark for the epithelial to mesenchymal transition of cells and progression of epithelial derived tumors (6). Paul J. Higgins (ed.), Transcription Factors: Methods and Protocols, Methods in Molecular Biology, vol. 647, DOI 10.1007/978-1-60761-738-9_11, © Springer Science+Business Media, LLC 2010
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The endogenous levels of b-catenin protein are tightly regulated by interaction with a multiprotein complex which phosphorylates and ubiquitinates b-catenin, thus tagging b-catenin for proteosome-mediated degradation (1). Mutations in genes encoding b-catenin, or its binding partners APC and axin, result in b-catenin stabilization and nuclear accumulation (2). In the nucleus, b-catenin binds and activates T cell factor (TCF) and Lymphoid Enhancer Factor (LEF) transcription factors and stimulates expression of a battery of genes involved in cell-cycle regulation, cell migration, and matrix metalloproteases (7). This abnormal nuclear accumulation of b-catenin and subsequent gene activation are thought to initiate cell transformation, and therefore cancer formation. Since the nuclear accumulation of b-catenin, like a range of other transcriptional regulatory factors, is associated with cancer progression, it is important to understand how b-catenin accumulates in the nucleus (8). b-catenin can shuttle in and out of the nucleus (9). We and others have shown that b-catenin is exported out of the nucleus by two distinct mechanisms: one is an indirect export mediated by binding to APC, which exits the nucleus via association with CRM1 (Exportin 1) via its nuclear export signal (NES) (10); the other pathway is a unique and poorly defined mechanism which is independent of CRM1 (11–13). Here, we outline techniques to study nuclear export of b-catenin.
2. Materials 2.1. Cell Culture
1. Mouse NIH 3T3 fibroblast, human SW480 colon carcinoma and human T47D breast cancer cell lines (American Type Culture Collection (ATCC)) are cultured in Dulbecco’s Modified Eagle’s Medium (DMEM; Invitrogen Corporation, Carlsbad, CA, USA) supplemented with 10% fetal bovine serum (Invitrogen Corporation), 2 mM glutamine and antibiotics (penicillin and streptomycin). 2. Cells were routinely passaged using a solution of 0.25% trypsin and 1 mM EDTA. Aliquots of trypsin/EDTA are stored at 4°C and frozen at −20°C for longer periods. 3. Leptomycin B (LMB; Sigma, St. Louis, MO, USA) is reconstituted according to the manufacturer’s instruction. 4. LipofectAMINE 2000 (Life Technologies, Inc.). 5. Opti-MEM media (Invitrogen Corporation). 6. Microscope cover glasses (Menzel GmbH + CoKG, Braunschweig, Germany). 7. Six-well trays (Becton Dickinson Labware, Franklin Lakes, NJ, USA).
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1. Phosphate-Buffered Saline (PBS) (10×); 1.37 M NaCl, 27 mM KCl, 100 mM Na2HPO4, 18 mM KH2PO4 (pH adjusted to 7.4 with HCl, if required). Dilute to 1× before use. 2. Formalin (Sigma): freshly diluted to 3.7% with 1× PBS. 3. Triton-X-100 (Ameresco Inc., Solon, OH, USA). Dilute to 0.2% with 1× PBS (see Note 1). 4. Bovine Serum Albumin (BSA, Sigma). Prepare 3% solution with 1× PBS. Stir and mix well. May add 0.01% sodium azide and store at 4°C for longer period. 5. b-Catenin monoclonal antibody (BD Transduction Lab), anti-mouse biotin (Dako), Avidin-conjugated to Texas red (Vector Lab), anti-mouse Alexa-Fluor-594 (Invitrogen), 0.05 mg/mL of Hoechst 33258. 6. Vectashield mounting medium (Vector Lab Inc., Burlingame, CA).
2.3. Nuclear Export Assays
1. Poly-l-lysine and digitonin (Sigma), creatine kinase and creatine phosphate, ATP and GTP (Roche Diagnostics, USA). 2. Cycloheximide and Actinomycin D (Sigma).
2.4. Nuclear Export Using Fluorescent Recovery After Photobleaching
1. Lab-Tek two- and four-well chamber cover glass slides (Nalge International, Rochester, NY, USA). 2. Human b-Catenin cDNA cloned into pEGFP-N1 vector (Clonetech, now Takara Bio Company, Mountain View, CA, USA).
3. Methods b-Catenin is a nuclear-cytoplasmic shuttling protein which functions in the nucleus, cytoplasm, and at the cell membrane. The localization of b-catenin varies in different cell lines; in nontumor cell lines (such as NIH 3T3 cells), it is predominantly seen at the cell–cell junctions, whereas in many colon cancer cell lines (e.g., SW480) it localizes in the nucleus and cytoplasm (Fig. 1). Using different assays, we can detect movement of b-catenin in and out of the nucleus. In this section we outline methods that involve use of epifluorescence microscopy and confocal microscopy to study the nuclear export of b-catenin in fixed and live cells. 3.1. Cell Culture, Transfection, and Drug Treatment
1. NIH 3T3 cells or SW480 cells are routinely maintained in 75 cm2 tissue culture flasks by passaging with trypsin/EDTA. At 24 h prior to transfection, cells are seeded on coverslips in a six-well tray such that they reach ~50% confluence at the time of transfection. One coverslip is used for each transfection.
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Fig. 1. Nuclear-cytoplasmic distribution of endogenous b-catenin. (a) NIH 3T3 and SW480 cells were fixed on coverslips and immunostained using b-catenin monoclonal antibody and Hoechst chromatin dye 33258 as outlined in Subheading 3.2. Cells were then analysed by fluorescence microscopy, and representative images are shown. (b) SW480 cells were seeded on coverslips and treated with digitonin and transport buffer as described in Subheading 3.3. Cells were then fixed and stained with b-catenin monoclonal antibody and images captured by fluorescence microscopy.
Prior to transfection remove the media and add 1 mL of complete media. 2. Mix 4 mL of lipofectAMINE reagent with 100 mL of OptiMEM media and incubate at room temperature for 5 min (tube “a”). In another microfuge tube, mix 2 mg of DNA with 100 mL of Opti-MEM media (tube “b”). Add contents of tube “a” into tube “b” dropwise and mix gently by pipetting (see Note 2). Incubate at room temperature for 20 min. Add this mix to cells in the appropriate well. Replace this media
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with complete media after 5 h (see Note 3). Incubate for 48 h at 37°C/5% CO2. Cells can then be immunostained (see Subheading 3.2). 3. Cells can be treated with LMB (a specific inhibitor of the CRM1 nuclear export pathway) at 6 ng/mL (diluted in the media) for 4 h. Use 1 mL of the diluted drug or a vehicle control per well. Optimum dose may vary according to the cell line. Cells are then ready for immunostaining (see Subheading 3.2). 3.2. Localization of b-Catenin by Immunofluorescence Microscopy
This is a general protocol to detect the steady state subcellular localization of b-catenin. The specificity of the antibody can be tested by transfecting b-catenin-GFP plasmid (increased staining of cells) as well as by transfecting siRNA for b-catenin (decreased stain). Cells stained with primary and secondary fluorescent conjugate antibodies are detected by use of a standard epifluorescence microscope. Figure 1a shows a typical staining pattern in NIH 3T3 cells and SW480 cells. NIH 3T3 cells show stronger accumulation at the cell membrane whereas SW480 cells show a stronger nuclear accumulation. The localization of the protein can be also confirmed biochemically by Western blot. 1. At 24 h prior to the experimental procedure NIH 3T3 and SW480 cells are seeded on coverslips in a six-well tray. One coverslip is required for each experimental procedure. Cells are processed when 50–100% confluent the next day, depending on the experimental conditions to be tested. Cells after transfection or drug treatments are stained using the following protocol. 2. Remove media and wash cells three times with PBS, then fix with 3.7% formalin (freshly diluted with PBS) for 20 min at room temperature. 3. Wash cells twice with PBS, then permeabilize with 0.2% triton-X-100/PBS for 10 min at room temperature. 4. Wash cells twice with PBS, then block with 3% BSA/PBS for 45 min at room temperature. (Do not wash after this.) 5. Apply 150 mL of b-catenin monoclonal antibody (primary antibody) diluted 1:100 for NIH 3T3 cells or 1:150 for SW480 cells in the blocking buffer. Incubate for 45 min at room temperature. Cells can be co-stained with other primary antibodies at the same time (see Notes 4 and 5). 6. Wash three times with PBS. 7. Apply 150 mL of anti-mouse biotin (1:500) or anti-mouse Alexa-Fluor-594 (1:1,500) (secondary antibody) diluted in the blocking buffer. Incubate 45 min at room temperature.
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8. Wash three times with PBS. 9. Apply 150 mL of avidin-texas red (1:800; only if anti-mouse biotin was used in step 8) and Hoechst diluted in the blocking buffer. 10. Wash three times with PBS. Mount the coverslip onto a drop of vectashield on the slide. The coverslips are dried very gently with tissue and sealed with clear nail polish. The slides are then viewed using a fluorescence microscope equipped with appropriate filter sets for detection of the fluorophore emissions, and are then stored at 4°C (for up to 2–3 months). 3.3. Nuclear Export of Endogenous b-Catenin in Permeabilized Cells
This assay enables the detection of nuclear export of endogenous cellular b-catenin in semipermeabilized cells, which display a strong nuclear level of b-catenin. Digitonin is a detergent that selectively permeabilizes the plasma membrane but not the nuclear membrane. Addition of an energy-generating transport buffer stimulates the energy-dependent active transport of proteins that can exit the nucleus in the absence of exogenously added nuclear transport factors (11). Nuclear export of proteins is generally energy (ATP) dependent and temperature sensitive. Figure 1b shows loss of nuclear staining from SW480 cells after treatment with digitonin and transport buffer. This assay requires use of a standard epifluorescence microscope. 1. Seed SW480 cells on coverslips in a six-well tray. Incubate overnight at 37°C/5% CO2 (see Note 6). 2. Preparation of digitonin: boil 500 mL of water and add 25 mg of high purity digitonin. Maintain heat while mixing slowly. Leave at 4°C overnight. Spin down and recover the supernatant. The saturated stock solution is taken as 1% solution (note that a precipitate will remain). Use at a final estimated concentration of 10–50 mg/mL depending on the cell line. The concentration needs to be optimized for each preparation of digitonin (see Note 7). 3. Preparation of transport buffer: 50 mM Tris–HCl (pH 7.5), 5 mM MgAc, 2 mM EGTA (pH 8), 50 mM KAc, 2 mM DTT. Freshly add the following protease inhibitors at final concentrations of: 50 mg/mL PMSF, 1 mM benzamidine, 50 mg/mL leupeptin. (a) Transport Buffer + LMB: Add higher dose (20 ng/mL) of LMB to the transport buffer. Incubate at 37°C/5% CO2 for 30 min prior to the export assay. (b) Transport Buffer + energy: Heat transport buffer to 30°C and add 1 mM of ATP, 0.5 mM GTP, 4 U/mL creatine kinase, 10 mM creatine phosphate just before use.
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(c) Transport Buffer – energy: Chill transport buffer to 4°C and add 10 mM sodium azide before use. 4. Wash cells three times with cold PBS. 5. Add 500 mL of digitonin per well. Always have a well without digitonin treatment as a control. Incubate on ice for 5–7 min (depending on the cell line). 6. Wash three times with PBS (see Note 8). 7. Add transport buffer +/− energy to respective wells. Incubate at 30°C for transport buffer + energy wells and at 4°C for transport buffer – energy wells for 30 min. (Make sure to seed cells in different trays for this step). 8. Wash twice with PBS. 9. Fix cells with 3.7% formalin and process for immunofluorescence microscopy as described in Subheading 3.2 (see also Fig. 1b). 3.4. Identification of Sequences with Nuclear Export Activity Using a TransfectionBased Assay
The first NES was identified in the HIV Rev protein. It comprises 11 amino acids (LQLPPLERLTL), where, leucines are critical for the export activity. Henderson and Eleftheriou developed a simple transfection-based assay to identify potential NESs. They constructed a GFP-tagged vector fused to a NES-mutated form of the Rev protein, called pRev(1.4)-GFP. This vector expresses a peptide that accumulates in the nucleus and is export defective (Fig. 2a, top panel); the addition of a functional Rev NES then translocates the GFP-reporter peptide to the cytoplasm (Fig. 2a, middle panel). This vector also contains a Rev nuclear localization
Fig. 2. Nuclear export assay using the Rev1.4-GFP vector. (a) T47D cells were transfected with the NES-defective plasmid pRev(1.4)-GFP, or the same construct into which was inserted DNA encoding the HIV Rev NES or the N terminal 132 amino acid sequence of b-catenin. At 48 h posttransfection, cells were untreated or treated for 3 h with 5 mg/mL actinomycin D (with or without 10 ng/mL LMB). All samples with treated with cycloheximide (CHX) to block new protein synthesis. Cell samples were then fixed and stained with Hoechst. (b) The proportion of cells with nuclear (N), nuclear-cytoplasmic (N/C) or cytoplasmic (C) localization was quantified and graphed.
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signal (NLS) and its nuclear retention (especially at the nucleolus) is disrupted by actinomycin D. This simple assay can be used to test putative NESs and compare the relative strength of different NESs (14). Figure 2a bottom panel shows minimal effect of LMB on nuclear export of the N terminus of b-catenin (amino acids 1–132). 1. Cloning of b-catenin sequence into pRev(1.4)-GFP vector: The N terminus of b-catenin (1-132 amino acids) has been shown to possess nuclear export activity (12). Therefore, we amplified a DNA sequence corresponding to the first 132 amino acids of b-catenin using PCR and cloned this fragment between the AgeI and BamHI sites of the pRev(1.4)-GFP vector. The plasmid (pb-catenin(1-132)-Rev(1.4)-GFP) integrity was confirmed by sequencing. 2. On day one, seed T47D cells onto coverslips in a six-well tray. T47D cells give a minimal cytoplasmic background localization when transfected with pRev(1.4)-GFP. Set up three wells for each construct and the pRev(1.4)-GFP vector. On day two, transfect 2 mg of the plasmid into each well using the protocol described in Subheading 3.1. 3. After 48 h treat each construct with: a. 20 mg/mL of cycloheximide (to prevent new protein synthesis) for 3 h at 37°C/5% CO2. b. Cycloheximide (as above in “(a)”) and 5 mg/mL of Actinomycin D (to prevent nuclear import of the cytoplasmic protein) for 3 h at 37°C/5% CO2. c. Cycloheximide and Actinomycin D (as in “(b)”) and 10 ng/mL of LMB (to prevent nuclear export) for 3 h at 37°C/5% CO2. 4. Wash cells with PBS three times and fix with 3.7% formalin for 20 min at room temperature. 5. Stain cells with Hoechst (prepared in 3% BSA–PBS) and mount with vectashield. 6. View and score cells by fluorescence microscopy as nuclear only, nuclear/cytoplasmic, or cytoplasmic accumulation of GFP-tagged peptide. The N terminal sequence of b-catenin was found to promote a modest degree of nuclear export and this was not blocked by LMB treatment (Fig. 2). 3.5. Nuclear Export of b-Catenin in Live Cells Using Fluorescence Recovery After Photobleaching
Cells are transfected with a GFP-tagged b-catenin expression plasmid (15) and then analysed with an inverted confocal microscopy system equipped with heated stage and live cell imaging, CO2 enclosure and the appropriate software for FRAP (fluorescence recovery after photobleaching) analysis of cells. Here we describe photobleaching assays using an Olympus FV1000 confocal live cell imaging system with Olympus software.
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1. Seed cells on a two well chamber cover glass (see Note 9) and transfect NIH 3T3 cells with GFP-tagged b-catenin using the protocol described in Subheading 3.1. (LMB may be added 3 h prior to imaging). The following protocol may be used for nuclear import also (see Note 10). 2. Settings of Olympus FV1000 confocal laser scanning microscope: All experiments are performed at 37°C with 5% CO2. Use 60× water objective; zoom = 2, scanning speed = fast, image format = 512 × 512, laser power during pre- and postbleach scanning is ~7–10% to minimize loss of fluorescence intensity, laser power during bleaching is 100%. Choose a region of interest (ROI) for bleaching covering ~90% of cytoplasm (e.g., using the polygon tool). Keep these parameters constant between cell samples (see Note 11). 3. Data acquisition: Start with five prebleach image scans of the whole cell. Bleach the ROI for 9–12 s (depending on the construct and stability of GFP tag in these cells). Postbleach imaging is done in three stages: first stage – 30 frames at fastest interval; second stage – 30 frames at 1.2 s interval; third
Fig. 3. Measurement of b-catenin nuclear export in live cells by FRAP analysis. (a) Subconfluent NIH 3T3 cells were transfected with b-catenin-GFP and after 48 h were then subjected to FRAP analysis with an Olympus FV1000 confocal microscope. In transfected cells, >90% 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
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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: S ignal 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
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.
35 35
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: 3.3. Protein–Protein Interaction Techniques 3.3.1. Coimmunoprecipi tation
Percent incorporation = (cpm of washed filter / cpm of unwashed filter) × 100. 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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3.4. Hydroxylation 3.4.1. HIF Hydroxylation
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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.
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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 Photo bleaching (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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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 27. 35S-L-Methionine, in vitro Translation Grade, >1,000 Ci/ mmol; >37 TBq/mmol (MP Biomedicals) 28. 35S-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
7. Matsumura T, Suzuki T, Aizawa K, Mune masa Y, Muto S, Horikoshi M, Nagai R (2005) The deacetylase HDAC1 negatively regulates the cardiovascular transcription factor Kruppel-like factor 5 through direct interaction. J Biol Chem 280:12123–12129 8. Oishi Y, Manabe I, Tobe K, Ohsugi M, Kubota T, Fujiu K, Maemura K, Kubota N, Kadowaki T, Nagai R (2008) SUMOylation of Kruppel-like transcription factor 5 acts as a molecular switch in transcriptional programs of lipid metabolism involving PPAR-delta. Nat Med 14:656–666 9. Chen C, Sun X, Ran Q, Wilkinson KD, Murphy TJ, Simons JW, Dong JT (2005) Ubiquitin-proteasome degradation of KLF5 transcription factor in cancer and untransformed epithelial cells. Oncogene 24: 3319–3327 10. Chen C, Sun X, Guo P, Dong XY, Sethi P, Cheng X, Zhou J, Ling J, Simons JW, Lingrel JB, Dong JT (2005) Human Kruppel-like factor 5 is a target of the E3 ubiquitin ligase
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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 <12 V, apply a set current of 100 mA per gel for 1 h. Ensure that the polarity is correct otherwise the proteins will be lost and the cathode plate may be damaged. 6. Stop the transfer, remove the cover and cathode plate and peel the 3MM paper and gel off the PVDF membrane. The pre-stained marker ladder should be clearly apparent on the membrane. Transfer the membrane to a shallow dish (protein face up) and incubate in 25–50 mL TBS-TM on a flat bed shaker for at least 30 min at RT. 7. Pipette 5 mL TBS-TM into a 50 mL Falcon tube; add the primary antibody to the recommended dilution (see Note 6). With a pair of tweezers transfer the membrane into the tube and incubate on the roller mixer over night at 4°C. 8. Transfer the membrane to a shallow dish and wash with five changes of 25–50 mL TBS-T for 5 min at RT (see Note 7). 9. After the final TBS-T wash, add enough TBS-TM to cover the membrane (10 mL is sufficient in a 100 × 80 mm dish) and add 1 µL of the HRP-conjugated secondary antibody (10,000-fold dilution) (see Note 8). Incubate on the flat bed shaker for 45 min at RT and then wash the membrane as before with five changes of 25–50 mL TBS-T for 5 min at RT. 10. Drain the TBS-T from the membrane and add 2 mL of freshly mixed ECL reagent evenly across the membrane. After 1 min transfer the membrane face down onto a clean acetate sheet (A5 size) and cover with a second acetate sheet (see Note 9). Image the membrane with a fluorescence imaging camera (10–30 min, depending on signal intensity) according to manufacturer’s instructions, or expose to ECL film in an X-ray cassette for an appropriate period of time. An example of the results produced is presented in Fig. 2. 11. The PVDF membrane can be stripped by incubation in 62.5 mM Tris–HCl pH 6.7, 100 mM 2-mercaptoethanol, 2% (w/v) SDS for 20 min at 65°C, followed by re-equilibration
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Fig. 2. Pre-initiation complexes on SRE promoters contain ERKs and MSK1. Nuclear extracts from HeLa cells stimulated with 15% FCS were used for PIC assembly on SRE (lanes 2 and 5), mutant SRE (DSE) (lanes 3 and 6) or TATA (lanes 4 and 7) templates. Complexes were isolated, washed and resolved by 10% SDS-PAGE for analysis. Aliquots of flow-through fractions were loaded in lanes 5–7. In this experiment the PVDF membrane was probed with four different antibodies in succession. The antibodies used and the order of application was as follows: MSK1, ERK1/2, SRF and MED23/Sur2. The sizes of molecular weight markers are shown to the left of the panels.
in TBS-T for 10 min at RT. It can then be re-probed with another antibody by repeating the procedure above from step 6.
4. Notes 1. The supernatant from the nuclear pellet can be worked up into a cytosolic extract following the original protocol (18) if required. 2. The dialysis tubing indicated only requires soaking in ddH2O for 30 min. 3. The magnetic beads are moved between tubes in an aqueous suspension and when taking aliquots, care must be taken to ensure the suspension is homogeneous. When in 1.5 mL microfuge tubes, beads are transferred quickly from one buffer to another by placing the tube into a magnetic holder, waiting a few seconds for the beads to move to the magnet, then removing the old buffer, replacing it with the desired one and removing the tube from the magnetic holder to resuspend the beads by shaking. 4. With some proteins we have observed high levels of binding to immobilised templates and a concomitant depletion of the
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protein from the unbound fraction. It may therefore be of interest to examine some of the unbound fraction(s) in parallel with the eluate(s). 5. At this stage immobilised PICs can be analysed for their transcriptional activity or for the presence of other specific enzymatic activities, for example phosphorylation or other post-translational modification of specific substrates. 6. Primary antibodies used were as follows: a sheep anti-MSK1 polyclonal (Upstate); a goat anti-ERK1/ERK2 polyclonal (C-14, Santa Cruz); a rabbit anti-SRF polyclonal (H-300, Santa Cruz) and a mouse anti-MED23/Sur2 monoclonal (D27, BD Biosciences). 7. The antibody solution in TBS-TM can be frozen and re-used at least once. 8. Secondary antibodies used were as follows: HRP-conjugated anti-sheep, anti-goat and anti-rabbit (Sigma). 9. To aid alignment of the molecular weight markers to the gel image when the outline of the membrane is not visible, tiny dots of fluorescent marker ink (Glow in the dark pen, Bostik) can be applied to the upper face of the acetate sheet with a needle or fine pipette tip to mark the corners of the gel.
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14. De Nadal E, Zapater M, Alepuz PM, Sumoy L, Mas G, Posas F (2004) The MAPK Hog1 recruits Rpd3 histone deacetylase to activate osmoresponsive genes. Nature 427:370–374 15. Pandey PK, Udayakumar TS, Lin X, Sharma D, Shapiro PS, Fondell JD (2005) Activation of TRAP/mediator subunit TRAP220/Med1 is regulated by mitogen-activated protein kinase-dependent phosphorylation. Mol Cell Biol 25:10695–10710 16. Ranish JA, Yudkovsky N, Hahn S (1999) Intermediaries in formation and activity of the RNA polymerase II preinitiation complex: holoenzyme recruitment and a postrecruitment role for the TATA box and TFIIB. Genes Dev 13:49–63 17. Drewett V, Molina H, Millar A, Muller S, von Hesler F, Shaw PE (2001) DNA-bound transcription factor complexes analysed by massspectrometry: binding of novel proteins to the human c-fos SRE and related sequences. Nucleic Acids Res 29:479–487 18. Dignam JD, Lebovitz RM, Roeder RG (1983) Accurate transcription initiation by RNA polymerase II in a soluble extract from isolated mammalian nuclei. Nucleic Acids Res 11:1475–1489
Chapter 19 Post-translational Modification of p53 by Ubiquitin Chunhong Yan Abstract Post-translational modification of p53 by ubiquitin resides in the center of a fine-tuned regulatory network that activates the tumor suppressor in response to genotoxic stress. Inhibition of p53 ubiquitination by DNA damage not only prevents p53 from degradation but also promotes its nuclear accumulation leading to transactivation of a number of downstream genes that are essential for the control of cell cycle progression, cell survival, and cellular senescence. Therefore, there are growing interests in studying p53 ubiquitination under physiological/pathological conditions. We describe herein a cell-based method and an in vitro reconstituted assay that are convenient, reproducible, and adaptable for various experimental conditions for detection of p53 ubiquitination. Wide application of these methods in studying mechanisms underlying regulation of p53 ubiquitination shall assist us in better understanding of the function of the tumor suppressor. Key words: p53, Ubiquitination, Ubiquitin, Post-translational modification, MDM2, Reconstituted assay system, Transfection
1. Introduction p53 is a guardian of the genome, and maintains genetic stability upon oncogenic challenges by transactivating a number of genes that are essential for the control of cell cycle progression, cell survival, and cellular senescence (1). The transcriptional activity of p53 is regulated by a growing number of proteins involving Posttranslational modifications (e.g., phosphorylation, ubiquitination, and acetylation) of the tumor suppressor (2, 3). Of these modifications, ubiqutination, a process adding ubiquitin moieties to p53, downregulates not only the stability but also the nuclear accumulation of the tumor suppressor (4, 5). Consequently, p53 ubiquitination resides in the center of a fine-tuned network that activates p53 in response to genotoxic stress (6). Indeed, whereas p53 remains at a low level in quiescent, unstressed conditions, DNA damage induces its phosphorylation (via ATM/ATR) and acetylation Paul J. Higgins (ed.), Transcription Factors: Methods and Protocols, Methods in Molecular Biology, vol. 647, DOI 10.1007/978-1-60761-738-9_19, © Springer Science+Business Media, LLC 2010
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(through p300/pCAF), which in turn disrupt the interaction of p53 with its major E3 ubiquitin ligase MDM2 thereby activating p53 by preventing it from ubiquitin-mediated degradation (7, 8). In a similar vein, we have shown that ATF3, a common sensor of DNA damage, participates in the genotoxic response through binding p53 and blocking its ubiquitination (9). Protein ubiquitination is an ATP-dependent process involving three sequential steps catalyzed by E1 ubiquitin-activating enzymes, E2 ubiquitin-conjugating enzymes, and E3 ubiquitin ligases, respectively (10). Currently, there are two widely used methods for studying p53 ubiquitination. The first method uses antibodies or nickelcharged resin to pulldown p53 that is coexpressed with ubiquitin and an E3 ubiquitin ligase such as MDM2, and subsequently detects modified p53 with immunoblotting. While this cell-based, “in vivo” method is convenient, cell context may compound the interpretation of experiment results. The second method, as a complement to the in vivo method, reconstitutes p53 ubiquitination reactions in vitro using purified E1, E2, and E3 enzymes, followed by detection of modified p53 with either radiography (11) or immunoblotting (9). Since most of the ubiquitination reaction components (URC) are commercially available, the latter method is accommodated by most laboratories providing that the investigated effectors are in defined forms (e.g., purified proteins, synthesized peptides, or chemical agents). Ideally, both “in vitro” and “in vivo” methods need to be utilized to delineate the regulation of p53 ubiquitination under various physiological/pathological conditions.
2. Materials 2.1. Cell Culture, Transfection, and Lysis
1. RPMI 1640 Medium supplemented with 10% fetal bovine serum, and Opti-MEM I medium (Invitrogen, Carlsbad, CA). 2. H1299 cells (see Note 1). 3. Trypsin (0.25%)/EDTA solution. 4. Fugene 6 (Roche, Indianapolis, IN) (see Note 2). 5. Phosphate buffered saline (PBS): 0.8% (w/v) NaCl, 0.02% (w/v) KCl, 0.144% (w/v) Na2HPO4, 0.024% (w/v) KH2PO4, pH 7.4. Store at 4°C. 6. MG-132 (Calbiochem, Gibbstown, NJ): dissolved in DMSO at 20 mM. Store in aliquots at −20°C. 7. Lysis buffer: 2 mM Tris–HCl, pH 7.5, 5 mM EDTA, 150 mM NaCl, 1% NP-40, 1% (w/v) sodium deoxylcholate, 0.025% (w/v) SDS. Store at 4°C. Before use, dissolve 1 Mini Protease Inhibitor Cocktail Tablet (Roche) into 10 ml Lysis buffer, and add MG-132 to a final concentration of 1 mM.
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8. RIPA buffer: 50 mM Tris–HCl, pH 7.4, 1 mM EDTA, 150 mM NaCl, 1% NP-40, 0.5% (w/v) sodium deoxylcholate, 0.1% (w/v) SDS. Add protease inhibitors and MG-132 before use as above. 9. Expression constructs for p53-myc (c-myc-tagged p53 protein), MDM2, and ubiquitin. 2.2. SDS-Polya crylamide Gel Electrophoresis (SDS-PAGE)
1. Tris–HCl buffers: 1.5 M pH 8.8 and 1.0 M pH 6.8. Store at 4°C. 2. 30% (w/v) acrylamide/bis solution (29:1) (Bio-Rad, Hercules, CA). 3. 10% (w/v) SDS. 4. N,N,N,N ¢-tetramethyl-ethylenediamine (TEMED) (Invit rogen). 5. 10% (w/v) ammonium persulfate (AP) solution: dissolve 1 g of AP in 10 ml H2O. Store at 4°C. 6. Running buffer: 25 mM Tris-HCl, 192 mM glycine, 0.1% (w/v) SDS. Store at 4°C. 7. 2× Sample Loading buffer: 100 mM Tris–HCl, pH 6.8, 4% SDS, 0.2% bromphenol blue, 20% glycerol. Store at room temperature. Add 1 M DTT to a final concentration of 100 mM before use. 8. Prestained molecule weight marker (Fisher).
2.3. Western Blotting for Ubiquitinated p53
1. Transfer buffer: 25 mM Tris-HCl, 192 mM glycine, 20% methanol, 0.0375% (w/v) SDS (see Note 3). Store at 4°C. 2. Nitrocellulose membrane (Bio-Rad), 3MM filter paper (Whatman, Maidstone, UK). 3. Tris-buffered saline with Tween (TBST): 0.24% (w/v) Tris, 0.8% (w/v) NaCl, adjust pH to 7.5. Add 500 ml Tween-20 before use. 4. Blocking solution: 5% (w/v) Blotto (Santa Crutz Biotechnology, Santa Crutz, CA) in TBST, or 3% (w/v) BSA (immunoblot grade, Santa Crutz). 5. Anti-Myc Agarose (Sigma, St. Louis, MI). 6. Anti-p53 antibody DO-1 (Santa Crutz) (see Note 4). 7. Anti-mouse IgG HRP conjugate (Promega, Madison, WI). 8. Enhanced chemiluminescent (ECL) reagents (Fisher).
2.4. In Vitro Reconstituted Ubiquitination Assay
1. E1 Ubiquitin-Activating Enzyme (UBE1) from rabbit (available from Boston Biochem, Cambridge, MA; see Note 5): diluted to 50 mg/ml with 50 mM Hepes (pH 7.6). Store at −80°C.
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2. E2 (UbcH5a) Ubiquitin-Conjugating Enzyme (Boston Biochem): diluted to 0.2 mg/ml with a buffer containing 50 mM Hepes (pH 7.6), 50 mM NaCl, 10% glycerol, and 1 mM DTT. Store at −80°C. 3. Human recombinant ubiquitin (Boston Biochem): dissolved in water at 5 mg/ml. Store at −20°C. 4. MgATP (Boston Biochem): 100 mM, store at −20°C. 5. 10× Ub buffer: 400 mM Tris–HCl, pH 7.5, 50 mM MgCl2, 20 mM DTT. 6. MDM2 protein (see Note 6). 7. p53 protein (Santa Crutz) (see Note 7): diluted to 10 mg/ml with PBS plus 5 mM DTT and 10% glycerol. Store at −20°C. 2.5. In Vitro Transcription/ Translation
1. TNT Quick Coupled Transcription/Translation System, which utilizes rabbit reticulocyte lysates to transcribe and translate proteins in vitro, is available from Promega. Upon receiving, dispense reticulocyte lysates into 40 ml aliquots and store them at −80°C. 2. p53-expressing construct (see Note 8). 3. RNase-free H2O: Add 1 ml of diethylpyrocarbonate (DEPC) into 1,000 ml purified H2O. Incubate the water overnight at 37°C, and then autoclave it for 20 min. Store at −20°C in aliquots.
3. Methods Steady-state levels of ubiquitin-modified proteins are often low in cells due to their rapid degradation by proteasomes. To measure levels of ubiquitinated p53 in vivo, cells need to be treated with MG-132, a proteasome-specific inhibitor, for a period ranging from 2 h to overnight to enrich for modified proteins. The in vivo method described here utilizes a c-myc-tagged p53 protein coexpressed with MDM2 and ubiquitin in p53-deficient cells by transfections. The p53 protein is then pulled down with agarose conjugated with the c-myc antibody, and subjected to immunoblotting with a p53-specific antibody to visualize the modified proteins that are often displayed as a number of slowly migratory bands unseen in conditions without MDM2 coexpression. Utilization of resin conjugated with an antibody to immunoprecipitate p53 is critical to the success of the experiments, because the IgG heavy chain with a size similar to p53 may interfere with immunoblotting results. This method can be modified in many ways. For example, p53 can be tagged with histidine and
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down-pulled with Ni+-NTA resin that binds histidine with high efficiency, or the ubiquitin moiety can be tagged with HA or FLAG to facilitate detection of modified p53. The in vitro assay reconstitutes the ubiquitination reaction by incubating the p53 protein with purified components of ubiquitination reactions (e.g., E1, E2, E3, ubiquitin, and ATP) and detecting ubiquitin modification using the p53-specific antibody DO-1. Commercially available p53 proteins are often purified from bacteria lysates and, hence, likely harbor conformations divergent from proteins of mammalian sources (12). In line with a notion that conformations may alter ubiquitination of the tumor suppressor (13, 14), we observed that the recombinant p53 protein was refractory to ubiquitin modification, yielding a limited number of ubiquitinated p53 bands (Fig. 1b).
Fig. 1. p53 ubiquitination detected in vivo and in vitro. (a) H1299 cells were transfected with indicated constructs expressing p53, ubiquitin, or MDM2 for 24 h, and then treated with 25 mM MG-132 or the solvent (DMSO) for 3 h before lysed for immunoprecipitations. Eluted proteins were subjected to immunoblotting to detect ubiquitinated p53 proteins. Modified p53 proteins are shown here as a number of slowly migratory bands above the major unmodified p53 band. (b) Five nanograms of recombinant p53 protein was incubated with indicated URC components at 37°C for 60 min, followed by immunoblotting to detect modified forms of p53. Asterisk indicates nonspecific bands that may be caused by impurity of the recombinant protein. (c) 0.5 ml of in vitro-translated p53 was incubated with other URC components as indicated, and subjected to immunoblotting. Ubiquitination reaction occurred without addition of E1 enzyme (the first lane), indicating that the rabbit reticulocyte lysate used for in vitro translation contained sufficient E1 enzyme for the ubiquitination reaction.
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In comparison, the p53 protein translated in vitro with rabbit reticulocyte lysates is more readily modified by MDM2-catalyzed ubiquitination reactions (Fig. 1c). We also found that rabbit reticulocyte lysates contain active E1 ubiquitin-activating enzymes sufficient for ubiquitination reactions (Fig. 1c), and therefore, the E1 protein can be omitted from the reconstituted reactions if p53 is obtained from in vitro translation. 3.1. Transfections and Prepration of Samples for Immunoprecipitation
1. H1299 cells are routinely cultured in RPMI 1640 medium, and passaged with trypsin/EDTA solution every 3–4 days. For transfections, the cells are detached from culture dishes with trypsin/EDTA solution and counted with a hematocytometer under an inverted microscope. Plate 6 × 105 cells onto 60-mm dishes, and cultured overnight (see Note 9). 2. In the following day, replace the culture medium with 4 ml fresh medium. 3. In a 1.5 ml tube, add 0.2 mg of the construct expressing p53-myc, 0.8 mg of ubiquitin, with or without 1 mg of MDM2 into 94 ml of Opti-MEM I medium (see Note 10). Mix well. Add 6 ml of Fugene 6 and leave at room temperature for 20 min. 4. Add the DNA-Fugene mixture to culture dishes drop by drop. Mix well and culture the cells in a CO2 incubator. 5. 24 h later, add MG-132 to a final concentration of 25 mM and treat the cells for 3 h to enrich ubiquitinated proteins. 6. Aspirate culture medium, and wash the cells for three times with cold PBS. 7. Add 500 ml of Lysis buffer and lyse the cells on ice for 20 min. Collect cell lysates into 1.5 ml tubes with cell scrapers. 8. Sonicate samples briefly (10 s, medium power output) with a cell sonicator. 9. Centrifuge the samples at 12,000 × g and 4°C for 15 min. Transfer supernatants to new tubes. The supernatants can be stored at −80°C for 1 week, or directly used for immunoprecipitation.
3.2. Immunopreci pitation
1. If samples are stored at −80°C, thaw out rapidly and remove precipitates that may be formed during storage by centrifugation at 12,000 × g at 4°C for 10 min. 2. Dilute 100 ml of samples with 400 ml of RIPA buffer. Leave samples on ice. 3. Pre-wash anti-myc agarose with 10 volumes of PBS for four times and then resuspend the agarose in an equal volume of PBS. Mix 40 ml of anti-myc agarose with each sample, and rotate the samples at 4°C for 4 h.
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4. Wash the agarose for three times using cold RIPA buffer at 4°C. For each wash, centrifuge tubes at 2,000 × g for 2 min, remove supernatants, resuspend the resin in 1 ml of RIPA buffer, and then rotate the tubes for 5 min. 5. After the last wash, centrifuge the tubes to remove trace amounts of RIPA buffer. 6. Add 40 ml of 2× Sample Loading buffer to each sample. Elute agarose-bound proteins by incubating the tubes in a boiled water bath for 10 min. 7. Centrifuge samples at 12,000 × g for 5 min. Use gel-loading tips to remove supernatants to new 1.5 ml tubes. Load supernatants immediately onto a polyacrylamide gel for SDS-PAGE. 3.3. SDS-PAGE and Western Transfer of Proteins
1. These instructions assume the use of Bio-Rad Mini-P4 system/Blot system and are adaptable for other commercial systems. First, prepare a 1.5-mm thick, 8% resolving gel (see Note 11) by mixing 3.7 ml of H2O, 2.2 ml of 30% acrylamide/bis solution, 2 ml of 1.5 M Tris–HCl (pH 8.8), 80 ml of 10% SDS, 80 ml of 10% AP, and 4.8 ml of TEMED. Pour the gel, and overlay with H2O. 2. Once the resolving gel polymerizes (~ 30 min), remove H2O on top of the gel. Prepare 5% stacking gel by mixing 2.7 ml of H2O, 0.67 ml of 30% acrylamide/bisacrylamide, 0.5 ml of 1 M Tri–HCl (pH 6.8), 40 ml of 10% SDS, 40 ml of 10% AP, and 4 ml of TEMED. Pour the gel solution and insert the comb. The stacking gel should polymerize within 20 min. 3. Assemble the gel unit, and add ~400 ml running buffer into the gel chamber and the tank. Carefully remove the comb. Load samples in wells with gel-loading tips; also load 5 ml of prestained molecular weight markers in a spare well to monitor gel running and transfer. Connect the unit to power supply, and run the gel at 150 V for 45–60 min until the dye front reaches the bottom of the gel. 4. Prepare a tray filled with transfer buffer. Cut a nitrocellulose membrane and four sheets of 3MM filter papers to a size equaling the resolving gel. Wet the membrane and the papers with transfer buffer. Stack two filter papers on top of the transfer-supporting foam provided by the Bio-Rad Blot system. Lay the nitrocellulose membrane on top of the papers. 5. Disconnect the power supply, and disassemble the gel unit. After removing the stacking gel, lay the resolving gel on top of the nitrocellulose membrane, and then cover the gel with two sheets of wet filter papers. As the transfer buffer contains SDS, air bubbles are easily trapped in the resulting sandwich. Carefully remove bubbles by rolling over the sandwich with a serological tube for several times.
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6. Assemble the transfer unit and connect it to power supply in a cold room or a chromatography refrigerator. Transfer the proteins at 100 V for 60 min. 7. Disassemble the transfer unit. Rinse the nitrocellulose membrane with 50 ml of TBST for two times. 8. Incubate the membrane in 50 ml of Blocking solution at room temperature with agitation for 1 h. Alternatively, the membrane can be blocked at 4°C overnight. 3.4. Immunoblotting for Detection of Modified p53 Protein
1. Prior to incubation with the primary antibody, rinse the nitrocellulose membrane with 50 ml of TBST twice. 2. Dilute the DO-1 antibody with TBST at 1:1,000. Incubate the membrane in the primary antibody solution at room temperature with agitation for 2 h. Alternatively, exposure to antibody can be performed at 4°C overnight. 3. Wash the membrane with 50 ml of TBST for three times (10 min for each wash). 4. Incubate the membrane with anti-mouse IgG-HRP conjugate (1:2,500 in TBST) at room temperature for 45 min. 5. Wash the membrane with 50 ml of TBST for three times (10 min for each wash). After the last wash, remove trace amounts of solution by blotting with Kim-Wipes. 6. Prepare 1–2 ml of ECL reagent based on the size of the membrane. Add the ECL reagent on top of the membrane and incubate for 1 min. 7. Remove the ECL reagent from the membrane. Blot the membrane with Kim-Wipes, and visualize the p53 proteins by exposure to X-ray film or using an imaging system as instructed by the manufacturer. An example result is shown in Fig. 1a. Modified p53 proteins are visualized as a number of slowly migratory bands, which are not apparent in the absence of MDM2.
3.5. Preparation of p53 Protein Through In Vitro Translation
1. Although recombinant protein is commercially available and can serve as a substrate for in vitro ubiquitination reactions, we found that it is more convenient to produce p53 protein by in vitro translation using rabbit reticulocyte lysates. For this purpose, p53 cDNA is subcloned in an expression vector (i.e., pcDNA3.1) that introduces a T7 primer sequence upstream of the p53-coding sequence. The TNT Quick Coupled Transcription/Translation System supplies reticulocyte lysates that can be used for preparation of p53 protein for in vitro ubiquitination reactions. 2. For in vitro translation, add 1 mg of the p53-expressing construct into a 1.5 ml tube containing 40 ml of rabbit
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reticulocyte lysate and 1 ml of methionine provided by the TNT system. Add RNase-free H2O to a total volume of 50 ml. Mix well and incubate the tube at 30°C for 60 min. 3. Store the reaction mixture at −80°C. 3.6. In Vitro Ubiquitination Assay
1. Thaw out the URCs including E1, E2, MDM2, recom binant or in vitro-translated p53 protein, MgATP and ubiquitin on ice. 2. In a 1.5 ml tube, add appropriate amount of H2O (to make the total reaction volume of 20 ml) and 2 ml of 10× Ub buffer. 3. Add 25 ng of E1, 100 ng of E2, 4 mg of ubiquitin, 100–500 ng of MDM2, 5 ng of recombinant p53 or 0.5 ml of in vitro-translated p53 prepared above, and MgATP to a final concentration of 2 mM. Incubate the tubes at 37°C for 60 min. 4. Stop the reactions by adding 20 ml of 2× sample loading buffer and heating in a boiling water bath for 5 min. 5. Load samples on 8% polyacrylamide gel to resolve proteins and detect p53 ubiquitination as above. An example result is shown in Fig. 1b (for recombinant p53 protein) and Fig. 1c (for in vitro-translated p53 protein).
4. Notes 1. The human lung cancer cell line H1299 is null for the p53 gene, and commonly used for studying p53 regulation. Other p53-null cell lines that have high transfection efficiencies (e.g., PC3) can be used to replace H1299 cells, but the transfection conditions need to be optimized accordingly. 2. We use Fugene 6 to transfect H1299 cells because this reagent yields high transfection efficiency with little toxicity to the cells. Other transfection reagents such as Lipofectamine 2000 (Invitrogen) can also be used. Follow the manufacturer’s protocol for transfections if another transfection reagent is used. 3. A low concentration of SDS in the transfer buffer promotes modified p53 (high molecular weight) to transfer to nitrocellulose membranes. 4. This antibody is also available from other commercial resources. Other p53 antibodies such as Pab 1801 can be used for this experiment. 5. The rabbit reticulocyte lysates contain E1 enzyme as shown in Fig. 1c. If in vitro-translated p53 protein is used for ubiquitnaiton reactions, this enzyme can be omitted from the assays.
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6. Purified MDM2 protein can be purchased from various commercial sources. For example, Boston Biochem sells histidine-tagged MDM2. We used GST-fused MDM2 protein that was expressed in E. coli and purified with glutathione agarose. We assume that a commercially available protein will also work in the described assay, providing that it is purified and preserved well. 7. This is a GST-fusion protein prepared from bacteria lysates. An alternative source is the in vitro-translated protein prepared using the TNT Quick Coupled Transcription/Translation assay. As discussed in Methods, in vitro-translated protein may preserve the native p53 conformation better than the bacterium-expressed protein, and thus can be more readily ubiquitinated by MDM2. 8. The region upstream of the p53-coding sequence must contain a T7 primer sequence to allow transcription initiation by rabbit reticulocyte lysates. Many mammalian expression vectors including the widely used pcDNA3.1 series contain such a T7 primer sequence. 9. The cells need to reach 50% confluence on the day of transfections. This cell density is optimized for Fugene 6 transfection reagents. If another transfection reagent is used, the plating cell density needs to be optimized following the instruction of the manufacturer. 10. The ratio between the constructs expressing MDM2 and p53-myc needs to be larger than 4:1 to ensure sufficient MDM2 expression. Like other E3 ubiquitin ligases, MDM2 expresses at a relatively low level in cells, presumably due to autoubiquitination-mediated degradation. The amounts of the constructs can be scaled down to allow for coexpression of a protein of interest so that the effect of the latter protein on p53 ubiquitination can be assessed (9). 11. Ubiquitin modification of p53 yields slowly migratory bands after electrophoresis. Therefore, 8% resolving gel is used to ensure better resolution of these ubiquitinated forms of p53.
Acknowledgments The author would like to thank Dr. Bert Vogelstein for providing the p53 and the MDM2 construct and Dr. Douglas D. Boyd for his advice and encouragement. This work was supported in part by Department of Defense grant PC061106.
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References 1. Levine A (1997) p53, the cellular gatekeeper for growth and division. Cell 88:323–331 2. Toledo F, Wahl G (2006) Regulating the p53 pathway: in vitro hypotheses, in vivo veritas. Nat Rev Cancer 6:909–923 3. Bode A, Dong Z (2008) Post-translational modification of p53 in tumorigenesis. Nat Rev Cancer 4:793–805 4. Geyer R, Yu Z, Maki C (2000) The MDM2 RING-finger domain is required to promote p53 nuclear export. Nat Cell Biol 2:569–573 5. Lohrum M, Woods D, Ludwig R, Balint E, Vousden K (2001) C-terminal ubiquitination of p53 contributes to nuclear export. Mol Cell Biol 21:8521–8532 6. Vogelstein B, Lane D, Levine A (2000) Surfing the p53 network. Nature 408:307–310 7. Vousden K (2002) Activation of the p53 tumor suppressor protein. Biochim Biophys Acta 1602:47–59 8. Appella E, Anderson C (2001) Posttranslational modifications and activation of p 53 by genotoxic stresses. Eur J Biochem 268:2764–2772
9. Yan C, Lu D, Hai T, Boyd D (2005) Activating transcription factor 3, a stress sensor, activates p53 by blocking its ubiquitination. EMBO J 24:2425–2435 10. Hershko A, Ciechanover A (1998) The ubiqtuitin system. Annu Rev Biochem 67:425–479 11. Fang S, Jensen J, Ludwig R, Vousden K, Weissman A (2000) Mdm2 is a RING fingerdependent ubiquitin protein ligase for itself and p53. J Biol Chem 275:8945–8951 12. Walerych D, Kudla G, Gutkowska M, Wawrzynow B, Muller L, King FW, Helwak A, Boros J, Zylicz A, Zylicz M (2004) Hsp90 chaperones wild-type p53 tumor suppressor protein. J Biol Chem 279:48836–48845 13. Nie L, Sasaki M, Maki C (2007) Regulation of p53 nuclear export through sequential changes in conformation and ubiquitination. J Biol Chem 282:14616–14625 14. Sasaki M, Nie L, Maki C (2007) MDM2 binding induces a conformation change in p53 that is opposed by heat-shock protein 90 and precedes p53 proteasomal degradation. J Biol Chem 282:14626–14634
Chapter 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 Abstract The chromatin organizer SATB1 regulates distant genes by selectively tethering matrix attachment regions (MARs) to the nuclear matrix. Post-translational modifications (PTMs) are important regulators of functional activities of proteins. Recently, a phosphorylation-dependent molecular switch that provided insights into the molecular mechanism of transcriptional repression and activation by SATB1 was discovered. SATB1 is specifically phosphorylated by protein kinase C (PKC) at serine 185 in vivo, and this modification leads to repression of transcription by SATB1 via increased association with the histone deacetylase 1 (HDAC1) corepressor. In this chapter, we describe methods for overexpression and purification of full length SATB1 protein and for its in vitro phosphorylation. We also describe method for in vivo phosphorylation of SATB1 upon immunoprecipitation using anti-SATB1. Finally, we describe a functional assay to monitor the effect of phosphorylation on transcription activity of SATB1 in vivo using MAR-linked reporter assay, in the presence and absence of PKC inhibitors. Key words: SATB1, Phosphorylation, HDAC1, Protein kinase C, Chromatin organization, Gene expression, Immunoprecipitation, Reporter assay
1. Introduction Special AT-rich sequence binding protein 1 (SATB1), the global chromatin organizer and transcription factor, has emerged as a key factor integrating higher-order chromatin architecture with gene regulation. SATB1 participates in the maintenance of chromatin architecture by organizing it into distinct loops via periodic tethering of matrix attachment regions (MARs) to the nuclear matrix (1–4). SATB1 organizes the T helper 2 (TH2) cytokine and MHC class-I loci into distinct chromatin loops (3, 4).
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SATB1 seems to play a role in dynamic organization of the transcriptionally poised chromatin (5). In addition to the well characterized role as an architectural component of chromatin, recently, SATB1 has also been shown to regulate genes by binding to the upstream regulatory elements and by recruiting chromatin modifiers (3, 4). SATB1 recruits several chromatin modifiers including ACF, ISWI, and HDAC1 (6, 7), and these chromatin modifiers were suggested to affect gene expression through histone modifications and nucleosome remodeling at SATB1-bound MARs (2, 6). SATB1 also regulates gene expression by recruiting corepressors (HDACs) and coactivators (HATs) directly to promoters (7, 8). In thymocyte nuclei, SATB1 forms a characteristic “cage-like” network that presumably demarcates heterochromatin from euchromatin (2). The ability of SATB1 to regulate transcription at global level stems from its unique ability to specifically bind to large number of regulatory sequences including promoters and MARs and recruit regulatory proteins to the binding site. How this is accomplished at the molecular level has become clear only recently. Various functional domains have been identified in SATB1 that together confer unique properties to this chromatin organizer. The ability of SATB1 to bind to a large number of genomic sequences in a sequence specific manner is conferred by a bipartite DNA-binding domain constituted by a cut repeat containing domain (CD) and a homeodomain (HD) in the C-terminal half (9, 10). Post-translational modifications of its N-terminal PDZlike domain act as molecular switches regulating the transcriptional activity of SATB1 via modulating its association with other proteins (8). The PDZ-like domain is also important for DNAand chromatin-binding ability of SATB1 through homodimerization (10, 11). The PDZ-like domain is indeed involved in interaction of SATB1 with other regulatory proteins such as the HDAC1 corepressor and the PCAF coactivator (7, 8). Post-translational modifications such as phosphorylation and acetylation have been shown to modulate interaction of many transcription factors with their interacting proteins as well as affect their DNA-binding affinities. Number of kinases and acetyltrasferases are involved in the phosphorylation and acetylation processes, respectively. Activity of many transcription factors is regulated through their interaction with coactivators, such as CBP (CREB-binding protein)/p300 (12) and PCAF/GCN5 (13). Phosphorylation also influences the association of different transcription factors with HDACs and HATs. Phosphorylation of NK-kB and p53 enhances their interaction with CBP/p300 (14, 15). Unphosphorylated MyoD showed higher interaction with PCAF (16), whereas dephosphorylation of pRb promoted the formation of pRb-HDAC1 complex in differentiated myotubes (17). Thus, the reversible and coordinated modifications of various
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transcription factors in the form of phosphorylation and acetylation seem to dictate their functional outcome. SATB1 is both phosphorylated and acetylated in vivo. Specifically, the residues K136 and S185 in the PDZ-like interaction domain are modified by PCAF acetyl transferase and protein kinase C (PKC), respectively (8). PKC is involved in divergent biological processes including transduction of extracellular signals during the T cells activation phenomenon (18). SATB1 is rapidly dephosphorylated during T cell activation (8). Interestingly, PKC itself is downregulated during T cell activation. More specifically, the PKC isoforms epsilon and beta are downregulated upon ionomycin treatment (19). SATB1 targeted genes are normally repressed due to recruitment of repressor complexes containing histone deacetylases (6–8) and are upregulated upon T-cell activation. Most importantly, phosphorylation status of SATB1 governs its mutually exclusive association with HDAC1 and PCAF (8). Thus, phosphorylation is a key Post-translational modification dictating the in vivo functions of SATB1. In this article, we describe methods to study the phosphorylation of SATB1 and its functional effects on the ability of SATB1 to regulate transcription. We also describe the method for purification of recombinant full length SATB1 protein for in vitro studies.
2. Materials 2.1. Purification of SATB1
1. Competent E. coli cells BL21(DE3) (Novagen, Gibbstown, NJ, USA; minimum efficiency 106 to 107 CFU per mg DNA). 2. Water processed with Milli Q (Millipore, Bedford, MA, USA) or double glass distillation. 3. LB (Luria-Bertani) ampicillin agar plates: 10 g bacto-tryptone, 5 g bacto-yeast extract, 10 g NaCl, and 15 g bacto-agar per liter containing 100 mg/ml ampicillin (see Note 1). 4. Isopropyl β-D-1-thiogalactopyranoside (IPTG) (SigmaAldrich Corp., St. Louis, MO, USA): dissolve in water to make 1 M stock and filter-sterilize by passing through 0.22 mm filter and distribute into 500 mL aliquots, store frozen at −20°C. 5. Ampicillin (Sigma-Aldrich Corp., St. Louis, MO, USA): dissolve in water at 50 mg/ml and filter-sterilize by passing through 0.22 mm filter and distribute into 500 mL aliquots, store frozen at −20°C. 6. Lysozyme: 20 mg/ml stock prepared in water. 7. Lysis buffer: 300 mM NaCl, 50 mM NaH2PO4, and 10 mM Imidazole.
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8. Wash buffer: 300 mM NaCl, 50 mM NaH2PO4, and 20 mM imidazole. 9. Elution buffer: 300 mM NaCl, 50 mM NaH2PO4, and 250 mM imidazole. 10. DC protein assay kit (Bio-Rad, Hercules, CA, USA). 11. LB broth containing 50 mg/ml ampicillin (Sigma-Aldrich Corp., St. Louis, MO, USA). 12. 6× loading buffer: 125 mM Tris–HCl, pH 6.7, 6% SDS, 20% glycerol, 10% 2-mercaptoethanol and 0.1% bromphenol blue. 13. Protease inhibitor cocktail, EDTA-free (Roche, Basel, Switzerland). 14. Nickel NTA agarose beads (Qiagen, Germantown, MD, USA). 15. Dialysis tubing, Snake skin (Pierce, Rockford, IL, USA). 16. Phosphate-buffered-saline (PBS) powder (Sigma-Aldrich Corp., St. Louis, MO, USA) is dissolved in 1 L water to make 1× solution or see Subheading 2.8. 17. Two by two ml disposable columns (Bio-Rad, Hercules, CA, USA). 18. Filters, 0.22 mm pore size (Millipore, Bedford, MA, USA). 19. Sonicator (Vibracell by Sonics, Newton, CT, USA). 2.2. SDSPolyacrylamide Gel Electrophoresis (SDS-PAGE) and Coomassie Brilliant Blue Staining
1. Separating buffer (4×): 1.5 M Tris–HCL, pH 8.7, 0.4% SDS. Store at room temperature. 2. Stacking buffer (4×): 0.5 M Tris–HCL, pH 6.8, 0.4% SDS. Store at room temperature. 3. Forty percent acrylamide solution (GE Healthcare, Uppsala, Sweden), 2% bis-acrylamide solution (GE Healthcare, Uppsala, Sweden) (see Note 2), and N,N,N,N ¢-Tetramethylethylenediamine (TEMED, Bio-Rad, Hercules, CA, USA). 4. Ammonium persulfate (GE Healthcare, Uppsala, Sweden): Prepare 10% solution in water. Store the solution at 4°C for 1 week. 5. Isopropanol (Sigma-Aldrich Corp., St. Louis, MO, USA). 6. Running Buffer (5×): 125 mM Tris–HCl, 960 mM glycine, 0.5% (w/v) SDS. Store at room temperature and dilute to 1× strength with distilled water prior to use. 7. Precision Plus prestained molecular weight markers (Bio-Rad, Hercules, CA, USA). 8. Fixer: (50% (vol/vol) methanol and 10% (vol/vol) acetic acid). 9. Coomassie Brilliant Blue (CBB) staining solution: 0.2% CBB in 50:40:10% methanol:water:acetic acid. 10. Destaining solution: (30% (vol/vol) methanol and 10% (vol/ vol) acetic acid).
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1. Recombinant protein kinase C, 25 mg/ml (Promega, Madison, WI, USA): single use aliquots should be made and stored at −80°C. 2. g32P-ATP (ICN, Santa Ana, CA, USA) specific activity 0.05 mCi/ml. (see Note 3). 3. Cold ATP (GE Healthcare, Uppsala, Sweden): 20 mM stock in water, pH adjusted to 7.0. 4. 5× PKC reaction buffer: 200 mM Tris–HCl, pH 7.5, 100 mM MgCl2, 2 mM CaCl2, 5 mM DTT (see Note 4). 5. Protease inhibitor cocktail EDTA-free (Roche, Basel, Switzerland). 6. Calf intestinal alkaline phosphatase (CIP) (New England Biolabs, Ipswich, MA, USA).
2.4. Drying of the Gel and Autoradiography
1. Cling wrap. 2. 3MM chromatography paper (Whatman, Maidstone, UK). 3. Bio-Max MS films (Kodak, Rochester, NY, USA). 4. Intensifying screen (Kodak, Rochester, NY, USA). 5. Gel dryer (Bio-Rad, Hercules, CA, USA).
2.5. Western Blotting of Phosphorylated SATB1
1. Transfer buffer: 20 mM Sodium phosphate buffer should be chilled to 4°C before setting up the transfer. 2. PVDF membrane (Millipore, Bedford, MA, USA) should be slightly bigger than separating gel. We use mini protean 3 system (Bio-Rad, Hercules, CA, USA) for which 9 × 11 cm membrane is appropriate. 3. 3MM chromatography paper (Whatman, Maidstone, UK). 4. Tris-buffered saline with Tween 20 (TBST), prepare 1× stock with 0.137 M NaCl, 25 mM Tris–HCL, pH 7.4, 0.1% Tween-20. 5. Blocking buffer: 5% nonfat dry milk in TBST. 6. SATB1 antibody (rabbit polyclonal generated in our lab or commercially available antibody from Abcam, UK) or phospho-serine antibody (mouse monoclonal, Sigma-Aldrich Corp., St. Louis, MO, USA) in TBST by diluting 1:1,000 and 1:2,000, respectively. 7. Secondary antibody: Anti-rabbit and anti-mouse IgG conjugated with horse radish peroxidase (Bio-Rad, Hercules, CA, USA) diluted in TBST to 1:20,000 and 1:3,000, respectively. 8. Dual substrate chemiluminescent reagent (Millipore, Bedford, MA, USA). 9. X-ray films (Kodak, Rochester, NY, USA).
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2.6. Cells, Cell Culture, and Treatments
1. Human embryonic kidney cell line (HEK 293T) obtained from American Type Culture Collection (ATCC, Manassas, VA, USA). 2. Dulbecco’s Modified Eagle’s Medium (DMEM) (Invitrogen, Carlsbad, CA, USA) supplemented with 10% fetal bovine serum (Invitrogen, Carlsbad, CA, USA) and 1× Penicillin– Streptomycin solution (Invitrogen, Carlsbad, CA, USA). 3. PKC inhibitors calphostin and rottlerin (Calbiochem, San Diego, CA, USA): 1 mM stock of each is prepared in DMSO, and single use aliquots are stored at −80°C. 4. Phosphate-buffered-saline (PBS) powder (Sigma-Aldrich Corp., St. Louis, MO, USA) is dissolved in 1 L water to make 1× solution. 5. Teflon scrapers (Nunc Inc. Rochester, NY, USA).
2.7. Immunoprecipitation (IP) of SATB1
1. Rapid extraction buffer (see Note 5): 20 mM Hepes, pH 7.9, 25% glycerol, 450 mM NaCl, 1.5 mM MgCl2, 0.2 mM EDTA, 0.5 mM DTT, 0.5 mM PMSF (see Note 6), 10 mM sodium orthovanadate (see Note 7) and all protease inhibitors (Roche, Basel, Switzerland). 2. Antibodies for IP: 1–2 mg of anti-SATB1 and normal rabbitIgG (as a negative control in immunoprecipitation). 3. Protein A/G beads slurry (Pierce, Rockford, IL, USA). 4. Washing buffer: PBS containing 0.1% Triton-X100. 5. Primary antibody: 1:2,000 dilution of anti-phosphoserine antibody (Sigma-Aldrich Corp., St. Louis, MO, USA) in TBST. 6. Secondary antibody: 1:3,000 dilution of anti-mouse IgG conjugated with horse radish peroxidase (Bio-Rad, Hercules, CA, USA) in TBST. 7. 2× loading buffer: 6× loading buffer diluted to 1: 3 with water.
2.8. Reporter Assay
1. Optiplates (Nunc Inc. Rochester, NY, USA). 2. Dulbecco’s Modified Eagle’s Medium (DMEM) (Invitrogen, Carlsbad, CA, USA) supplemented with 10% fetal bovine serum (Invitrogen, Carlsbad, CA, USA) and 1× PenicillinStreptomycin solution (Invitrogen, Carlsbad, CA, USA). 3. Phosphate buffered saline (PBS): Prepare 10× stock with 1.37 M NaCl, 27 mM KCl, 100 mM Na2HPO4, and 18 mM NaH2PO4. Adjust pH to 7.4 if necessary using HCl. Autoclave and store under sterile conditions. Prepare a 1× working stock (see Note 8). 4. DNA constructs to be transfected (see Note 9). 5. Lipofectamine 2000 reagent (Invitrogen, Carlsbad, CA, USA).
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6. Steadylite substrate (Perkin Elmer, Waltham, MA, USA) (see Note 10). 7. Plate based luminiscence counter (such as TOPCOUNT from Packard, Ramsey, NJ, USA).
3. Methods 3.1. Purification of SATB1
1. Transform competent E. coli BL21(DE3) cells with the pTriEX-SATB1 DNA construct (see Note 11) using the heatshock method and spread on LB ampicillin agar plate (see Note 12). 2. Inoculate a single colony in 5 ml LB media containing 50 mg/ml ampicillin and incubate at 37°C for 12 h (overnight) in a bacterial shaker incubator at 200 rpm. 3. Transfer 1 ml culture into 250 ml prewarmed LB media (37°C) supplemented with 50 mg/ml ampicillin. 4. Induce protein expression with 1 mM IPTG at an A600nm of 0.6– 0.8 for 3–4 h at 25°C. Prior to addition of IPTG, remove 5 ml aliquot as uninduced culture and keep the pellet in refrigerator. 5. Remove out an aliquot of 5 ml uninduced and induced culture and test for the expression level and solubility of the expressed protein by resolving the soluble bacterial lysate on SDS-PAGE. Meanwhile, pellet the rest of induced culture by centrifugation at 4,000 × g at 4°C for 10 min. Steps 6–8 describe the preparation of soluble and insoluble fractions (see Note 13). 6. Resuspend the cell pellet from 5 ml induced culture in 500 ml lysis buffer, and keep it on ice for 10–15 min. Sonicate the cell suspension until the sample becomes clear (e.g., using a miniprobe, 30% output power supply, and 2 S pulse for 1–2 min using the Sonics instrument). Centrifuge at 17,000 × g at 4°C for 15 min (see Note 14). 7. Transfer the supernatant (soluble fraction) to a new tube. Add 10 ml of 6× SDS loading buffer to 50 ml supernatant, and boil the sample for 10 min at 95°C. 8. Dissolve the pellet (insoluble fraction) in 1 ml of lysis buffer followed by centrifugation at 17,000 × g at 4°C for 15 min. 9. Transfer the supernatant to fresh tube. Take an aliquot of 50 ml lysate, and mix it with 10 ml of 6× SDS loading buffer. 10. Dissolve the pellet in 600 ml of 2× SDS loading buffer, and heat it for 10 min at 95°C. 11. Centrifuge the boiled samples at 15,000 × g for 10 min at room temperature.
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12. Prepare a 10% SDS-PAGE gel, and resolve the proteins by electrophoresis. Compare the protein profile of the soluble and insoluble fractions (see Note 15). 13. After confirming the protein in soluble fraction from steps 6 to 12, proceed to purification using the larger cell pellet. Resuspend the cell pellet (from the 245 ml induced culture) in 12 ml lysis buffer (10 ml lysis buffer per gram of wet pellet) and incubate on ice for 30 min or at room temperature for 15 min for adequate lysis of the cells. Lysis of the bacterial cell wall leads to an increase in viscosity. 14. The viscosity can be reduced by sonication until the lysate becomes clear (see Note 16). 15. Centrifuge the lysate at 17,000 × g for 30 min at 4°C. 16. Filter the lysate using a 0.22 mm syringe-driven filter. 17. Incubate 500 µl of Nickel NTA agarose beads with 12 ml of cleared lysate at 4°C for 30 min with proper mixing using a end-to-end rotator. 18. Pack the protein-bound beads in 2 × 2 ml Econo-column. 19. Allow the solution to drain out by gravity. 20. Wash the column three times with ten volumes of wash buffer. 21. Elute the fusion protein by passing 500 ml of elution buffer five times. Check the quality by running a small fraction of it on 10% gel (see Note 17) (see Fig. 1). 22. Pool all the elutions containing protein, and dialyze against PBS twice for 2 and 4 h each at 4°C. 3.2. SDS-PAGE and Coomassie Staining
1. These instructions assume the use of a mini protean 3 assembly system (Bio-Rad, Hercules, CA, USA); however, a comparable mini-gel apparatus from any manufacturer may be used. Clean the large and short plate thoroughly by using liquid detergent first followed then water. Finally, spray absolute ethanol and then dry them by wiping with lint-free tissue paper. Assemble the plates in plate holder and fix on casting stand. 2. Prepare a 10% gel by mixing 1.25 ml of 4× separating buffer, 1.24 ml of 40% acrylamide, 450 mL of bis-acrylamide, 2.0 ml distilled water, 50 ml ammonium persulfate solution and 5 ml TEMED. Pour the gel, leaving the space for stacking gel and overlay with isopropanol to ensure a flat surface and to exclude air. 3. Drain the isopropanol layer and rinse the top of separating gel with water.
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Fig. 1. Purification of recombinant SATB1 protein: Elution profile of 6× His tagged full length human SATB1 expressed and purified as described in Subheading 2.1. The figure shows Coomassie brilliant blue stained SDS-PAGE analysis of fractions from various stages of purification as indicated above each lane. Molecular weight standards in kDa are indicated on left. Full length SATB1 along with the 6× His tag migrates at approximately 110 kDa (marked by asterix). Induction level of SATB1 is moderate (lane 3), and the protein undergoes rapid degradation during purification and upon storage even in the presence of multiple protease inhibitors. The bands below full length SATB1 in the eluates (lanes 4–6) indicate the degradation products that accumulate rapidly during purification. The proportion of these bands with respect to the full length varies from preparation to preparation; however, in our experience, it has never been possible to obtain a preparation with more than 20% protein corresponding to the full length SATB1. A representative gel profile is depicted to give a realistic idea of the protein preparation. The degradation seems to occur by proteolysis at specific sites since most of these bands appear in different preparations of SATB1. The prominent bands observed at around 65–70 kDa are the most common degradation products of SATB1.
4. Prepare the stacking gel by mixing 500 mL of 4× stacking buffer, 200 ml of 40% acrylamide, 100 ml of bis-acrylamide, 1.18 ml distilled water, 20 ml ammonium persulfate solution, and 2 mL TEMED. Pour the gel and carefully insert the comb. 5. Prepare 1× running buffer by diluting 100 ml 5× running buffer with 400 ml of water. 6. Remove the glass plate assembly along with comb from the casting stand once the stacking gel is polymerized. Remove the comb, and wash the wells under running water.
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7. Arrange the plates with gel in electrode assembly, and fill the inner tank with running buffer. 8. Load 30 ml of sample in a well along with an unstained molecular weight marker in one well. 9. Complete the assembly of the gel unit, and perform the electrophoresis at 30 mA for 1 h such that the dye front reaches bottom of the gel. 10. Carefully detach the gel from glass plates and place it in clean staining box (glass/plastic) containing water. Rinse the gel with water followed by fixation in 30 ml fixer. Decant the fixer, add 30 ml of CBB staining solution, and incubate for 1–2 h at room temperature with gentle agitation. 11. Destain the gel with destaining solution until background of the gel turns to transparent and the bands are visible. 3.3. In Vitro Phosphorylation of SATB1
1. Prepare in vitro phosphorylation reaction mixture in a microcentrifuge tube on ice. Add the following: (a) 5 ml of 5× kinase reaction buffer. (b) 2.0 ml of PKC (50 ng). (c) 5.0 ml of ATP (cold or radiolabeled). (d) 1–5 mg of bacterially expressed and purified SATB1 protein. Adjust the final volume to 50 ml with water. 2. Supplement the reaction mixture with protease inhibitors. 3. Incubate the reaction mixture at 30°C for 30 min. 4. Terminate the kinase reaction by incubating at 70°C for 5 min. 5. Remove an aliquot of 10 ml reaction mixture, mix it with 10 ml of 2× loading buffer and boil for 5 min. 6. For confirmation of phosphorylation, treat one aliquot of the above reaction mixture with 10 units of CIP for 60 min at 37°C (see Fig. 2a). 7. Separate the reaction mixture by SDS-polyacrylamide gel electrophoresis on a 10% gel. 8. If radiolabeled ATP has been used, then remove the gel from glass plates and rinse it with 1× SDS running buffer (see Note 18). 9. Transfer the gel to 3MM Whatman chromatography paper, and cover it with cling wrap (see Note 19). 10. Dry the gel for 60 min at 80°C using a vacuum dryer (BioRad, Hercules, CA, USA). 11. Expose the gel to BioMax MS X-ray film at −80°C (usually 6–12 h). An intensifying screen can be used to enhance the signal (see Fig. 2b).
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Fig. 2. In vitro phosphorylation of SATB1: (a) Immunoblot analysis of recombinant 6× His-tagged SATB1 (lane 1) that was phosphorylated by PKC using cold ATP (lanes 2 and 3) as described in Subheading 2.3. Antiphosphoserine antibody was used to detect the in vitro phosphorylation. Half of the reaction mixture was treated with CIAP to digest the phosphorylated amino acid and the loss of immunoreactivity to antiphosphoserine confirms that the signal obtained in lane 2 corresponds to that of phosphorylated SATB1 (lane 3). (b) Autoradiogram depicting the phosphorylation of SATB1 using g 32P-ATP. Lane 1 represents unmodified SATB1, and lane 2 shows SATB1 phosphorylated in the presence of PKC. Upper band corresponds to the full length protein whereas lower band corresponds with one of its truncation product. For both panels, the position of band corresponding to SATB1 is marked by Asterix, and positions of the molecular weight standards are indicated on the left side in kDa.
12. If radiolabeled ATP is not used, then the proteins in the gel can be transferred to a PVDF membrane, followed by a standard immunoblotting analysis with anti-phosphoserine antibody (see Fig. 2a). 13. Make aliquots of the remaining labeled protein, and store at −80°C until needed for various biochemical assays. 3.4. Western Blotting
1. After gel electrophoresis, the proteins are electroblotted onto a PVDF membrane in transfer buffer at 4°C. 2. These directions assume the use of a Bio-Rad transfer assembly. In a plastic tray, PVDF membrane is wetted in methanol for 1 min, washed with water, and then saturated with transfer buffer. A tray of transfer buffer is prepared that is large enough to accomodate an opened transfer cassette with its two pieces of fiber pads and with one sheet of 3MM chromatography paper submerged on one side. 3. Remove the stacking gel from the separating gel while it is still on the glass plate. Apply a dry sheet of 3MM paper onto the separating gel so that wet gel sticks to the sheet, and
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carefully lift paper along with gel away from the glass plate and place it on a fiber pad with paper side touching the fiber pad. Fiber pad should already be placed on black flap of cassette (cathode side). 4. With a clean glass rod remove air bubble, if any, is trapped between the gel and the filter sheet. Then carefully place the PVDF membrane on the gel using flat end forceps. Cut a corner of the PVDF membrane for marking the orientation of the gel. Remove bubbles if any with glass rod. 5. Place one more prewetted sheet of 3MM paper on to the membrane, followed by a fiber pad. Close the cassette and transfer the assembly to a tank, which is filled with chilled transfer buffer. Keep a frozen cooling unit into the tank for to prevent over-heating. 6. Cover tank with lid such that electrodes are in contact and turn the power supply on. The transfer assembly is placed inside a 4°C refrigerator to avoid heating. Transfer of SATB1 can be accomplished within 1 h at 600 mA. 7. After transfer, remove the cassette from assembly and pick the membrane using flat end forceps. Block the membrane by incubating in 50 ml of blocking buffer for 1 h at room temperature on a rocking platform. 8. Discard the blocking buffer, and rinse the membrane in TBST. 9. Incubate with 1:2,000 dilution (in TBST) of antiphosphoserine antibody for 2 h in a sealed polythene pouch at room temperature. 10. Wash the membrane three times for 5 min each with TBST. 11. Incubate with a 1:3,000 dilution of HRP-conjugated secondary antibody in TBST for 45 min. 12. Discard the antibody, and wash the membrane three times with TBST for 5 min each. 13. In a dark room, prepare the luminescence detection reagent as per the manufacturer’s guidelines. 14. Add the mixed detection reagents directly to the membrane in a tray, and incubate for 1 min at room temperature. 15. Cover the membrane using cling wrap, and expose to X-ray film for different time intervals. 3.5. Immunoprecipitation of SATB1
Endogenously phosphorylated SATB1 can be detected using this assay. 1. Aspirate the culture media, and rinse the cells with ice cold 1× PBS. 2. Collect the cells by trypsinization or scraping and wash twice with 1× PBS.
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3. Centrifuge the cells at 2,000 × g for 5 min at 4°C. 4. Aspirate the PBS, and freeze the pellet in −80°C for 30 min. 5. Remove the cell pellet from the freezer, thaw rapidly, add five volumes of rapid extraction buffer and resuspend the cells by pipetting. Keep it on ice for 10 min, and repeat the pipetting. Lyse the cells by freeze thawing three times. 6. Centrifuge the lysate at 10,000 × g for 12 min at 4°C. Collect the supernatant into a fresh tube. 7. Centrifuge the supernatant at 142,000 × g for 30 min at 4°C. 8. Use 500 mg of total protein lysate in a final volume of 500 ml made up with 1× PBS containing 0.1% Triton-X100 and protease inhibitor cocktail mixture (see Note 20). 9. Add 1.0 mg of normal rabbit IgG to the lysate, and incubate at 4°C with rotation for 1 h. 10. Add 10 ml of protein A/G bead cocktail, and incubate at 4°C for 1 h on rocker. 11. Centrifuge at 500 × g for 5 min at 4°C. 12. Collect the supernatant in a fresh tube, and store the pelleted beads at 4°C. To the supernatant, add 1.0 mg of anti-SATB1 antibody. Incubate at 4°C for 2 h, add 10 ml protein A/G bead cocktail, and incubate for 2 h at 4°C. 13. Pellet beads by centrifugation at 500 × g for 5 min in a refrigerated centrifuge. 14. Wash the beads from steps 12 to 13 with PBS containing 0.1% Triton-X100. 15. Pellet the beads by centrifugation at 500 × g for 5 min. 16. Repeat steps 13–15 three times. 17. Add 20 ml of 2× loading buffer to the beads, mix by vortexing and heat denature at 95°C for 5 min (see Note 21). Load the sample onto a 10% SDS-PAGE gel. 18. After gel electrophoresis, the proteins are electroblotted onto a PVDF membrane followed by standard immunoblot analysis with antiphosphoserine antibody (see Fig. 3). 3.6. Treatments to the Cells and Reporter Assays
1. Culture HEK 293T cells in Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% fetal bovine serum at 37oC in 5% CO2 atmosphere. 2. Plate the cells in a 24 well culture plate 12–18 h prior to the transfection (see Note 22). 3. At 60% confluency, transfect the cells with DNA constructs using Lipofectamine 2000 as per the manufacturer’s instructions with slight modifications as follows (see Note 23).
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Fig. 3. Immunoblot analysis of in vivo phosphorylated SATB1: HEK 293T cells were treated with PKC inhibitors, rottlerin and calphostin. Cell extracts were then prepared and subjected to immunoprecipitation using anti-SATB1 as described in subheading 2.7. Phosphorylated SATB1 was detected by western blot using anti-phosphoserine. Lane 1 depicts the control IgG immunoprecipitate, and lane 2 shows the level of SATB1 phosphorylation at serine residue in control cells. Upon inhibition of PKC by rottlerin, SATB1 phosphorylation is reduced drastically (lane 3) and is completely lost upon calphostin treatment (lane 4). Position of band corresponding to SATB1 is marked by asterix. Positions of the molecular weight standards are indicated on the left side in kDa.
(a) Both DNA (0.25 mg/well) and Lipofectamine (2 ml/ well) are independently diluted in plain DMEM (25 and 50 ml, respectively) and incubated at room temperature for 5 min. (b) Mix the diluted DNA with Lipofectamine by rigorous pipetting, and incubate the Lipofectamine-DNA transfection mixture for 30 min at room temperature. (c) Aspirate the media from the cells just prior to transfection. (d) Gently layer the transfection mixture over the cells, and swirl the plate to ensure uniform spreading of the transfection mixture. (e) Incubate the cells with this transfection mixture for 6 h in an incubator at 37°C in 5% CO2 atmosphere. (f) Aspirate the transfection mixture after 6 h posttransfection, add 0.5 ml fresh DMEM with 10% FBS, and incubate for 48 h. 4. Intermittently monitor the growth of cells under microscope. 5. Treat the cells with either calphostin (25 mM) or rottlerin (10 mM), 1 h prior to cell harvest.
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6. Aspirate the media 48 h post transfection from each well. 7. Add 1 ml of chilled sterile 1× PBS, and collect the cells by scraping. 8. Pellet the cells by centrifugation at 2,000 × g for 5 min at 4°C. 9. Aspirate the PBS and resuspend each cell pellet in 50 ml of fresh PBS supplemented with 1 mM MgCl2 and 1 mM CaCl2 (see Note 24). The cell suspension is then transferred to the optiplate. 10. Add equal volume of Steadylite reagent to each well. Mix by pipetting, and incubate further for 10 min at room temperature (see Note 25). 11. Read the luciferase counts for each well by using a luminiscence counter, and plot the graph after calculating the fold change with respect to vector control (see Fig. 4).
Fig. 4. Regulation of SATB1 function by PKC mediated phosphorylation. In vivo luciferase reporter assay was performed upon cotransfection of SATB1 and IgH-MAR reporter construct (21) in HEK 293T cells (lane 1) as described in Subheading 3.6. As expected, SATB1 represses MAR-linked reporter activity (lane 2). Upon treatment with the PKC inhibitors calphostin (lane 3) and rottlerin (lane 4), the MAR-linked reporter activity is upregulated. The effect is more pronounced in calphostin treated cells than in rottlerin treated cells, presumably due to their differential effects on inhibition of phosphorylation of SATB1 (Fig. 3). Inhibition of phosphorylation of SATB1 leads to decreased affinity to MAR, therefore leading to derepression of MAR-linked transcription. Fold change in reporter activity is calculated with reference to that of the cells in which only the reporter vector (pGL3basic) is transfected. Each error bar represents the standard deviation calculated from three experiments.
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4. Notes 1. Prepare LB media or LB-Agar without ampicillin and autoclave. Allow the media to cool down such that while adding ampicillin the temperature of the media should not exceed 55°C. 2. Bis-acrylamide is neurotoxic in unpolymerized form, and therefore, mask and gloves should be worn while handling it. 3. pH of ATP is very important for efficient donation of g-phosphate to substrate protein. 4. Phosphorylation buffer should be made fresh each time. 5. A similar buffer is described in a protocol by Dignam et al. (20). The original method is very elaborate and time consuming and involves isolation of nuclei followed by extraction of nuclear proteins by incubation in this buffer, which is described as “buffer C” in the original method. 6. Add DTT and PMSF just before use. Dissolve PMSF in ethanol or isopropanol to prepare 0.25 M stock solution, add dropwise, and stir so that it will go into solution. 7. To prepare sodium orthovandate solution, boil 0.2 M stock solution until it becomes translucent; pH will be ~10. 8. 1× PBS is supplemented with 1 mM MgCl2 and 1 mM CaCl2 prior to use. 9. Supercoiled plasmid DNAs can be prepared by alkaline lysis using standard protocol and purified further using kits available from many commercial vendors. However, for long shelf life and best transfection efficiency and results, we recommend purifying the plasmid construct(s) by CsCl density centrifugation method. DNA purified by any of these methods is passed through sterile syringe-driven 0.22 mm filter membrane in biosafety hood. 10. Reconstitute Steadylite reagent just before use as per the manufacturer’s instructions. This reagent is light and temperature sensitive; hence once reconstituted, it should be stored away from light at −80°C. 11. We chose the pTriEX-3 Neo vector (Novagen, Gibbstown, NJ, USA) to clone full length human SATB1 because this expression vector enables rapid characterization of target genes in E. coli, insect and vertebrate cells and also allows rapid selection of stably transfected vertebrate cells expressing high levels of target gene. The 2.2 kb human SATB1 cDNA is cloned into the BamHI site in the multiple cloning site of this vector.
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12. The colonies on the plate should be present in moderate numbers and should be well spaced such that they grow sufficiently large to be picked singly and still do not merge into each other. 13. To minimize proteolysis, the cell pellet should not be stored frozen at −80°C for many days and should be preferably processed for purification immediately. 14. The volume of lysis buffer used for resuspension of the pellet can be varied depending on the size of the pellet obtained after centrifugation of the induced culture. The cells should be properly resuspended in the lysis buffer such that there are no clumps. 15. Samples should not be viscous before loading on SDSpolyacrylamide gels. Increase the boiling time or sonicate briefly to reduce the viscosity. After sonication, the samples should be clarified by centrifugation. While loading, the particulate material from bottom should be carefully avoided and only the top layer of the sample should be loaded in the wells. Although induction can be achieved at 25°C to a greater extent, lowering the temperature helps to keep the metabolic activity of the cell low and also ensures that the proteins are in a more soluble form than in the inclusion bodies. Shortening the duration of induction also helps to increase the soluble fraction of protein. 16. Keep the sample on ice during sonication and avoid frothing. Sonication generates heat, and therefore pulses should be given intermittently after a short period of incubation on ice during each break. For preparation of full-length SATB1, we recommend not to use enzymatic methods for clarification of the lysate. 17. SATB1 protein is very labile and is degraded rapidly; therefore, all steps should be performed on ice as quickly as possible and all buffers should contain freshly added protease inhibitors. 18. If kinase reaction is performed using radiolabeled ATP, then all apparatus that came into contact with the samples should be washed separately, the electrophoresis buffer should be discarded in appropriate waste container and the dried gel should also be disposed safely after autoradiography. Gloves and other protection for radiation exposure such as acrylic shield should be used throughout. 19. Cling wrap should cover the entire gel and the 3MM supporting sheet only on the top surface; otherwise, the gel may shatter into pieces during or after drying.
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20. Protease inhibitors and phosphatase inhibitors should be added while making up the volume. The dilution should be such that the final concentration of salt in the IP mix should be lower than 150 mM. 21. Never store the beads after immunoprecipitation without boiling them in SDS loading buffer. Once boiled in this buffer, the IPed products can be stored at −20°C for several weeks. 22. Seed cells at ~50% confluence so that they are not overgrown, 48 h post-transfection. 23. Since the HEK 293 cells are loosely adherent, aspiration of media and layering of the transfection mixture should be performed carefully. 24. Washing the cells with PBS is recommended to ensure complete removal of media. Media contain phenol red as pH indicator, which interferes with the reporter read out. 25. The reaction is performed away from direct light, and the plate is then sealed and covered with aluminium foil to avoid quenching.
Acknowledgments We thank Ranveer Jayani for help with protein purification and Prabhat Kumar Purbey for suggestions. Work was supported by grants from the Department of Biotechnology, Government of India and the Wellcome Trust, UK. Dimple Notani is supported by fellowship from the Council of Scientific and Industrial Research, India and Amita Limaye is supported by fellowship from the University Grants Commission, India. Sanjeev Galande is an international senior research fellow of the Wellcome Trust, UK. References 1. deBelle I, Cai S, Kohwi-Shigematsu T (1998) The genomic sequences bound to special AT-rich sequence-binding protein 1 (SATB1) in vivo in Jurkat T cells are tightly associated with the nuclear matrix at the bases of the chromatin loops. J Cell Biol 141:335–348 2. Cai S, Han HJ, Kohwi-Shigematsu T (2003) Tissue-specific nuclear architecture and gene expression regulated by SATB1. Nat Genet 34:42–51 3. Cai S, Lee CC, Kohwi-Shigematsu T (2006) SATB1 packages densely looped, transcriptionally active chromatin for coordinated
expression of cytokine genes. Nat Genet 38:1278–1288 4. Kumar PP, Bischof O, Purbey PK, Notani D, Urlaub H, Dejean A, Galande S (2007) Functional interaction between PML and SATB1 regulates chromatin-loop architecture and transcription of the MHC class I locus. Nat Cell Biol 9:45–56 5. Galande S, Purbey PK, Notani D, Kumar PP (2007) The third dimension of gene regulation: organization of dynamic chromatin loopscape by SATB1. Curr Opin Genet Dev 27:408–414
Phosphorylation-Dependent Regulation of SATB1, the Higher-Order Chromatin Organizer 6. Yasui D, Miyano M, Cai S, Varga-Weisz P, Kohwi-Shigematsu T (2002) SATB1 targets chromatin remodelling to regulate genes over long distances. Nature 419:641–645 7. Kumar PP, Purbey PK, Ravi DS, Mitra D, Galande S (2005) Displacement of SATB1bound histone deacetylase 1 corepressor by the human immunodeficiency virus type 1 transactivator induces expression of interleukin-2 and its receptor in T cells. Mol Cell Biol 25:1620–1633 8. Kumar PP, Purbey PK, Sinha CK, Notani D, Limaye A, Jayani RS, Galande S (2006) Phos phorylation of SATB1, a global gene regulator, acts as a molecular switch regulating its transcriptional activity in vivo. Mol Cell 22:231–243 9. Dickinson LA, Dickinson CD, KohwiShigematsu T (1997) An atypical homeodomain in SATB1 promotes specific recognition of the key structural element in a matrix attachment region. J Biol Chem 272:11463–11470 10. Purbey PK, Singh S, Kumar PP, Mehta S, Ganesh KN, Mitra D, Galande S (2008) PDZ domain-mediated dimerization and homeodomain-directed specificity are required for high-affinity DNA binding by SATB1. Nucleic Acids Res 36:2107–2122 11. Galande S, Dickinson LA, Mian IS, Sikorska M, Kohwi-Shigematsu T (2001) SATB1 cleavage by caspase 6 disrupts PDZ domainmediated dimerization, causing detachment from chromatin early in T-cell apoptosis. Mol Cell Biol 21:5591–5604 12. Chan HM, La Thangue NB (2001) p300/ CBP proteins: HATs for transcriptional bridges and scaffolds. J Cell Sci 114:2363–2373 13. Poux AN, Marmorstein R (2003) Molecular basis for Gcn5/PCAF histone acetyltransferase selectivity for histone and nonhistone substrates. Biochemistry 42:14366–14374
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14. Zhong H, Voll RE, Ghosh S (1998) Phosphorylation of NF-kappa B p65 by PKA stimulates transcriptional activity by promoting a novel bivalent interaction with the coactivator CBP/p300. Mol Cell 1:661–671 15. Lambert PF, Kashanchi F, Radonovich MF, Shiekhattar R, Brady JN (1998) Phospho rylation of p53 serine 15 increases interaction with CBP. J Biol Chem 273:33048–33053 16. Tintignac LA, Sirri V, Leibovitch MP, Lecluse Y, Castedo M, Metivier D, Kroemer G, Leibovitch SA (2004) Mutant MyoD lacking Cdc2 phosphorylation sites delays M-phase entry. Mol Cell Biol 24:1809–1821 17. Puri PL, Iezzi S, Stiegler P, Chen TT, Schiltz RL, Muscat GE, Giordano A, Kedes L, Wang JY, Sartorelli V (2001) Class I histone deacetylases sequentially interact with MyoD and pRb during skeletal myogenesis. Mol Cell 8:885–897 18. Hug H, Sarre TF (1993) Protein kinase C isoenzymes: divergence in signal transduction? Biochem J 291:329–343 19. Strulovici B, Daniel-Issakani S, Baxter G, Knopf J, Sultzman L, Cherwinski H, Nestor J Jr, Webb DR, Ransom J (1991) Distinct mechanisms of regulation of protein kinase C epsilon by hormones and phorbol diesters. J Biol Chem 266:168–173 20. Dignam JD, Lebovitz RM, Roeder RG (1983) Accurate transcription initiation by RNA polymerase II in a soluble extract from isolated mammalian nuclei. Nucleic Acids Res 11:1475–1489 21. Rampalli S, Kulkarni A, Kumar P, Mogare D, Galande S, Mitra D, Chattopadhyay S (2003) Stimulation of Tat-independent transcriptional processivity from the HIV-1 LTR promoter by matrix attachment regions. Nucleic Acids Res 31:3248–3256
Chapter 21 In Vivo and In Vitro Tools to Identify and Study Transcriptional Regulation of USF-1 Target Genes Marie-Dominique Galibert and Sébastien Corre Abstract In response to environmental stress, cells trigger a number of molecular mechanisms to control their survival and growth. These include changes in gene expression with corresponding Post-translational modifications to critical transcriptional-control proteins. Transcription is a highly-regulated process that is impacted by a large number of ubiquitous and specific factors. In order to determine how gene expression is regulated in response to environmental cues, it is necessary to correlate modifications to specific transcription proteins with an accurate assessment of the transcriptional response. This chapter details quantitative Real Time PCR (qPCR) and Luciferase assay protocols to illustrate, both in vivo and in vitro, the role of the USF-1 transcription factor in the UV-dependant regulation of pigmentation genes (POMC and MC1R). The procedures have been optimized for the USF-1 transcription factor and the regulation of specific target genes in response to physiological UV doses. Key words: USF transcription factor, Expression regulation, RT-qPCR, Luciferase assays
1. Introduction Promoter architecture, defined by the composition and the exact positioning of nucleosomes relative to transcription factor binding sites, dictates the capacity of a gene to respond appropriately to extracellular and developmental signals and is, therefore, critical to the maintenance of cell lineages. The establishment of correct promoter architecture is particularly important during DNA replication and on silencing of a promoter postactivation. Gene expression is dependent on chromatin remodeling and DNA occupancy at the different promoter regions to facilitate the recruitment of transcriptional machinery (1, 2). Transcription factors interact with different target genes during the cell cycle and promoter activation/shutdown to regulate and establish Paul J. Higgins (ed.), Transcription Factors: Methods and Protocols, Methods in Molecular Biology, vol. 647, DOI 10.1007/978-1-60761-738-9_21, © Springer Science+Business Media, LLC 2010
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the program of gene expression necessary for commitment, maintenance, and function of the cell lineage. Upstream Stimulating Factors, USF-1 and USF–2, are members of the eukaryotic evolutionary conserved basic-HelixLoop-Helix-Leucine Zipper transcription factor family (3). They interact with high affinity to cognate E-box regulatory elements (CANNTG), which are largely represented across the whole genome in eukaryotes (4). The ubiquitously expressed USF transcription factors participate in distinct transcriptional processes, mediating recruitment of chromatin remodeling enzymes and interacting with coactivators and members of the transcription preinitiation complex (PIC) (5–7). Results obtained from both cell lines and knock-out mice indicate that USF factors are key regulators of a wide number of gene regulation networks, including the stress and immune responses (8–10), cell cycle and proliferation (11–14), and lipid and glucid metabolism (15, 16). In melanocytes, USF-1 has been also largely implicated in vivo and in vitro as a key UV-activated regulator of genes associated with pigmentation (Tyrosinase, POMC, MC1R…) (17–19). Here, we propose to describe the required tools and techni ques used to establish the importance of USF transcription factors in the control of gene expression. Our presentation is focused on the USF-1 bHLH-LZ transcription factor, which is involved in cellular-stress response, following activation of the p38 kinase, by UV-radiation. Understanding the regulation of pigmentation gene expression has a physiological impact, as the tanning process constitutes one of the most important mechanisms at the skin level to protect the organism against UV-induced damage. 1. USFs transcription factors bind E-box regulatory elements, and although protein/DNA interaction suggests a strong transcriptional regulation, it remains to be proved. A combination of in vivo and in vitro assays (ChIP and EMSA) are required to map protein/DNA interactions in order to investigate the impact of USFs factors on transcriptional regulation of candidate E-box containing genes. 2. In vivo analysis of gene expression regulation is based on the use of real-time polymerase chain reaction (qPCR) to quantify specifically RNA levels of target genes (20). Real-time PCR monitors the progress of the PCR as it occurs (i.e., in real time). Data is collected throughout the PCR process in the exponential growth phase making this technique ideal for gene expression studies. Real-time chemistries, with the SYBR® Green I dye are very sensitive and can detect PCR amplifica tion during early phases of the reaction. This provides a distinct advantage over traditional PCR methods that identify PCR amplification only at the final phase or end-point of the reaction. The delta-delta (DD) Ct method (fluorescence
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threshold) provides tight and accurate measurements of variation in the level of gene expression under different cell conditions and as a function of time of treatments. The accuracy of real-time PCR is dependent on the quality of the extracted RNA that should be checked before any investigation (Fig. 1). Gene expression is then normalized to an invariant transcript defined for each studied model, usually a housekeeping gene (GAPDH) or the 18S rRNA (Fig. 2). 3. Real-time RT-PCR is a highly sensitive and an efficient method to characterize weak variations of target RNA level between two different conditions. Using RT-qPCR, this protocol will illustrate that expression of two of the most important pigmentation genes, POMC and MC1R, is significantly increased in response to physiological UV irradiation using human 501mel melanoma cell lines. Induced expression is directly linked to the activation of the stress dependent p38 kinase as use of the specific p38 kinase inhibitor (SB203580) prevents induction. Furthermore, when 501mel cells express the T153E mutated USF-1 protein that mimics the transcriptionally-active phophorylated-USF-1 form, a significant increase in endogenous POMC and MC1R transcripts are detected by real-time PCR (Fig. 3). 4. To establish the definite transcription factors and the corresponding cis-regulatory elements involved in the regulation of target genes, in vitro transcription methods are valuable tools to use in combination to DNA-binding assessments (ChIP and EMSA). Luciferase assays allow for the determination of the interactions and transcriptional regulation of target promoter genes in a simple model (21, 22). This technique is based on linking a target promoter sequence upstream to a genetic reporter, firefly luciferase. The chemiluminescence reporter activity is very sensitive and directly correlates with promoter activation and transcription. Mutations to potential cis-elements are additional experiments required to identify the transcriptionally-active promoter elements. Coexpression of trans-regulatory factors provides for the definition of the involved transcriptional mediators. Luciferase assays are thus complementary to RT-qPCR assays. They confirm in vitro activation of transcription, define the synergic cooperation of transcription factors, and establish correlations between specific DNA binding-site occupancy and induction of gene expression. Using luciferase assays, this chapter describes a protocol to show that expression of specific target genes (POMC and MC1R) is specifically induced after UV induction or following the coexpression of the transcriptionally active USF-1 protein (T153E-USF-1). Mutating consensus and conserved E-box motifs, that interact in vitro with USF factors
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Fig. 1. Analysis of RNA purity and integrity. (a) After purification from cell extracts, quantification of RNA is performed with the Nanadrop™ 1000 spectrophotometer (Labtech). (b) The NanoDropTM 1000 Spectrophotometer quantifies accurately and reproducibly nucleic acid samples from 15 up to 3,000 ng/ml. Absorbances are given specifically at 260 and 280 nm in addition to the calculated 260/280 and 260/230 ratios and the absorbance curve. (c) The Agilent 2100 Bioanalyzer is a microfluidics-based platform for the analysis of RNA integrity. Assessment of RNA integrity is a critical first step in obtaining meaningful gene expression data, as the use of intact RNA is a key element for successful RT-qPCR analyses. (d) Profiles generated with the Agilent 2100 bioanalyzer, allow a visual inspection of RNA integrity, by the integration of the electrophoretic trace. (e) RNA integrity, scoring between 0 and 10, is determined by the entire electrophoretic trace, taking in account the ratio between the 18S and 28S signal but also the shape of the ribosomal peaks and the nature of the baseline. Typical RNA profiles presenting degradation marks with a RIN of 4.3, or intact RNA RIN of 9.6 are shown.
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(EMSA and supershift assays) permits the identification of the UV response elements and the USF-1 binding sites required for subsequent promoter regulation (Fig. 4). 5. Real-time PCR (qPCR) and Luciferase assays are complementary techniques, both required for gene expression analysis in response to environmental cues. They are straightforward methodologies when performed with respect to definite protocols and prerequisites (cell confluence, RNA quality, and qPCR settings). In addition, real-time PCR also constitutes a powerful tool for complementary in vivo studies associated with siRNA, shRNA, or knock-out cell lines experiments targeting the depletion of specific cellular factors.
2. Materials 2.1. Cell Culture Reagents
1. RPMI-1640 medium (Gibco BRL, Invitrogen). 2. Foetal Bovine serum FBS (PAA). 3. Penicillin-streptomycin antibiotics (Gibco BRL, Invitrogen). 4. Cold-PBS: 8 g NaCl; 0.2 g KCl; 1.15 g Na2HPO4⋅7H2O; 0.2 g KH2PO4 to make 1 L. 5. Specific p38 kinase inhibitor SB203580 (Sigma) Stratalinker (Stratagene).
2.2. RNA Purification, Quality Control, and Reverse Transcription (RT) Reaction
1. RNAlater RNA Stabilization Reagent (Qiagen). 2. NucleoSpin® RNA II (Macherey Nagel). 3. Nanodrop™ 1000 Spectrophotometer (Thermo Scientific) (http://www.nanodrop.com/nd-1000-overview.html). 4. Agilent 2100 Bioanalyzer (Agilent Technologies) (http:// www.chem.agilent.com/en-us/products/instruments/ lab-on-a-chip/2100bioanalyzer/pages/default.aspx). 5. 2100 Bioanalyzer. 6. RNA 6000 Nano total RNA (Agilent technologies). 7. SuperScript™ II Reverse Transcriptase (Invitrogen). 8. RNase Inhibitor RNaseOUT™ (Invitrogen). 9. Random hexamers 50 ng/µl (Invitrogen). 10. dNTP mix (10 mM each).
2.3. R eal Time PCR
1. Primers (10 mM): about 20 bases, about 50% GC, amplimers of 70–100 bp designed with Universal Probe Library (Roche: https://www.roche-applied-science.com/servlet/RCConfig ureUser?URL=StoreFramesetView&storeId=10305&catalo gId=10304&langId=-1&countryId=uk).
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Fig. 2. Primer-set validation and gene-expression analysis by real time PCR. (a) Specificity of qPCR assays is measured by the dissociation curve profile. Dissociation curves analysis, which is performed after a completed PCR, is extremely useful when selecting optimal primers and primer concentrations for a SYBR® Green qPCR. Two dissociation curves are observed in the present example: one narrow peak corresponding to the specific product (1) and one to primer-dimer formation (2). This typical primer-dimer formation curve indicates that lower primer concentrations should provide more optimal results with a larger linear dynamic range. (b) Primer efficiency also affects the amplification process and threshold of detectable fluorescence. To determine primer efficiency, construction of a relative standard curve should be done (plot of cDNA concentration versus Ct). Serial cDNA dilutions amplified under low efficiency conditions yield to a standard curve with a different slope than the one obtained under high efficiency amplification conditions. The blue standard curve has an efficiency of 100% (slope is −3.3) while the green one has an efficiency of 78% (slope is −4). An 90–110% efficiency is considered acceptable. (c) To properly evaluate PCR efficiency, a minimum of three replicates and a
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2. Power SYBR® Green, PCR Master Mix (Applied Biosystems). 3. Thermocycler used: Applied Biosystems 7900 HT Fast Real Time PCR System. 4. PCR-program: 1 × 95°C 2 min; 40×(95°C 10 min; 95°C 15 s; 60°C 1 min). 2.4. Transient Tranfections Assays of Recombinant Proteins and Promoter-Genes
1. USF-1 recombinant proteins are obtained from the wild type transcription factor cloned into the pCMV vector. Point mutations were introduced into the WT pCMV-USF-1 expression construct, using the Stratagene® QuickChange® Site-Directed Mutagenesis kit (Stratagene). 2. Gene-promoter sequences are isolated by PCR and cloned into the luciferase reporter pGL3-Basic plasmid (Promega). E-boxes are mutated with the GeneTailor™ Mutagenesis System (Invitrogen) using specific oligonucleotides (for the POMC and MC1R promoters the sequences are available on request). 3. FuGENE® 6 Transfection Reagent (Cat N°11814443001, Roche).
2.5. L uciferase Assays
1. Cold-PBS: 8 g NaCl; 0.2 g KCl; 1.15 g Na2HPO4⋅7H2O; 0.2 g KH2PO4 for 1 L. 2. Luciferase lysis buffer: 1 ml KPO4 pH 7.8, 200 µl 10% Triton X-100, 10 µl DTT 1 M to make 10 ml. 3. Luciferase Assay Reagent (Promega). 4. Centro LB 960 Luminometer (Berthold Technologies). 5. Bovine Serum Albumin (BSA). 6. Bradford Reagent (Sigma). 7. Spectrophotometer to read OD at 595 nm wavelength.
3. Methods 3.1. C ell Culture and RT-qPCR 3.1.1. Day 1 Procedures
1. Plate 1.0–3.0 × 106 501mel cells in 10 ml of RPMI-1640 medium supplemented with 10% FBS and 1% of penicillinstreptomycin in 10 cm dishes. Prepare one plate for each experimental condition. Cells are maintained in a controlled atmosphere (10% CO2, 37°C).
Fig. 2. ( continued ) minimum of 5 logs of template concentration are necessary. With a twofold dilution with 5 points (orange), the potential artifact is higher than for the tenfold dilution with 5 points (blue). (d and e) Considering the specific efficiency of the primer sets, analysis of the threshold cycle (Ct) for the different samples is used to calculate the variation of expression according to the DDCt method (ref). This value corresponds to the fractional cycle number at which the fluorescence passes the threshold (linear: d or log: e). (f) This table describes the required parameters to perform accurately gene expression analysis by real time PCR.
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Fig. 3. Analysis of regulation of pigmentation genes expression after UV induction by real time PCR. (a) Typical induction of POMC (pro-opio melanocortin) expression after physiological UVB induction of 501mel cells. Cell extracts are recovered upon an 8 h kinetic after UVB induction (200 J/m² Stratalinker™) for RNA extraction and gene expression analysis by real time PCR (Syber™ Green Applied Biosystems®). The variation of expression is calculated according to the DDCT method, using the G3PDH as internal control gene and noninduced cell as reference (0 h). Each biological condition is performed twice, and the data are presented as mean ± SEM. They are considered significant (*) if P < 0.05, using the two samples Wilcoxon test (S-PLUS 6, Insightful™). A significant increase in POMC expression is evident 3 h after UVB induction, which is dependent on p38 kinase pathway activation. Indeed, induction is prevented in the presence of the specific p38 inhibitor (SB203580 10 µM) in the culture medium. (b) Typical induction of MC1R (Melanocortin receptor), which is the membrane receptor of the agonist (a-MSH POMC) by melanocytes, after physiological UVB (200 J/m²) induction of 501mel cells. Comparable results as previously described for POMC gene expression are obtained for MC1R gene with a significant increase of expression after UVB induction, which is also dependent on p38 kinase activation. (c) Endogenous expression of the pigmentation POMC gene is analyzed after transfection of 501mel cells and overexpression of different
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2. When cells are attached (about 8 h), change medium with fresh RPMI-1640 supplemented with 1% of FBS (see Note 1) and 1% of penicillin-streptomycin. Maintain the cells in controlled atmosphere to reach 80% confluence on day 2 (UV-induction day). 3.1.2. Day 2 Procedures
1. In order to specifically inhibit the p38 kinase, add directly in the medium the SB203580 compound (10 µM final concentration), 30 min to 1 h before UV-induction. 2. After 1 h, remove the medium from the cells and retain under the hood. 3. Irradiate cells with defined UVB doses (80–200 J/m²). (Always check that the proper UV-bulbs are present in the Stratatlinker™ apparatus (Stratagene), for UVB induction, the wavelength is 314 nm). 4. Replace the stored medium onto the cells, and maintain them in the incubator until the harvesting time according to the defined kinetic (appropriate delay following UV induction). 5. At the defined time, wash cells twice with cold-PBS (2 × 10 ml), and add 1 ml of PBS (see Note 2). 6. Scrape cells and transfer to a sterile 1.5 ml Eppendorf tube (see Note 3). 7. Pellet the cells by centrifugation (2500 g), and remove the supernatant. 8. For RNA extraction, gently resuspend cell-pellets, by pipetting up and down several times with 350 µl Lysis Buffer RA1 that is maintained at room temperature. 9. At this step of the protocol, samples can be used immediately for RNA extraction or stored at −20°C in RA1 lysis buffer for less than 1 month before RNA extraction (see Note 4).
3.1.3. Day 3 Procedures
1. RNA purification is performed on cell-pellets recovered in RA1 lysis buffer using the specific NucleoSpin® RNA II kit, according to the manufacturer’s recommandations (http:// www.mn-net.com/Portals/8/attachments/Redakteure_Bio/ Protocols/RNA%20and%20mRNA/UM_TotalRNA.pdf). RA1 lysis buffer efficiently inactivates RNases, which are present in virtually all biological materials and creates appropriate binding conditions that favor adsorption of RNA
Fig. 3. ( continued ) USF-1 recombinant proteins (pCMV-USF1 -WT, -T153E, -T153A), corresponding to specific activated or inactivated forms of the transcription factor. At 48 h after transfection and UV irradiation (if done), cells are recovered for RNA extraction and expression analysis by real time PCR (Applied Biosystems®). In the presence of the UV-activated USF-1 transcription factor form (WT-pCMV-USF1 + UV), and following transfection of the T153E constitutive activated form independently of additional UV-stimulation, a significant increase in POMC gene expression is evident.
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Fig. 4. In vitro analysis of transcriptional activation by Luciferase assays. (a) Luciferase assays are performed in the presence of both POMC and MC1R gene promoter regions, cloned upstream of a Luciferase reporter (in the pGL3 vector), and after physiological activation of 501mel cells with UVB (200 J/m²). At 48 h after transfection of 501mel cells with the different reporter-constructs (WT or mutated E box motifs, previously described to be USF-1 binding site by CHIP and EMSA), and 6 h after UVB stimulation of cells, whole cell extracts are prepared in Luciferase lysis buffer.
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to the silica membrane. Contaminating DNA, which is also bound to the silica membrane, is removed with a DNase solution, by direct addition onto the silica membrane. Washing steps are performed with two different buffers to eliminate salts, metabolites and macromolecular cellular components. Pure RNA is finally eluted under low ionic strength conditions with RNase-free Water. The standard protocol allows the purification of up to 70 µg of total RNA per NucleoSpin® RNA II Column from up to 5 × 106 cultured cells or 30 mg of tissue. 2. After RNA purification, 1.2 µl of purified sample is used to quantify RNA, using Nanodrop™ 1000 Spectrophotometer (LabTech) (Fig.1a). This spectrophotometer detects and quantifies accurately very low level of RNA. It permits also to evaluate the presence of potential contamination (protein, solvent…) by calculating both 260 nm/280 nm and 260 nm/ 230 nm ratios, which have to be between 1.8 and 2 (Fig. 1b). 3. Controlling RNA integrity is essential to follow up with RT-qPCR and is performed using Agilent 2100 Bioanalyzer (Agilent Technologies) (Fig. 1c). This technology is based on the loading of RNA in microfluidic lab-on-a-chip. This capillary electrophoresis is an integrated system coupling electrophoresis to staining, detection, and signal integration. (Agilent recommendations can be found on the following site: http://www.chem.agilent.com/Library/usermanuals/ Public/G2938-90035_QuickRNA6000Nano.pdf). RNA integrity is based on the migration profile (Fig. 1d), and the RNA Integrity Number (RIN), scoring from 0 to 10, is calculated according to several parameters (e.g., the area under the 18S and 28S narrow-peaks, the baseline profile). Finally, the higher the RIN, the higher is the quality and integrity of the RNA sample (Fig. 1e). 4. First-Strand cDNA Synthesis Using SuperScript™ II RT; a 20-µl reaction volume is used for 1 µg of total RNA. 5. Add the following components to a nuclease-free microcentrifuge tube: 1 µl of Random Hexamers (50 ng/µl), 1 µg of total controlled RNA, 1 µl dNTP Mix (10 mM each), RNase free water to 12 µl.
Fig. 4. ( continued ) A Berthold Centro luminometer is used to measure luminescence of lysate aliquots (20 ml) and values compared with those obtained for the empty vector (empty-pGL3). Using this method, it is possible to identify the UV elements required for transcriptional activation of the POMC and MC1R genes, which is dependent on specific USF-1 binding sites. (b) Similar Luciferase assays are conducted in the presence of both POMC and MC1R gene promoter regions and after cotransfection of different recombinant USF-1 proteins in 501mel cells. These data illustrate that the activated form of USF-1 is involved in the regulation of both POMC and MC1R expression after specific interaction with the consensus E box sequences on the respective promoters. The increase in expression of pigmentation genes is linked to the presence of phosphorylated-USF-1 form on the E box-promoter sequences.
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6. Heat mixture to 65°C for 5 min and quick chill on ice. 7. Collect the contents of the tube by brief centrifugation and add: 4 µl of 5× First-Strand Buffer (250 mM Tris–HCl (pH 8.3), 375 mM KCl, 15 mM MgCl2), 2 µl of DTT 0.1 M, 1 µl of 0.1 M DTT, 1 µl RNase Inhibitor RNaseOUT™ (40 units/µl). 8. Mix contents of the tube gently. Incubate at 25°C for 2 min. 9. Add 1 µl (200 units) of SuperScript™ II RT, and mix by pipetting gently up and down. Incubate tube at 25°C for 10 min and then at 42°C for 50 min. 10. Inactivate the reaction by heating at 70°C for 15 min. The cDNA can now be used as a template for amplification by qPCR. 3.1.4. Day 4 Procedures
Real-Time PCR is based on the collection of data in the exponential growth phase of PCR, during which amplification and integration of fluorescent SYBR® Green I dye in double stranded DNA is proportional with the initial amount of DNA or RNA in the tube. 1. Primers design and validation: For real-time PCR, primer sequences were designed using the Universal ProbeLibrary Assay Design Center (Roche) (https://www.roche-appliedscience.com/sis/rtpcr/upl/index.jsp?id=UP030000). This software is appropriate to design quickly and efficiently primers from fasta gene-sequences or using genes accession number from NCBI Genebank or Ensembl. The Universal Probe Library software is based on designing primers for real-time PCR using specifically Syber Green I dye, which takes in consideration several parameters and settings to optimize their efficiency (more specific than Primer 3). Notably it detects exon–exon junction to minimize the risk of false assay signals, from remaining genomic DNA and avoids intronspanning assays targeting introns that are too short to be effective targets. It also compares the sequence designed against the whole genome to minimize the risk of false assay signals from unrelated transcripts generated by splice variants and homologous genes. Finally, it designs relatively short primers (20 bp), with the same melting temperature (60°C) and a 40–60% GC-content. 2. To perform real-time PCR, you have to first validate the speci ficity and the efficiency of the amplification for each primer-set. The specificity of the amplification is confirmed by the detection of only one narrow pick on the dissociation curve (Fig. 2a). 3. To validate that each primer sets presents the same amplification efficiency, you have to perform real-time PCR with different dilutions of the same cDNA sample on at least a
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5 log range. The correlation coefficient for standard deviation has to be as close as possible to 1 and the slope for the different curves has to be the same (−3.3) (Fig. 2b, the blue curve has a good efficiency). It is also important to have a wide range of dilutions (5 log) to avoid amplification problems with low or high levels of cDNA, linked to the integration of the fluorescent dye (Fig. 2c). 4. After validating the primer-sets, corresponding to each target genes and to the internal control (GAPDH), you can carry on with gene expression analysis. To analyze the gene expression level of a target gene according to cell conditions, you should compare the Ct values, which correspond to the threshold of detectable fluorescence (linear: Fig. 2d; log: Fig. 2e) and use the delta delta CT method for relative quantification of cDNA. 5. Real-time PCR is an efficient and powerful technique but requires specific recommendations to optimize the analysis and to compare expression, such as the amplification efficiency of the different primer-sets, the precision of analysis with at least 3 replicates, and also the sensitivity of the reaction, which is associated to different steps of the protocol, including the RNA integrity. Also, the choice of the internal control gene and the reference sample (T0) can change according to experimental settings. 6. Real Time PCR reaction is obtained by mixing 3 pmol 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) (http://www3.appliedbiosystems. com/cms/groups/mcb_support/documents/generaldocuments/cms_046776.pdf) to 1 ml of diluted cDNA or Input DNA samples. Complete with sterile H2O to a final volume of 10 ml. PCR is run in appropriate 96 well or 384 well plate with Applied Biosystems 7900 HT Fast Real Time PCR System, following PCR program (1 × 95°C 2 min; 40×(95°C 10 min; 95°C 15 s; 60°C 1 min). Each amplification is performed in triplicate. 3.1.5. Data Analysis
1. At the end of the qPCR run, the Applied Biosystems 7900 HT Fast Real software gives all the Ct values for the different genes and experimental conditions, with the average and standard deviation for triplicates (<0.16). The variation of expression for each target gene is calculated according to the DDCt method (http://www3.appliedbiosystems.com/ AB_Home/applicationstechnologies/Real-TimePCR/ AbsolutevsRelativeQuantitation/index.htm). Relative variation of expression = 2 (∆Ct1-∆Ct2),
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where: DCt1 = (Ct condition X − Ct reference (T0)) target gene (in our case POMC or MC1R) and: DCt2 = (Ct condition X − Ct reference (T0)) internal control gene (in our case GAPDH). 2. Each biological experiment is performed twice and each real-time PCR is done in triplicate for a good precision in order to be statistically significant. Statistical test, using the two samples Wilcoxon test (S-PLUS 6, Insightful™) is used to evaluate significant increase of target gene expression with a 5% confidence threshold. Here is described (Fig. 3a, b) classical response for the analysis of expression by real-time PCR for pigmentation genes in response to UV (POMC and MC1R). This technique allowed us to show that the mRNA level of the target genes are significantly increased 3–6 h post physiological UV-radiation (UVB). This increase is also linked to the activation of the stress dependent p38 kinase. In the presence of the SB203580 specific compound (10 µM), no increase occurs. 3.2. Luciferase Assay 3.2.1. Day 1 Procedures
3.2.2. Day 2 Procedures
1. Plate 5.0 × 104 501mel cells in 500 µl of RPMI-1640 supplemented with 10% Foetal Bovine Serum (FBS) and 1% of Penicillin-Streptomycin in 12 wells plate. Three distinct luciferase measurements are performed for each condition, so prepare proper number of wells for all the experiments. Cells are maintained overnight in a controlled atmosphere (10% CO2, 37°C) to reach 50% confluence on the transfection day. 1. In tube N°1, add 500 ng of the adequate pGL3-promoter-Luc construct according to the condition tested (control: empty pGL3-Luc, normal or mutated promoter-Luc), to the USF-1 expressing vectors (WT, or mutant USF-1) or not (empty pCMV), with 100 µl of serum and Pen-Strep free RPMI-1640 medium. Keep 5 min at RT. (see Note 5). 2. In tube N°2, add 3 µl of Fugene to 50 µl of serum and Pen-Strep free RPMI-1640 medium. Keep 5 min at RT. 3. Mix gently each tube N°1 (100 µl) with one tube N°2 (50 µl) and keep at RT for 20 min. 4. Add to each tube 50 µl of serum free RPMI-1640 medium, mix gently, and add drop wise onto the cells.
3.2.3. Day 4 Procedures
1. 48 h after transfection, wash the cells two times with cold PBS, add 100 ml of Luciferase Lysis buffer, and harvest cells by scrapping using 1 ml syringe tip. 2. Recover the cell lysate, and spin quickly (2500 g) to pellet the membranes. At this step of the protocol, samples can be stored at −80°C for few weeks before analysis. 3. Transfer 20 µl of each sample in a 96-well luciferase assay plate.
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4. Dissolve luciferase powder in 10 ml buffer (from Luciferase assay reagent kit). 5. After configuration, washes and prime of Centro LB 960 Luminometer, read 96-well plate for luminescence measure ment, by using automatic injection of luciferase reagent (see Note 6). 6. Each result is normalized to protein concentration (Bradford) and then triplicate corrected luminescence data are averaged. Here, we show typical luciferase assay data using POMC or MC1R promoters link to the luciferase reporter genes following transfection of 501mel cells (Fig. 4a, b). This technique confirms that both promoters are UV-induced in vitro and permits the identification of the UV response DNA elements within each promoter-sequence (Fig. 4a). Coexpression of WT or T153E- USF-1 form indicates that the transcriptionally active USF-1 protein is required in combination with the presence of specific E box element to drive transcription (Fig. 4b).
4. Notes 1. It is thoroughly recommended not to overgrow the cells before use, to passage them regularly, and to ensure that cell confluence will not exceed 80%. Serum should be removed to minimize the impact of growth factor and to allow visualizing the effect of cell stimulation. 2. It is important to recover in parallel to RNA extraction samples, cell-pellets in Laemmli buffer for protein analysis by western blot. 3. 2 ml syringe tips are quite useful for scraping and cheaper than rubber-policeman! 4. For RNA protection and long sample storage, cells can be submerged in an appropriate volume (300 µl) of RNAlater RNA Stabilization Reagent (Qiagen) immediately after harvesting. RNAlater quickly stabilizes and protects RNA expression patterns and ensures that downstream analyses reflect the expression profile of the intact cells. Samples can be archived at −20°C or −80°C for several months without risk of RNA degradation, even after multiple freeze–thaw cycles. This reagent is compatible with RNA extraction protocol, using NucleoSpin® RNA II kit from Macherey Nagel. 5. To compare accurately Luciferase data, the amount of transfected DNA per cells should be the same for each condition. DNA level should be adjusted by the addition of
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the corresponding empty vectors (empty pCMV vector). The amount of Fugene used should be adjusted to the amount of DNA transfected. Increasing concentration of recombinant protein permit to prove that transcription is dose dependent. In this case, DNA content has to be adjusted to remain identical. You can also analyse the cooperation of different factors. 6. If you do not use the dual luciferase system, which allows to adjust light signal with the level of transfected reporter plasmid in the cell, by measuring the two independent signals, the Luciferase reporter one (gene promoter specific) and the Renilla reporter (proportional of plasmid content but independent on transcription activation), you have to quantify protein level in the different samples. For this, proceed to protein quantification on one 10 µl aliquot of sample using the Bradford method, referring to standard curve established with serial dilution of BSA.
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. Lam FH, Steger DJ, O’Shea EK (2008) Chromatin decouples promoter threshold from dynamic range. Nature 453:246–250 2. Urnov FD, Wolffe AP (2001) Chromatin remodeling and transcriptional activation: the cast (in order of appearance). Oncogene 20:2991–3006 3. Corre S, Galibert MD (2005) Upstream stimulating factors: highly versatile stressresponsive transcription factors. Pigment Cell Res 18:337–348 4. Ferre-D’Amare AR, Pognonec P, Roeder RG, Burley SK (1994) Structure and function of the b/HLH/Z domain of USF. EMBO J 13:180–189 5. Bungert J, Kober I, During F, Seifart KH (1992) Transcription factor eUSF is an essential component of isolated transcription complexes on the duck histone H5 gene and it mediates the interaction of TFIID with a
6.
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TATA-deficient promoter. J Mol Biol 223: 885–898 Chiang CM, Roeder RG (1995) Cloning of an intrinsic human TFIID subunit that interacts with multiple transcriptional activators. Science 267:531–536 Meisterernst M, Horikoshi M, Roeder RG (1990) Recombinant yeast TFIID, a general transcription factor, mediates activation by the gene-specific factor USF in a chromatin assembly assay. Proc Natl Acad Sci U S A 87:9153–9157 Chang LA, Smith T, Pognonec P, Roeder RG, Murialdo H (1992) Identification of USF as the ubiquitous murine factor that binds to and stimulates transcription from the immunoglobulin lambda 2-chain promoter. Nucleic Acids Res 20:287–293 Carter RS, Ordentlich P, Kadesch T (1997) Selective utilization of basic helix-loop-
In Vivo and In Vitro Tools to Identify and Study Transcriptional Regulation helix-leucine zipper proteins at the immunoglobulin heavy-chain enhancer. Mol Cell Biol 17:18–23 10. Trepicchio WL, Krontiris TG (1993) IGH minisatellite suppression of USF-bindingsite- and E mu-mediated transcriptional activation of the adenovirus major late promoter. Nucleic Acids Res 21:977–985 11. Peter M, Herskowitz I (1994) Joining the complex: cyclin-dependent kinase inhi bitory proteins and the cell cycle. Cell 79: 181–184 12. North S, Espanel X, Bantignies F et al (1999) Regulation of cdc2 gene expression by the upstream stimulatory factors (USFs). Oncogene 18:1945–1955 13. Cogswell JP, Godlevski MM, Bonham M, Bisi J, Babiss L (1995) Upstream stimulatory factor regulates expression of the cell cycledependent cyclin B1 gene promoter. Mol Cell Biol 15:2782–2790 14. Jung HS, Kim KS, Chung YJ et al (2007) USF inhibits cell proliferation through delay in G2/M phase in FRTL-5 cells. Endocr J 54:275–285 15. van Deursen D, Jansen H, Verhoeven AJ (2008) Glucose increases hepatic lipase expression in HepG2 liver cells through upregulation of upstream stimulatory factors 1 and 2. Diabetologia 51:2078–2087
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16. Nowak M, Helleboid-Chapman A, Jakel H et al (2008) Glucose regulates the expression of the apolipoprotein A5 gene. J Mol Biol 380:789–798 17. 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 18. 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 19. Corre S, Mekideche K, Adamski H, Mosser J, Watier E, Galibert MD (2006) In vivo and ex vivo UV-induced analysis of pigmentation gene expressions. J Invest Dermatol 126:916–918 20. Heid CA, Stevens J, Livak KJ, Williams PM (1996) Real time quantitative PCR. Genome Res 6:986–994 21. Colosimo A, Goncz KK, Holmes AR et al (2000) Transfer and expression of foreign genes in mammalian cells. Biotechniques 29:314–318, 320–2, 324 passim 22. de Wet JR, Wood KV, DeLuca M, Helinski DR, Subramani S (1987) Firefly luciferase gene: structure and expression in mammalian cells. Mol Cell Biol 7:725–737
Chapter 22 Measuring the Absolute Abundance of the Smad Transcription Factors Using Quantitative Immunoblotting David C. Clarke and Xuedong Liu Abstract In the age of systems biology, biologists seek to quantify the absolute number of molecules in experimentally treated samples. Immunoblotting remains a technique of choice for assessing the relative differences between the protein levels in different samples. Here we discuss how to exploit immunoblotting for estimating the number of Smad transcription factor molecules per cell. We focus on describing the calculations needed to analyze the data. Our methods are generally applicable to the quantification of other cellular proteins. Key words: Transforming Growth Factor-b, Smad, Cell signaling, Quantitative, Immunoblot, Western blot
1. Introduction In the age of systems biology, we seek to prioritize the importance of putative molecular mechanisms and to determine the way in which they are integrated to give rise to phenotype. These goals depend on our ability to quantify and model the system of interest, which calls for experimental methods that can accurately quantify the absolute number of biological molecules in a sample. For example, the quantitative properties of Transforming Growth Factor-b (TGF-b)/Smad signaling determine the specific cell response (1). Understanding TGF-b biology therefore requires investigating these properties, such as kinetic parameter values and the cellular concentrations of relevant molecular species. The principal intracellular components of the TGF-b signaling pathway are the Smad transcription factors (2). Eight Smad isoforms exist in mammalian cells, which transduce signals from the TGF-b
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superfamily member ligands. Five of the isoforms (1, 2, 3, 5, and 8) are classified as the “receptor-regulated” Smads (R-Smads); one of the isoforms, Smad4, is classified as the “common-mediator” Smad (Co-Smad); and the two remaining isoforms, 6 and 7, are classified as the “inhibitory” Smads (I-Smads). Smads 2, 3, and 4 are the principal downstream mediators of TGF-b signaling proper, and we focus on quantifying Smad2 in the following discussion. Ultimately, we hope to quantify the Smads in various cell states, as well as the amounts of Smads with different Posttranslational modifications, and their abundance in different cellular locations (see the chapter by Chapnick and Liu in this volume). In this chapter, we describe our methods for quantifying the number of Smad transcription factor molecules per cell using quantitative immunoblotting. Normally, immunoblotting cannot be used to derive quantitative information by direct comparison of band optical densities because of the nonlinear relationship between protein abundance and optical density. However, by running a standard curve of proteins of known abundance alongside the experimental samples, immunoblotting can be used to estimate Smad transcription factor abundance in absolute units (e.g., moles per Liter or molecules per cell). 1.1. Relevant Background Information on Immunoblotting
Immunoblotting, otherwise known as “Western blotting”, is a protein analysis technique that involves separating proteins using polyacrylamide gel electrophoresis, transferring them to a nitrocellulose or polyvinylidene difluoride (PVDF) membrane, and then exposing the membrane to antibodies that specifically bind the protein of interest. Usually, a second round of antibody incubation follows in which the secondary antibody linked to an enzyme, a fluorescent moiety, or an otherwise detectable molecule, targets the first antibody. The bands are then detected, typically using chemiluminescence, fluorescence, or chemifluorescence. The signals are captured on autoradiography film or digitally and the bands are quantified by densitometry. A key issue in immunoblotting is that the band intensity is nonlinearly related to the abundance of target protein in the sample. This is especially true for chemiluminescence detection using autoradiography film because the incident energy is related to the amount of energy passing through the film (the “emergent energy”) by Beer–Lambert’s law, such that the band intensity is a power law function of the protein abundance (3). People often speak of the “linear range of detection” in immunoblotting and optimizing one’s blotting conditions so that the band intensity of their sample falls in this range. In reality, there is no true linear range of detection – a linear range can be approximated, however,
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by employing blotting conditions that fall either on the steep initial rise of the power law curve or the more gradual rise that occurs more distally in the curve. Therefore, there are two options for quantifying immunoblot data using standard curves – (1) attempt to optimize the conditions to achieve an approximately linear standard curve that encompasses the sample signal intensity or (2) use a power law function to fit a broader standard curve. The first option is useful when you seek to quantify the band intensity of a single sample. For example, we discuss below how to use immunoblotting to quantify the abundance of endogenous Smad2 in the lysates of cells cultured in basal conditions (Fig. 1). The second option is likely better when you seek to quantify the band intensities of a number of samples in which the signal intensity is expected to vary over a broad dynamic range. We used this strategy to quantify the abundance of C-terminally phosphorylated Smad2 (phospho-Smad2) in cells exposed to a range of concentrations of TGF-b over time (Fig. 2). 1.2. Outline of a Quantitative Immunoblotting Experiment
Two sets of proteins are needed: (1) purified protein standards whose abundance or concentration is either already known or can be measured and (2) experimental samples for which the protein abundance is to be measured based on the standards. The stock concentration of the protein standards can be measured using gel-based methods such as Coomassie staining or the extinction coefficient method (4–6). Next, the specific dilutions to use for the immunoblot are determined by performing a series of immunoblots with different ranges of dilutions of the protein standard until a range of standards that encompasses the signal from the experimental samples is obtained. Once the conditions for the standard curve are determined, the immunoblot that includes both the standards and experimental samples is performed. Once a successful immunoblot is reproducibly obtained, the data is then analyzed to calculate the number of protein molecules contained in the experimental samples. In addition, the experimental design must include measurement or calculation of other variables, including the molecular weights of both the protein standard and of the target endogenous protein (these will often be different because the protein standard will likely be tagged) and the number of cells that comprise the experimental sample. The cell number can either be counted directly using the sample from which the proteins will be extracted or estimated by performing the experimental treatment in parallel on a separate group of cells that are subsequently counted. Deciding how to estimate cell number depends on whether the cells need to be rapidly processed at the end of the experiment.
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Fig. 1. Quantification of the abundance of endogenous Smad2 molecules per cell in PE25 cells. (a) Measurement of the protein standard concentration. The recombinant GST-Smad2 protein standard was quantified using Coomassie staining with BSA standards (0.03–0.1 mg/mL in increments of 0.01 mg/mL in a loading volume of 18 mL sample + 6 mL 4× SDS buffer). Three replicates of the GST-Smad2 protein sample were run alongside the BSA standards. (b) A linear standard curve was generated using Excel’s Trendline tool, from which the mass per band of each GST-Smad2 sample was interpolated. The mean mass per band was used to calculate the concentration of the stock GST-Smad2 protein standard. (c) Estimation of the number of Smad2 molecules per cell. Cells were seeded in 10 cm plates and grown to confluence,
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2. Materials 2.1. Cell Culture and TGF-b Treatment
1. Cell culture consumables: 6-well plates (Sarstedt), 0.05% trypsin-EDTA (GIBCO), Dubecco’s phosphate-buffered saline (D-PBS, GIBCO). 2. Cell culture medium: Dubecco’s modified Eagle’s Medium (DMEM) (GIBCO) supplemented with 100 IU/mL penicillin (ICN Biomedicals), 100 mg/mL streptomycin (Sigma), 2 mM l-glutamine (GIBCO), and 10% fetal bovine serum (SAFC Biosciences). 3. PE25 cells, a derivative of the mink lung epithelial cell line (Mv1Lu or CCL-64) (7, 8). 4. Reichert Bright-Line Hemocytometer. 5. Transforming growth factor-b, human recombinant isoform 1 (R & D Systems). Reconstitute in sterile 4 mM HCl containing 0.1% (w/v) fatty-acid-free bovine serum albumin (Sigma) and 75 mM NaCl (Sigma) to a final TGF-b1 concentration of no less than 1 mg/mL. Aliquot in single use amounts (e.g., 20–50 mL) in 0.7 mL microfuge tubes, snap freeze in N2, and store at –80°C.
2.2. Cell Lysis and Sample Preparation
1. Protein standards: We used recombinant glutathione-Stransferase (GST)-tagged Smad proteins for the quantification of Smad2 in whole-cell lysates (9). Phospho-Smad2 molecules in TGF-b-treated cells were quantified using recombinant phospho-MH2 domain phosphopeptide that was constructed by synthetic protein ligation. This polypeptide was a generous gift of Dr. Yigong Shi at Princeton University. General information on synthetic protein ligation is available in several excellent reviews (10, 11) while methods specific for the Smad MH2 domain are available in (12). 2. Cell lysis buffer: 50 mM Tris (Tris base, Research Products International) (pH 7.4), 400 mM NaCl, 1 mM ethylenediaminetetraacetic acid (EDTA, Fisher), 1% (v/v) NP-40 alternative (Calbiochem), 15% (v/v) glycerol (Sigma). The lysis buffer is kept at 4°C and lasts for months. Prior to use,
Fig. 1. (continued) trypsinized and counted using a hemocytometer, followed by lysis with 500 mL of lysis buffer. Serial dilutions of the GST-Smad2 protein were separated by SDS-PAGE alongside the six independent lysates from untreated PE25 cells. The 12%, 15-well SDS-polyacrylamide gel was loaded with 24 mL of each sample. Proteins were subsequently transferred, immunoblotted with a Smad2-specific antibody, and detected using enhanced chemiluminescence. (d) The GST-Smad2 dilutions conferred a reasonably linear standard curve, which was fit using Excel’s Trendline tool, and was then used to estimate the abundance of endogenous Smad2 molecules per band. These numbers were then divided by their respective number of cells, and the six estimates were used to calculate 95% confidence intervals for the number of Smad2 molecules per cell (we reported that 8.5–12 × 104 Smad2 molecules per cell exist in PE25 cells (14)).
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Fig. 2. Quantification of phospho-Smad2 levels in whole cell lysates of PE25 cells exposed to 10 pM TGF-b over 8 h. (a) Quantification of the phospho-MH2 standards using Coomassie-staining. Two replicates of the phospho-MH2 (P-MH2)
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supplement buffer with complete protease inhibitor cocktail (Roche) and 2 mM sodium fluoride (Fisher) and 1 mM sodium orthovanadate (Na3VO4, Sigma) phosphatase inhibitors. The Na3VO4 must first be activated in the following manner (13). First, prepare a 200 mM solution of Na3VO4. Adjust the pH to 10.0 using either 1 N NaOH or 1 N HCl (the starting pH of the Na3VO4 will depend on the source). At pH 10.0, the solution will be yellow. Boil the solution until it becomes colorless (approximately 10 min), followed by cooling it to room temperature, and then readjust its pH to 10.0. Repeat these steps until the solution remains colorless and the pH is stable at 10.0. Aliquot Na3VO4 and store at −20°C. 3. Cell lifter (Corning). 4. 1.7 mL microfuge tubes (Life Science Products). 5. Bovine albumin standard (2 mg/mL, Pierce). 6. BCA protein assay kit (Pierce). 7. 96-well microplate (Becton Dickinson). 8. Parafilm. 2.3. SDS-PAGE
1. Gel apparatus: For large gels, we used R. Shadel Inc. Discovery 2040 Vertical Gel Unit; for mini-gels, we used Invitrogen Novex XCell SureLock gel boxes with Bio-Rad glass rear spacer plates (1.0 mm spacer) and custom-made glass front plates. Note that the methods are compatible with many gel systems. 2. Polypropylene bags (VWR). 3. Heat Sealer: American International Electric Impulse Sealer.
Fig. 2. (continued) polypeptide were separated by SDS-PAGE alongside BSA standards (15 mL each of 0.03, 0.05, 0.06, 0.07, 0.09, 0.1, and 0.125 mg/mL serially diluted concentrations were loaded). The gel was then Coomassie-stained and quantified by densitometry. (b) A three-parameter logistic equation was used to fit the standard curve. Protein abundance per band of the phospho-MH2 peptides were numerically interpolated. The concentration of the standard was obtained by dividing the mass per band by the loading volume (15 mL). (c) Raw immunoblot data of a time series of lysates from cells treated with 10 pM TGF-b loaded alongside serial dilutions of cell lysate spiked with a known abundance of phosphoMH2 (P-MH2) polypeptide. An antibody specific to C-terminally phosphorylated Smad2 (P-Smad2) was used to probe the membrane. (d) Quantification of the immunoblot data. Phospho-MH2 band optical densities (OD) were quantified by densitometry and plotted versus the number of phospho-MH2 molecules per band. The data were fit to a power law equation in accordance with the chemiluminescence signal following the Beer-Lambert law (3). The fitted power law equation was used to calculate the estimated number of phospho-Smad2 molecules in the bands from the cell lysates. The number of molecules per cell was estimated by dividing the number of molecules per band by the estimated number of cells in the volume of lysate loaded on the gel. (1.5 × 106 cells were seeded in each well and, at the end of the experiment, were lysed in 200 mL of lysis buffer, giving a ratio of approximately 7.5 × 103 cells/mL of lysate.) Specific loading volumes for each sample were determined to achieve equal total protein abundance across samples. From this type of experiment, we derived estimates for phospho-Smad2 molecules per cell over time in response to 10 pM TGF-b (15).
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4. Protogel (30% acrylamide-0.8% bis-acrylamide) (National Diagnostics). Store at room temperature. Note that this solution is toxic and proper safety precautions should be taken. 5. 4× Resolving buffer: 1.5 M Tris (pH 8.8), 0.4% (w/v) sodium dodecyl sulfate (SDS, Sigma). Store at room temperature. 6. Stacking gel buffer: 1.0 M Tris (pH 6.8), 0.4% (w/v) SDS. Store at room temperature. 7. 10% ammonium persulfate (Sigma), made fresh in H2O. 8. N,N,N¢,N¢-Tetramethylethylenediamine (TEMED, Research Products International Corp.). 9. 4× SDS buffer: 200 mM Tris (pH 6.8), 40% (v/v) glycerol, 8% (w/v) SDS, 0.1% bromophenol blue (Sigma), and 400 mM b-mercaptoethanol (Sigma). Mix all components except b-mercaptoethanol, and store at room temperature (lasts for months). To make working reagent, aliquot 1.46 mL of solution into a 1.7 mL microfuge tube, and add 43 mL b-mercaptoethanol. Keep working reagent frozen at –20°C when not in use. 10. Electrophoresis buffer: 25 mM Tris, 192 mM glycine, 0.1% (w/v) SDS. Store at room temperature. 11. Protein marker: Spectra Multicolor Broad Range Protein Ladder (Fermentas). 2.4. Coomassie Staining
1. Destain solution: 10% (v/v) acetic acid (Mallinkrodt Chemicals), 45% (v/v) methanol, 45% (v/v) H2O. Store at room temperature. 2. Coomassie Brilliant Blue solution: 2.5 mg/mL Coomassie Brilliant Blue R-250 (Fisher) in destain solution. Filter with Whatman #1 filter paper. Store at room temperature.
2.5. Immunoblotting
1. Blotting membrane: Whatman Schleider Schuell BA83 Protran Nitrocellulose membrane. 2. Transfer buffer: 48 mM Tris, 39 mM glycine (Fisher), 0.37 g/ L SDS, 20% (v/v) methanol (Mallinckrodt Chemicals). Store at room temperature. 3. Semi-dry transfer unit: Hoefer TE 70 connected to a Hoefer Scientific Instruments PS500XT DC power supply. 4. Blotting containers (Research Products International Corp). 5. Filter paper: 0.8 mm thick (Life Science Products). 6. Ponceau S staining solution: 0.2% (w/v) Ponceau S (Sigma) in 1% acetic acid. Store at room temperature. 7. Tris-buffered saline-Tween (TBS-T): 0.05 M Tris, 0.138 M NaCl, 0.0027 M KCl, 0.05–0.1% Tween (Sigma). Store at room temperature.
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8. Blocking buffer: 4% (w/v) Safeway nonfat dried milk dissolved in TBS-T. Store at 4°C. 9. Primary antibodies: Smad2 (Zymed 51-1300), diluted to 1:1,000 in blocking buffer; phospho-Smad2 (Cell Signaling Technology #3108), diluted to 1:150 in blocking buffer. 10. Secondary antibody: Amersham Biosciences ECL Rabbit IgG, HRP-Linked Whole Ab from donkey. 11. Autoradiography cassette (Fisher Biotech). 12. Super-sensitive enhanced chemiluminescence substrate (Pierce SuperSignal West Dura Extended Duration Substrate). 13. Autoradiography film (ISC Bioexpress Blue Lite Autoradio graphy film F-9024-8×10).
3. Methods 3.1. Cell Culture and TGF-b Treatment
1. PE25 cells are cultured at 37°C/5% CO2 in cell culture medium in a humidified incubator. 2. Seed 1.5 × 106 PE25 cells per well in 6-well plates. Let the cells settle for at least 6 h or overnight. 3. If cells cultured under basal conditions (i.e., no TGF-b treatment) are to be analyzed, then the cell number in each well can be counted prior to lysis. Trypsinize the cells, collect them in a 15 mL conical tube, and store on ice. Dilute the cells with cell culture medium to a volume that confers approximately 50–75 cells per 1 mm2 (i.e., the area of one of the nine smaller squares in the hemocytometer counting area). Count the cells within at least five squares, calculate the average, divide the average by the volume above each square, and multiply by the dilution factor if necessary. Spin down the remaining cells at 300 × g for 5 min and proceed with lysis. 4. For experiments involving TGF-b stimulation, add the desired amount of TGF-b to sufficient culture medium to treat all the samples in the experiment. Warm the medium to 37°C in a water bath. Aspirate the old media, apply the medium containing the TGF-b to the cells, and incubate the cells for the desired time. 5. At the end of the experiment, aspirate the medium, wash the cells with 2 mL ice-cold D-PBS, aspirate D-PBS, and snap freeze the cells by carefully pouring N2 into each well. Store plates at –80°C until ready to lyse.
3.2. Cell Lysis and Sample Preparation
1. Thaw the frozen 6-well plates at 4°C for a few minutes to warm up.
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2. Pipette 200 mL of lysis buffer onto the cells and scrape them using the cell lifter. Pipette the cells into a microfuge tube. Save an aliquot of lysis buffer to serve as a background control in the BCA assay. 3. Rotate the cells for 45 min at 4°C, followed by spinning down the insoluble material in a microfuge for 10 min at 16,100 × g at 4°C. Transfer lysate into a fresh microfuge tube and either store at −80°C for later use or proceed directly to the following steps, keeping tubes on ice. Avoid repeated freeze-thaw cycles, because this can reduce the signal on subsequent immunoblots. 4. Measure the protein concentration of the lysates using the microplate version of the bicinchoninic acid (BCA) method following the manufacturer’s instructions (see Note 1). 5. Combine the volume of lysate necessary for the desired amount of total protein loaded per sample (we typically load 30–36 mg of protein per well), dilute to the desired volume with H2O, followed by adding 4× SDS buffer to the final volume (we typically load each well with 38 mL, such that 27.75 mL of diluted sample is combined with 9.25 mL of 4× SDS buffer). Cap the samples tightly and boil them in a heatblock at 90°C for 5 min, let them cool at room temperature, followed by centrifuging the samples at 3,300 × g for 3 min in a microfuge at room temperature (see Note 2). 6. Use samples for SDS-PAGE or store at –80°C for future use. 3.3. SDS-PAGE
1. Determine the percentage acrylamide, the size, and the number of wells needed for the gel. For quantification of the protein standards, we run mini-gels with the BSA standards and 2–3 replicates of the protein standard (GST-Smad2 or phospho-MH2 polypeptide) (14, 15). For quantification of the Smad2 transcription factor under basal conditions, we use 15-well minigels loaded with the molecular weight marker, 6 standards, and 6 independent cell lysates (14). For quantification of phospho-Smad2 molecules per cell during TGF-b signaling, we run a large 20-well gel (15). For mini-gels, follow the manufacturer’s instructions. We provide instructions for making and running large gels below. 2. Wash the glass plates and combs with dish soap and water, followed by wiping down with a Kimwipe and glass cleaning solution or ethanol. 3. Prepare gel-casting bags by heat-sealing the polypropylene bags using a heat sealer. The bag dimensions should just be big enough to snugly fit the front and rear glass plates and spacers for two gels. Check for leaks by filling with water and looking for drips. Empty the bag.
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4. Very carefully, insert the glass plates for two gels into the gel-casting bag. In particular, be careful not to cause any small tears in the bag, because this will cause leaks upon pouring the gels (see Note 3). Insert two spacers in between each set of front and rear glass plates at their lateral edges. 5. Using large office supply binder clamps, sandwich an additional glass plate to the front and back of the gel-casting bag. 6. Insert one of the combs, measure approximately 1 cm below the teeth, and mark this spot on the glass plate used to sandwich the gel plates. This represents the line to which the running gel will be poured. Remove the comb. 7. In a 125 mL filtering flask, combine 21 mL H2O, 15.6 mL resolving gel buffer, 25 mL Protogel, and 625 mL 10% APS. Degas gel for about 1 min by capping flask with a rubber stopper and attaching to a vacuum pump. Add 23 mL TEMED to solution, swirl gently, and immediately pour the gel (see Note 4). 8. Pipette 500 mL butanol slowly and evenly onto gel. Avoid disturbing the gel otherwise during polymerization (~15– 30 min). Let the gel solution remaining in the flask polymerize. When that gel is polymerized, so should be the poured gel. Another sign is the presence of excluded H2O in a separate layer under the butanol at the top of the polymerized gel. 9. Soak up the butanol with filter paper. Remove the polymerized gel from the filter flask and wash. Rinse the flask with distilled H2O, and dry it with paper towels. 10. Prepare the stacking gel by combining 18.75 mL H2O, 7.5 mL stacking gel buffer, 4.5 mL protogel, and 300 mL 10% APS. Degas the solution for about 1 min. Add 23 mL TEMED, and pour the gel up to the top of the glass plates. Insert combs, ensuring that no bubble gets between the teeth (small bubbles below the teeth are acceptable). Let the gel polymerize for 15–30 min, and use the gel immediately or store in electrophoresis buffer at 4°C for up to 3 weeks. 11. Remove the combs carefully by clamping the glass plates at the bottom with an office supply binder clamp and gently and evenly pulling out the comb. The vacuum created by removing the combs will distort the well walls. Use a narrow strip of 1 mm thick Teflon to straighten the well walls. 12. Set up the gel apparatus, and fill the upper and lower reservoirs with electrophoresis buffer. Wash out the gel wells with electrophoresis buffer using a 5 mL syringe fitted with a 20 g1 needle.
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13. Load gels using pipettor fitted with gel-loading pipette tips (see Note 5). Load 3–5 mL of the molecular weight marker, and load any empty wells with the same volume of 4× SDS buffer that is included in each sample. 14. Attach the gel apparatus to the power supply, and run on constant current mode at ~35 mA per gel. If desired, the gel can be run at lower currents to prolong the run time (e.g., overnight). 15. Run gel until the dye line has passed 5.5 cm from the border of the stacking and resolving gels. This is to ensure that all the proteins from the sample will be transferred onto a membrane that fits in the blotting containers. 3.4. Coomassie Staining
1. Carefully remove the resolving gel from the plates by cutting the stacking gel with a gel tool and by gently lifting the gel off the plates and placing it into the stain solution. Incubate the gel at room temperature for 1 h on a rocking shaker at a low speed (e.g., 3 out of 10). 2. Destain the gel by replacing the stain solution with destain solution. Replace the destain solution periodically (e.g., every 10–30 min) until the unbound stain has exited the gel and the stained bands are left behind (Figs. 1a and 2a). Document the gel, and perform densitometry (see Subheading 3.7).
3.5. Immunoblotting
1. Cut a piece of nitrocellulose membrane and four pieces of 0.8 mm thick filter paper to dimensions that match the intended size of the section of gel to be transferred. Cut a small notch in the upper left-hand corner to orient the membrane. Wet nitrocellulose membrane with H2O, and store in transfer buffer until use. Keep filter paper dry until just prior to use. 2. When the gel is finished running, lay the plates flat and gently remove the front plate using a gel tool. Cut off the stacking gel, and cut the unused portion of the resolving gel such that the dimensions of the resolving gel containing the separated proteins are the same as those of the membrane and filter paper. Equilibrate the gel in transfer buffer for about 5 min. 3. Load and run the transfer unit according to manufacturer’s instructions. Briefly, create a sandwich (in order from bottom to top) of two 0.8 mm thick transfer-buffer-soaked pieces of filter paper, the wetted nitrocellulose membrane, the gel, and two more pieces of soaked filter paper. As each layer is added, gently roll a test tube along the layer to press out any bubbles. 4. Upon completion of the transfer, check its effectiveness by soaking the membrane in Ponceau S solution for a few seconds, and then rinse with H2O until the bands are clearly visible.
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5. Perform the following steps at room temperature on a rocking shaker set at low speed. Block the membrane by incubating it for 1 h in ~10 mL blocking buffer. 6. Replace the blocking buffer with primary antibody, and incubate the membrane for ~2 h (see Note 6). Afterwards, collect primary antibody back into its tube and refreeze. Wash the membrane 3 × 5 min in TBS-T (see Note 7). 7. Incubate the blot in secondary antibody (5–6 mL for a small membrane and 8 mL for a large membrane) for 45–60 min (see Note 8) followed by washing the membrane 3 × 10 min in TBS-T. Discard the used secondary antibody. 8. Detect the blot by enhanced chemiluminescence (see Note 9) and visualize using autoradiography film. Perform exposures of several durations (typically between 30 s and 5 min) to optimize the signal relative to the background (e.g., Fig. 1c and Fig. 2c). Afterwards, align the developed film with the membrane, and mark the location of the molecular weight markers. Dry the membrane, label, and store (see Note 10). 3.6. Densitometry Using ImageJ
1. For Coomassie-stained gels, use a gel documentation system (e.g., AlphaImager, AlphaInnotech Corp.) to obtain a digital image of the gel. For immunoblots, scan the film using a digital scanner (e.g., we use a UMAX PowerLook III with MagicScan v4.5 software) in transmissive mode, grayscale, and at a resolution of 300–600 dots per inch. Save as TIFF files. 2. Perform densitometry using ImageJ (Scion Corp.). Open the file containing the gel image, and ensure that the corresponding bands in each lane are as horizontal as possible by rotating the image (see Note 11). 3. Define boundaries for each band by tracing a vertical line and pressing “Backspace” in the space between each band and at the outer edges of the first and last bands. This deletes pixel density between the bands, which demarcates the bands in subsequent steps. 4. Trace a box around the entire set of bands and press
. If bands from different molecular weights must be quantified (which is typically the case if tagged recombinant proteins or protein fragments are used as standards), then move the box around those bands and press . Press to display the plotted optical densities (see Note 12). 5. In the plotted optical densities window, trace a horizontal line across the bands at the level of the optical densities at the outer edges of the gel. This removes the background optical density from each lane.
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6. Quantify each band density using the magic wand tool by clicking within the space defined by each band and the background line. A yellow tracing should appear with each click showing that the band has been quantified. The results window will appear automatically with the densitometric measurements (arbitrary units). 7. Select “Copy All” from the Edit menu, and paste the results into a spreadsheet. 3.7. Data Analysis and Calculations for Coomassie-Stained Gels Using Microsoft Excel
1. Plot the mass of BSA in each lane versus the corresponding OD. Examine the shape of the curve – if the curve appears reasonably linear, go to step 2 (Fig. 1a). If the standard curve resembles an S-shape (Fig. 2a), a three-parameter logistic equation (or Hill equation) will be required, and its use is described in steps 4–9 of this subheading. 2. In the case of a linear standard curve, use Excel’s Add Trendline feature by right-clicking on the data points within the plot. Select “Add Trendline,” and then select the “Linear” type. Click the “Options” tab, and check the “Display Equation on Chart” option and the “Display the R2-value on chart” if desired (as one way to evaluate the goodness of fit). The equation will be of the form y = mx + b, where y = OD, m = slope of the line, x = mass of BSA per lane, and b = the y-intercept (Fig. 1b). 3. Interpolate the mass of protein per lane for the protein standards by solving the equation with respect to x, substituting the OD values of the protein standards into the equation and calculating the mass of protein per lane. Compute the mean and standard deviation if multiple replicates of the protein standard were measured. 4. If an S-shaped curve results, then the following equation can be used to fit the data: ODmax· (protein abundance )
n
OD =
(
)
K n + protein abundancen
where OD is the optical density of the band, ODmax is the asymptotic maximum OD, K is the protein abundance that confers the 50% of ODmax, and n is a parameter that determines the steepness of the S-curve. This equation is most easily solved using a numerical approach implemented using a spreadsheet. 5. First, generate the standard curve equation by estimating the values for the parameters ODmax, K, and n. This equation will then be used to estimate protein abundances that correspond to the experimentally observed OD values. Start by entering
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the model parameters in three cells just below the data columns. Enter guess values for ODmax, K, and n. Sensible guess values include the maximum observed OD value for ODmax, the mass of BSA that approximately corresponds to the OD that is half of ODmax for K, and between 0.5 and 2 for n. 6. In the column next to the data columns, enter the model equation for each BSA mass using the parameters entered above (use absolute cell references or cell names). 7. In the following column, calculate the “error” or “residuals” by subtracting the modeled OD value by the observed OD value. At the bottom of this column, use the SUMSQ function applied to residuals to calculate the sum of squared errors (SSE). 8. Use Excel’s Solver tool to find the parameter values that best fit the data. From the menu bar, select Tools, and from the dropdown menu, select Solver (see Note 13). In the Solver dialog box, set the “Target Cell” as the cell containing the SSE value, choose the “Min” radio button for the “Equal to” option, enter the cells containing the model parameters in the “By Changing Cells” box, and press “Solve” (see Note 14). The SSE value should change to a smaller value, the model parameter values should change, and, in the standard curve plot, the model line should run through the data points (Fig. 2b). The Solver step should be repeated several times to ensure convergence (i.e., the solution ceases to change). 9. Use the standard curve to interpolate the protein masses per band for the protein standards. In the column adjacent to the OD values corresponding to the experimental samples, enter guess values for the protein abundances of the protein standards using the standard curve data as a guide. 10. In the adjacent column, write the model equation with the fitted parameters to calculate an OD value corresponding to the guess value. 11. Subtract the calculated OD value from the observed OD value (the residual) and square it. 12. For each protein standard replicate, use Excel’s Solver tool to minimize the squared error value by changing the guess value for protein abundance. Repeat the Solver step until convergence is reached (i.e., until the calculated OD and observed OD values are equal through at least three decimal places, which should correspond to a very low squared error value). Calculate the mean and standard deviation of the protein standard masses if more than one replicate was run on the gel.
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13. Calculate the concentration of the protein standard by dividing the mass by the volume of protein standard loaded on the gel. 3.8. Data Analysis and Calculations for Immunoblot Data Using Microsoft Excel
1. Estimate protein molecular weights using the ExPASy “Compute pI/MW” tool (http://www.expasy.ch/tools/ pi_tool.html), entering the primary sequence of the proteins as the input. Using this method, we found that Smad2 has a predicted molecular weight (MW) of 52,306 Da, GST-Smad2 has a predicted MW of ~78,306 Da, and phospho-MH2 polypeptide has a predicted MW of 25,517 Da. 2. Create the standard curve by fitting an appropriate function to the data. First, express the standard curve in the correct units. Because antibody staining is proportional to the number of epitopes present in the sample, express the concentration of the protein standard in units of moles per microliter by dividing the concentration (in units of g/mL) by the molecular weight of the protein standard. 3. Calculate the protein abundance per band (in units of mol) of the standards by multiplying the protein standard concentrations by the volume of standard loaded. (If desired, the protein abundance can also be expressed as the number of molecules per band simply by multiplying by Avogadro’s number, instead of performing the same operation in step 9.) 4. Plot the known protein abundances versus the OD values. The shape of the curve will likely be either reasonably linear (Fig. 22.1d) or resemble the power law form (Fig. 2d). 5. If the curve is linear, use the same procedure outlined in Subheading 3.7 steps 2–3 to generate the standard curve and to interpolate the protein abundances per band from the experimental samples. 6. If the curve resembles a power-law function, enter the following equation adjacent to the observed OD values:
OD = C*(Protein abundance)n
where C and n are the model parameters and Protein abundance is the abundance of protein per band calculated in step 3. Choose initial guess values for C and n. Plot the calculated values alongside the experimental data, and manually adjust the parameters until the model curve is reasonably close to the data points. A reasonable initial guess value for n should be between 0.3 and 0.7, as we and others (3) typically observe fitted values of n from this range. Choose C such that the plot of the model curve scales well with the data. 7. Follow the procedure described in Subheading 3.7 steps 4–8 to fit the C and n parameters to the data. Briefly, calculate the
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residuals for the modeled and observed OD values, calculate the SSE, and minimize the SSE by using Excel’s Solver tool to change the values of C and n until the minimum SSE is found (Fig. 2d). 8. Use the equation with fitted parameter values to estimate the protein abundances in the experimental samples from their corresponding optical densities. The solution to the power law equation with respect to Protein abundance is
Protein abundance = (OD/C)(1/n).
9. Convert the protein abundance per band into molecules of protein per band by multiplying Protein abundance by Avogadro’s number (6.02 × 1023 molecules/mol) (if not already done in step 3 for the standard). 10. Calculate the cell concentration: Cell concentration = (Number of cells in lysate)/(Lysate volume) For example, we estimated 1.5 × 106 cells in the lysates of the phospho-Smad2 kinetics experiment (Fig. 2), and each well was lysed with 200 mL of lysis buffer. Therefore, we estimate a cell concentration of 7.5 × 103 cells/mL. The result is then multiplied by the volume of lysate loaded to estimate the number of cells that contributed to the signal of the corresponding band in the immunoblot. 11. Calculate the number of molecules per cell by dividing the number of molecules per band by the number of cells.
4. Notes 1. To bring the protein concentration of the lysate within the range of the standard curve, we dilute the lysate 10× in water, e.g., 3 mL of lysate + 27 mL of H2O, followed by loading 10 mL of the diluted lysate and combining with 200 mL of the BCA working reagent in two separate replicates. Cover the microplate with parafilm, and incubate at 37°C for 30 min. It is also possible to use the Bradford method, but our lysis buffer reacts with the Bradford reagent, resulting in high background, and the standard curve is not as linear as that for the BCA assay. 2. Loading the optimal (not necessarily the most) amount of sample is imperative for achieving clear separation between the bands in adjacent lanes. Our experience has been that gels loaded with too much protein tend to cause the adjacent bands to overlap.
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3. We line up the glass plates upside down and slowly slide the gel-casting bag down over the plates, taking care to avoid cutting the bag with the leading corners of the plates. 4. Pour the resolving gel at a consistent rate until the gel line approaches the marked line. Let it settle for a few seconds, and then pour slowly until the gel line matches up with the marked line. Watch gel for leaks. Minor slow leaks can be replenished as needed until leaks cease as the gel polymerizes. Note that the use of properly poured, relatively fresh gels is crucial for obtaining sharp, straight bands. Distorted or smeared bands usually result from imperfections in the gel and are unusable for quantification purposes. 5. Insert tip as far into well as possible and slowly pipette sample into well until the sample lies above the tip orifice. Continue pipetting while slowly drawing tip upward to the first stop of the pipette. Purge remaining sample with tip orifice well above the sample (to avoid blowing out sample from well). 6. Using concentrated primary antibody exceeding the usual recommended dilution is typically needed for endogenous protein detection. Our antibodies are often diluted to 1:150– 1:250. We also emphasize the need to have sufficient volume of antibody solutions to maximize the contact time of the antibody solution with the entire surface of the membrane. For mini-gels (5 × 8 cm), we typically use >6 mL of antibody solution, and for large gels (14 × 5.5 cm), we use 10–15 mL of antibody solution. While this might seem exhorbitant, we save reagent and costs by reusing the antibody stock multiple times and freezing between uses at –20°C (we often use the same antibody stock ten or more times this way). In addition, we have observed that antibody incubations performed at room temperature for as long as feasibly possible (we typically do 2 h incubations) confer better signal than incubations performed overnight at 4°C. 7. We shorten the wash times from the manufacturer’s recommendation and adjust the Tween concentration in the Trisbuffered saline-Tween (TBS-T) wash solution to achieve a blot with good signal and low background. Typical wash times are 3 × 5 min after the primary antibody and 3 × 10 min after the secondary antibody incubation, and the Tween concentration is 0.05% (v/v) and increased to 0.1% if there is too much background. 8. We keep the secondary antibody incubation to less than 1 h. Intuitively, one might think that prolonged secondary antibody incubations would confer stronger signal. This is not true, because in standard immunoblotting technique, the
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secondary antibody solution contains no primary antibody. Since antibody binding is reversible, primary antibodies bound to the membrane will experience a driving force to dissociate according to the laws of equilibrium, thus reducing the overall signal if left too long. 9. To detect endogenous proteins (which are often expressed at levels that are difficult to detect), we use Thermo Scientific (formerly Pierce) SuperSignal West Dura Extended Duration Substrate (34076). This chemiluminescent substrate delivers a much stronger and longer-lasting signal than standard chemiluminescent reagents. 10. Nitrocellulose (and presumably PVDF) membranes can be readily dried and rewetted at a later time for subsequent use with no loss of signal. Therefore, whenever a membrane is not in use, we always dry and store it in a plastic sheet protector in a binder. One caveat is that membranes become brittle over time, and great care is needed to avoid tearing the membrane until rewetted. 11. We do not use ImageJ’s “Subtract background” function, because the optical densities of the bands can be unevenly affected. Instead, we process the background as part of our quantification procedure. 12. The box should be just large enough to encompass all the bands, with some space at the outer edges to measure the background levels. If multiple ranges of bands are to be quantified, the same box must be used to ensure compatibility between sampled ranges. Therefore, the box dimensions must be sufficiently large to incorporate the bands with the largest area and all the bands within each range. 13. If the Solver option does not appear, it may need to be installed. 14. This step should be repeated several times until convergence is reached. In addition, different initial guesses for the parameter values should also be tried to enhance the probability of finding the best-fit parameter values.
Acknowledgments We thank Scott Dixon, Dana Ungermannova, and Meredith Brown for critical reading of the manuscript. This work was supported by a National Institutes of Health grant (GM083172) to X. L.
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References 1. Clarke DC, Liu X (2008) Decoding the quantitative nature of TGF-b signaling. Trends Cell Biol 18:430–442 2. Massague J, Seoane J, Wotton D (2005) Smad transcription factors. Genes Dev 19: 2783–2810 3. Pitre A, Pan Y, Pruett S, Skalli O (2007) On the use of ratio standard curves to accurately quantitate relative changes in protein levels by Western blot. Anal Biochem 361: 305–307 4. Pace CN, Vajdos F, Fee L, Grimsley G, Gray T (1995) How to measure and predict the molar absorption coefficient of a protein. Protein Sci 4:2411–2423 5. Gill SC, von Hippel PH (1989) Calculation of protein extinction coefficients from amino acid sequence data. Anal Biochem 182: 319–326 6. Pierce. Extinction coefficients. Rockford, IL; 2006 7. Hua X, Liu X, Ansari DO, Lodish HF (1998) Synergistic cooperation of TFE3 and smad proteins in TGF-beta-induced transcription of the plasminogen activator inhibitor-1 gene. Genes Dev 12:3084–3095 8. Macdonald M, Wan Y, Wang W et al (2004) Control of cell cycle-dependent degradation
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of c-Ski proto-oncoprotein by Cdc34. Oncogene 23:5643–5653 Zhang Y, Feng X, We R, Derynck R (1996) Receptor-associated Mad homologues synergize as effectors of the TGF-beta response. Nature 383:168–172 Muralidharan V, Muir TW (2006) Protein ligation: an enabling technology for the biophysical analysis of proteins. Nat Methods 3:429–438 Muir TW (2003) Semisynthesis of proteins by expressed protein ligation. Annu Rev Biochem 72:249–289 Wu JW, Hu M, Chai J et al (2001) Crystal structure of a phosphorylated Smad2. Recognition of phosphoserine by the MH2 domain and insights on Smad function in TGF-beta signaling. Mol Cell 8:1277–1289 Gordon JA (1991) Use of vanadate as protein-phosphotyrosine phosphatase inhibitor. Methods Enzymol 201:477–482 Clarke DC, Betterton MB, Liu X (2006) Systems theory of Smad signalling. Systems Biology (Stevenage) 153:412–424 Clarke D, Brown M, Erickson R, Shi Y, Liu X (2009) Transforming growth factor-beta depletion is the primary determinant of Smad signaling kinetics. Mol Cell Biol 29:2443–2455
Chapter 23 Flow Cytometry Analysis of Transcription Factors in T Lymphocytes Diana I. Albu, Danielle Califano, and Dorina Avram Abstract Detection of transcription factors in immune cell populations, particularly in subpopulations that are represented at low frequencies in lymphoid and nonlymphoid organs, presents a particular challenge when using traditional methods such as western blot analysis. Therefore, development of flow cytometrybased methods which allow identification of transcription factors in specific immune cell populations is of main interest. Here we developed and optimized a methodology for rapid and convenient detection of the transcription factor BCL11B in T lymphocyte subpopulations using flow cytometry. The optimal protocol employs saponin and Tween 20 both during the fixation and permeabilization steps, and we demonstrate that it is efficient for three anti-BCL11B antibodies covering distinctive BCL11B epitopes. In addition, we prove that the method preserves the staining of surface markers. Key words: Transcription factors, BCL11B/CTIP2, T lymphocytes, CD4+ T lymphocytes, CD8+ T lymphocytes, Flow cytometry
1. Introduction Western blot analysis of total cell lysates or nuclear fraction is most commonly employed for transcription factors detection in various types of cells. Different from most cell lines, primary cells are composed of heterogeneous populations. Therefore, analysis of transcription factors by western blot analysis has major limitations because it requires prior purification of subpopulations of cells, and a large number of cells (at least 0.5 × 106). For cells of the immune system such separation of subpopulations can be achieved by fluorescent-activated cell sorting (FACS), based on surface markers. However, many subpopulations contain limited numbers of cells, which makes western blot analysis difficult. On the other hand, immunocytochemistry offers the advantage of detection of transcription factors in small populations of cells Paul J. Higgins (ed.), Transcription Factors: Methods and Protocols, Methods in Molecular Biology, vol. 647, DOI 10.1007/978-1-60761-738-9_23, © Springer Science+Business Media, LLC 2010
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and also permits to determine whether the transcription factor is present in the cytoplasmic or nuclear fraction. This technique also allows surface staining, and therefore identification of subpopulations, however with limited number of fluorochromes at the same time, and limitations in terms of evaluation of significant number of cells. The method of choice that permits rapid and simultaneous identification of subpopulations expressing the transcription factor of interest in small but significant number of cells, without prior purification, is flow cytometry. The identification of subpopulations which express specific transcription factors can be achieved by knocking in reporter genes, such as the Green Fluorescence Protein (GFP) downstream of the transcription factor promoter, as it has been achieved for Foxp3 (1) and RORgammat (2). In the absence of such models, the populations expressing specific transcription factors can be identified by surface staining for subpopulation specific markers, and intracellular staining with fluorescence-labeled antibodies for specific transcription factors. This second goal requires permeabilization of the cellular and nuclear membranes to allow antibodies to reach the nuclear epitopes. Specifically, following surface staining, fluorochrome-labeled antibodies bound to surface markers are fixed, following which cells are permeabilized, allowing exposure of nuclear epitopes. Next, transcription factors are identified either directly by staining with fluorochrome-labeled transcription factor-specific antibodies, or indirectly, with fluorochromelabeled secondary antibodies. At the end, samples are run on a flow cytometer and specific subpopulations are identified based on the surface markers together with the transcription factor of interest. This technique offers the advantage of single cell analysis, which enables the determination of both the presence of the transcription factor of interest in subpopulations of primary cells, and the frequencies of the primary cell subpopulations which express the transcription factor, without the need of the prior purification of the subpopulations. The technique was proven to render significant data after acquisition and analysis of a small number of cells, as little as 3 × 104. In addition, the technique allows the expression quantification of specific transcription factors in subpopulations of cells by evaluating the mean fluorescence intensity (MFI). The transcription factor BCL11B, known also as CTIP2, is a sequence-specific DNA binding protein (3) expressed in brain (4–8), skin (9, 10), and immune system, where its presence is confined to T lymphocytes (11–14). We previously demonstrated that ectopic expression of the BCL11B homolog BCL11A/CTIP1 in HEK293 cells resulted in restrictive localization of the protein to the nuclei (4). Similarly, endogenous BCL11B was restricted to the nuclei in CD4+ T lymphocytes,
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Fig. 1. BCL11B transcription factor is localized in the nucleus of CD4+ T lymphocytes. (a) Prior to immunocytochemistry and western blot analysis the CD4+ T lymphocytes were isolated from spleen and lymph nodes by MACS technology, using anti-CD4 antibody-coated magnetic microbeads and magnetic column separation, as described in Material and Methods. The degree of purification was evaluated by staining an aliquot of the purified CD4+ T lymphocytes with anti-CD4-FITC-labeled antibodies and analyzed by flow cytometry. Purity of the separated CD4+ T lymphocytes was 92.4%. (b) Left: immunocytochemistry staining of cells cytospun on coverslips with anti-BCL11B antibodies (Clone BL1801) or prebleed rabbit serum, followed by FiTC-conjugated secondary antibodies, as described in Material and Methods. Middle: Hoechst counterstained of the cells shown in the left; right: Overlay. Images were obtained on an Olympus BX61 Microscope with 100× objectives. (c) Left: Western blot analysis of cytoplasmic and nuclear extracts from CD4+ T lymphocytes purified as described in a. Right: Western blot analysis of nuclear extracts from 0.5 × 106 (left lane) or 1.5 × 106 CD4+ T lymphocytes (right lane). Actin and HDAC2 were used as loading controls.
as demonstrated both by immunocytochemistry and western blot analysis of cytoplasmic and nuclear fractions (Fig. 1). However, as shown above, these two techniques have limitations for detection of transcription factors in T cell subpopulations, and other cells of the immune system. It is therefore critical to establish a methodology based on flow cytometry analysis that allows determination of various T cell subpopulations which express BCL11B.
2. Materials 2.1. Mice
5–10 weeks C57BL/6 female or male mice were housed in the Albany Medical Center animal research facility and all the animal procedures were approved by the Institutional Animal Care and Use Committee.
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2.2. Instruments and Disposables
1. Sterile surgical instruments: forceps and scissors (Roboz Surgical Instrument Co., Gaithersburg, MD). 2. 40 mm nylon cell strainers (BD Falcon, Franklin Lakes, NJ). 3. 50 ml conical tube (BD Biosciences, Franklin Lakes, NJ). 4. 5 ml syringes (BD Biosciences, Franklin Lakes, NJ). 5. V-bottomed 96 well plates (Corning Costar Corp, Cambridge, MA). 6. FACS tubes (Krackeler Scientific, Albany, NY). 7. Sterile transfer pipettes (Fisher Scientific, Pittsburgh, PA). 8. MACS multistand and separation unit, MS columns and mouse CD4 L3T4 Microbeads (Miltenyi Biotec, Bergisch Gladbach, Germany). 9. Sorval Legend RT with cytospin system and plate holders (Thermo Scientific, Waltham, MA). 10. Olympus BX61 Microscope (Olympus, Center Valley, PA). 11. FACS Calibur Flow Cytometer (Beckton Dickinson, San Jose, CA) upgraded to five colors (Cytek, Fremont, CA).
2.3. Antibodies and Reagents
1. Supernatants of HB-197 hybridoma (ATCC, Rockville, MD) containing 2.4G2. 2. Anti-CD4 antibody, clone GK1.5, (FITC) (eBioscience, San Diego, CA). 3. Anti-CD8 antibody, clone 53-6.7, (PE-Cy7) (eBioscience, San Diego, CA). 4. Anti-BCL11B antibodies, clones BL1801, BL1800, BL1798, and BL1799 (Bethyl laboratories, Montgomery, TX) (Note 1). 5. Anti-HDAC2 antibody, clone H54 (Santa Cruz Biotech nology, Santa Cruz, CA). 6. Anti-b-actin mouse monoclonal antibody (Sigma, St. Louis, MO). 7. Alexa 647-conjugated goat antirabbit secondary antibodies (Invitrogen Corp., Carlsbad, CA). 8. Fluorescein isothiocyanate (FITC)-conjugated goat antirabbit secondary antibodies (Southern Biotechnology Associates, Inc., Birmingham, AL). 9. Formaldehyde 37% (VWR, West Chester, PA). 10. Saponin (Sigma, St. Louis, MO). 11. Tween 20 (Fisher Scientific, Pittsburgh, PA). 12. RNase (Sigma, St. Louis, MO). 13. Fluoromount (Southern Biotechnology Associates, Inc.). 14. Protease inhibitor cocktail (Sigma, St. Louis, MO). 15. HEPES (Fisher Scientific, Pittsburgh, PA).
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16. Heat-inactivated fetal bovine serum FBS (Hyclone, Logan, UT). 17. Penicillin/Streptomycin solution (Gibco/BRL, Bethesda, MD). 18. l-glutamine (Gibco/BRL, Bethesda, MD). 19. Beta-mercaptoethanol (Sigma, St. Louis, MO). 20. NH4Cl (Sigma, St. Louis, MO). 21. KHCO3 (Sigma, St. Louis, MO). 22. EDTA (Sigma, St. Louis, MO). 23. Bovine serum albumin (BSA) (Sigma, St. Louis, MO). 24. Sodium azide (Sigma, St. Louis, MO). 25. NaCl (Sigma, St. Louis, MO). 26. KCl (Sigma, St. Louis, MO). 27. Sodium Phosphate dibasic (Sigma, St. Louis, MO). 28. Potassium Phosphate monobasic (Sigma, St. Louis, MO). 29. Hoechst 33258 (Molecular Probes, Eugene, OR). 2.4. M edia and Buffers
1. Complete Medium (CM): RPMI 1640 media (Gibco/BRL, Bethesda, MD) supplemented with 10% heat-inactivated FBS, 100 U/ml Penicillin, 100 U/ml Streptomycin solution, 2 mM l-glutamine, 0.05 mM beta-mercaptoethanol and 25 mM HEPES. 2. Phosphate Buffer Saline (PBS): 137 mM NaCl, 2.7 mM KCl, 10 mM Sodium Phosphate dibasic, 2 mM Potassium Phosphate monobasic, pH 7.4 3. Red Blood Cell Lysis Buffer (RBCLB): 0.83% NH4Cl, 0.1% KHCO3, and 0.0037 % EDTA. 4. Staining buffer (SB): 1% BSA and 0.01% sodium azide in PBS. 5. Fixation buffers for flow cytometry (Note 2): (a) 3% formaldehyde in PBS (protocols 1–3) (Fig. 2). (b) 3% formaldehyde, 0.1% saponin in PBS (Note 3) (protocol 4) (Fig. 2). (c) 3% formaldehyde, 0.1% saponin, 0.5% Tween 20 in PBS (protocols 5 and 6) (Fig. 2). 6. Permeabilization buffers for flow cytometry: (a) 0.5% Tween 20 in SB (protocol 1) (Fig. 2). (b) 0.5% saponin in SB (protocols 2 and 4) (Fig. 2). (c) 0.5% saponin, 0.5% Tween 20 in SB (protocols 3 and 5) (Fig. 2). (d) 0.5% saponin, 0.5% Tween 20, 10 mg/ml RNAse in SB (protocol 6) (Fig. 2). 7. MACS buffer: PBS supplemented with 0.5% BSA and 2 mM EDTA.
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Protocol 2
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Fig. 2. Detection of BCL11B by Flow Cytometry. Total splenocytes were surface stained with anti-CD4-FITC and anti-CD8PE-Cy7 antibodies as described in detail in Material and Methods. The cells were further fixed and permeabilized for exposure of the nuclear epitopes. Protocols 1–6 only differ in the composition of the fixation and permeabilization buffers, as detailed in steps 5 and 6 of Subheading 2.4. In all six protocols anti-BCL11B antibody clone BL1801 was used, followed by secondary Alexa 647-conjugated goat antirabbit secondary antibodies. Histograms depict BCL11B in the gated CD4+ T lymphocytes (solid black line), CD8+ T lymphocytes (dotted black line), and CD4−CD8− cells (solid gray line); secondary antibodies are shown in shaded gray.
8. BN Buffer: 15 mM Tris pH 7.5, 60 mM KCl, 5 mM MgCl2, 15 mM NaCl, 250 mM sucrose, 0.3% NP40, protease inhibitor cocktail. 9. NEB Buffer: 25 mM Tris pH 8.0, 250 mM NaCl, 10% glycerol, 0.2% NP40, protease inhibitor cocktail. 10. Blocking buffer for immunocytochemistry: 4% rabbit serum in PBS. 11. Immunocytochemistry wash buffer: 1% Tween in PBS.
3. Methods 3.1. Overview
Here we developed and optimized a methodology for rapid and convenient detection of the transcription factor BCL11B in T lymphocyte subpopulations using flow cytometry, which is likely suitable, with further optimization, for detection of other transcription factors which are expressed in T lymphocytes. The methodology was
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optimized based on the use of various fixation and permeabilization buffers (items 5 and 6 in Subheading 2.4). Detection of BCL11B by western blot analysis of the nuclear fraction required at least 0.5 × 106 purified CD4+ T lymphocytes (Fig. 1c). Conversely, the use of flow cytometry only required 3 × 104 cells and allowed us to detect BCL11B simultaneously in the CD4+ and CD8+ T lymphocyte subpopulations without their prior purification (Figs. 2 and 3). In addition, flow cytometry enabled the quantification of BCL11B BL1800 100
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CD4+ CD8+ CD4–CD8– Fig. 3. Additional anti-BCL11B antibodies recognize nuclear epitopes exposed through the optimized method of staining. Total splenocytes were surface stained with anti-CD4-FITC and anti-CD8-PE-Cy7 antibodies, fixed and permeabilized using Protocol 6, with the buffers detailed in steps 5 and 6 of Subheading 2.4. Anti-BCL11B antibodies, clones BL1800, BL1799, and BL1798, were used for BCL11B detection, followed by Alexa-647-conjugated goat antirabbit secondary antibodies. The histograms show BCL11B on gated CD4+ T lymphocytes (solid black lines), CD8+ T lymphocytes (dotted black lines), and CD4−CD8− cells (solid gray line); secondary antibodies are shown in shaded gray. Mean fluorescence intensities (MFI) are indicated.
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levels through MFI (Fig. 3). In brief, to detect BCL11B by flow cytometry, total lymphocytes were first stained for the surface markers CD4 and CD8, followed by fixation with the buffers described in item 5 of Subheading 2.4, further followed by a permeabilization step with the buffers described in item 6 of Subheading 2.4, preceding the staining with anti-BCL11B antibodies. The protocols only differ in the compositions of the fixation and permeabilization buffers. We found that the addition of the nonionic detergents during both the fixation and permeabilization buffers (items 5 and 6 in Subheading 2.4) allowed for the optimal detection of the transcription factor BCL11B (Fig. 2, Protocols 5 and 6). We used two nonionic detergents, saponin, known to permeabilize cellular membranes (15), and Tween 20, a polysorbate surfactant, known to permeabilize membranes and stabilize nuclear antigen–antibody binding (16). Neither saponin nor Tween 20 alone, or saponin and Tween 20 together added only in the permeabilization buffer (items 5 and 6 in Subheading 2.4) (Fig. 2, Protocols 1–3) allowed optimal detection of BCL11B. Addition of saponin in both the fixation and permeabilization buffers (items 5 and 6 in Subheading 2.4) improved the detection (Fig. 2, Protocol 4). Addition of both saponin and Tween 20 in both the fixation and permeabilization buffers (items 5 and 6 in Subheading 2.4) further improved the detection (Fig. 2, Protocol 5). Due to the high abundance of RNA in the cells, we rationalized that the addition or RNase may further reduce the nonspecific background. Indeed, further addition of RNase in the permeabilization buffer (item 6 of Subheading 2.4) improved the staining and reduced the background (Fig. 3, Protocol 6) (Note 4). Importantly, the optimized protocol allowed the preservation of the surface markers CD4 and CD8 (Fig. 4) (Note 5).
Fig. 4. Staining for the surface markers CD4 and CD8 is preserved following fixation and permeabilization with the optimized method of staining. Total splenocytes were surface stained as in Figs. 2 and 3. Fixation and permeabilization employed the buffers described in steps 5 and 6 of Subheading 2.4 for Protocol 6, followed by staining with anti-BCL11B antibody clone BL1801. The dot plot graph depicts the percentages of CD4+ and CD8+ T lymphocytes in the spleen.
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Three additional antibodies covering distinctive epitopes of BCL11B, BL1800, BL1798, and BL1799, were tested with the optimized protocol 6 containing saponin and Tween 20 in both fixation and permeabilization buffer, as well as RNase in the latter (items 5 and 6 in Subheading 2.4). BL1800, BL1798 allowed the detection of BCL11B (Fig. 3), while BL1799, though previously demonstrated to work both in western blots and immunoprecipitations (data not shown) could not detect BCL11B by flow cytometry (Fig. 3) (Note 6). In conclusion, antibodies BL1801, BL1800, and BL1798 allow successful detection of BCL11B by flow cytometry (Note 6). 3.2. Tissue and Cells Suspension Preparation
1. Spleen and lymph nodes were removed from the mouse using surgical forceps and cut into pieces using surgical scissors and then placed on a PBS prewet 40 mm cell strainer on the top of a 50 ml conical-bottom tube. 2. Spleen pieces were gently pressed against the strainer with a 5 ml syringe plunger using an up and down motion (Note 7). The strainer was washed with 10 ml CM. 3. The splenocytes were centrifuged at 300 × g for 10 min at 4°C, resuspended in 1 ml prewarmed RBCLB and incubated at 37°C for 5 min. 4. CM was immediately added and the cells were centrifuged at 300 × g for 10 min. Cells were further resuspended in 10 ml CM, mixed, and a small aliquot was removed for counting. The same procedure was also used for lymph nodes without the red blood cell lysis step.
3.3. Staining of Lymph Node or Spleen Cellular Suspensions for Flow Cytometry Analysis
We tested six staining protocols, which only differ in the composition of the fixation and permeabilization buffers (items 5 and 6 in Subheading 2.4), for the detection of the transcription factor BCL11B by flow cytometry.
3.3.1. Surface Staining for CD4 and CD8 Markers
1. In all six protocols staining was performed in V-bottomed 96 well plates. 2. 1 × 106 cells/well were used for each staining. 3. Cells were incubated in 100 ml HB-197 hybridoma supernatant on ice for 20 min to block Fc receptors. 4. 0.4 mg anti-CD4-FITC and 0.08 mg anti-CD8-PE-Cy7 antibodies were then added to the cells to perform the surface staining of these markers. 5. Cell suspensions were then incubated for 30 min on ice in the dark followed by two washes with 150 ml SB.
3.3.2. Fixation for Flow Cytometry
Following surface staining, cells were fixed for 30 min at 4°C in 150 ml fixation buffers. The fixation step was identical for all
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six protocols, except for the composition of the fixation buffers (see above item 5 in Subheading 2.4). 3.3.3. Permeabilization for Flow Cytometry
Permeabilization was further conducted to allow the antibodies against nuclear epitopes to reach their targets. The permeabili zation step was identical for all six protocols, except for the composition of the permeabilization buffers (see above item 6 in Subheading 2.4). 1. Fixed cells were spun at 450 × g for 10 min, and washed twice with 150 ml of the respective permeabilization buffer (see above item 6 in Subheading 2.4). 2. Cells were resuspended in 75 ml permeabilization buffers (see above item 6 in Subheading 2.4), and incubated for 30 min at 4°C.
3.3.4. Addition of Anti-BCL11B and Secondary Antibodies
This step was identical for all six protocols, except for the composition of the permeabilization buffers (see above item 6 in Subheading 2.4). 1. 0.1 mg of anti-BCL11B antibodies (clones BL1801, BL1800, BL1798, and BL1799) were added in 25 ml permeabilization buffers (see above item 6 in Subheading 2.4) for another 30 min on ice (Note 8). 2. Unbound primary antibodies were washed out with permeabilization buffers (item 6 in Subheading 2.4) for a total of three washes. 3. To allow fluorescent detection, 0.16 mg antirabbit IgG-Alexa 647 secondary antibodies were added in 25 ml to the 75 ml cell suspension in permeabilization buffers (item 6 in Subheading 2.4). 4. Cells were incubated for another 20 min at 4°C in the dark, and washed three times, each time with 150 ml of permeabilization buffers (item 6 in Subheading 2.4). 5. Cells were then resuspended in SB buffer and transferred into flow tubes right before reading on the flow cytometer.
3.3.5. Flow Cytometry Data Acquisition and Analysis
3 × 104 cells were acquired on a FACS Calibur flow cytometer, and data was analyzed using Flowjo software (Tree Star, Inc, Ashland, OR).
3.4. Purification of CD4+ T Lymphocytes
1. Single cell suspensions isolated from spleen and lymph nodes by the protocol presented above were washed two more times in PBS and counted. 2. The cells were then washed once in MACS buffer and further resuspended in MACS buffer at 1 × 107 cells/30 µl MACS buffer.
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3. 5 µl CD4 MACS beads were added to each 1 × 10 7 cells followed by incubation in the dark at 4°C for 15 min to allow for the beads to bind. 4. The cells were then washed with 1–2 ml MACS buffer to remove any unbound CD4 MACS beads and then resuspended in 500 µl MACS buffer and applied to the MS column. 5. The columns were hung on the magnetic column holder and the cells were passed through the columns. 6. The columns were washed three times with 500 µl MACS buffer. 7. After the final wash, the column was removed from the magnetic column holder and placed into a fresh 15 ml conical tube. 8. 1 ml MACS buffer was then applied to the column and the CD4+ T cells were flushed out by firmly pushing the plunger into the column (Note 9). 3.5. Immunocyto chemistry
1. CD4+ T lymphocytes were cytospun on 0.1% gelatin-coated coverslips using a Cytospin system adapted for Sorval Legend RT. 2. Following this, cells were fixed in 4% formaldehyde in PBS at 4°C for 20 min, followed by treatment with 0.1% Tween 20 in PBS for 30 min for permeabilization. 3. The cells were then incubated with blocking buffer for 1 h, and then with anti-BCL11B, clone BL1801 primary antibodies (1 µg in 200 µl) or prebleed rabbit serum in blocking buffer, plus 0.1% Tween 20, overnight at 4°C. 4. After four washes the cells were incubated with the FITCconjugated secondary antibodies, 1 µg in 500 µl blocking buffer, plus 0.1% Tween 20, for 1 h at room temperature, followed by three washes and 10 min staining with 1 mg/ml Hoechst 33258 in PBS. 5. Cells were then mounted in Fluoromount and visualized under Olympus BX61 Microscope.
3.6. Cytoplasmic/ Nuclear Fractionation
1. CD4+ T lymphocytes purified as described above were spun down and the cell pellet was washed once with 5 ml cold PBS and then resuspended in 1 ml cold PBS and transferred to a 1.5 ml microfuge tube and spun down again. 2. The supernatant was removed and the cells were resuspended in 100 µl BN buffer plus protease inhibitor cocktail. 3. The cells were incubated for 5 min on ice and then spun down for 10 min at 200 × g in a microfuge at 4°C. 4. The supernatant (cytosolic fraction) was transferred to a new tube.
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5. The pellet was then resuspended in 50 µl NEB buffer plus protease inhibitor cocktail, and incubated for 30 min on a rotator at 4°C. 6. After incubation, the samples were sonicated 3× for 5 s and then spun down at 13,000 rpm (12,000 × g) for 15 min. 7. The supernatant (nuclear fraction) was transferred to a fresh tube. 8. A volume corresponding to at least 0.5 × 106 CD4+ T lympho cytes was used to run on a 7.5% SDS-polyacrylamide gel.
4. Notes 1. The epitopes recognized by BL1800, BL1801, BL1798, and BL1799 map to the following regions of the human BCL11B using the numbering given in Gene ID 64919: residues 500–550 (BL1800), residues 881–894 (BL1801), residues 11–26 (BL1798), and residues 245–259 (BL1799), respectively. The epitopes recognized by BL1800, BL1801, BL1798, and BL1799 are 90, 100, 100, and 94%, respectively, identical between mouse and human proteins. 2. Fixation and permeabilization buffers for flow cytometry were prepared fresh. Due to formaldehyde toxicity, fixation buffers should be prepared in the fume hood. 3. Saponin was prepared as a 10% stock solution in PBS. Saponin is light-sensitive, thus, the stock solution should be kept in the dark. 4. This might not be appropriate for transcription factors known to be associated with RNA, as their three-dimensional organization and epitope exposure might be disturbed by elimination of RNA. 5. It is important that this protocol preserves the surface markers. The optimized protocol is likely to work for other T cell-specific transcription factors using monoclonal or polyclonal antibodies. Tittering of the antibodies and further optimization of the protocol might be necessary. 6. Clone BL1799 did not detect BCL11B in any of the T lymphocytes subpopulations tested. Therefore, BL1799 is not recommended for detection of BCL11B by flow cytometry. Antibodies BL1800, BL1801, and BL1798 allowed detection of BCL11B both in gated CD4+ and CD8+ T lymphocytes, while the CD4−CD8− fraction, known to lack expression of BCL11B, was negative, in agreement with our
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previously published data. BL1798 clone was less optimal in detecting BCL11B in CD8+ T lymphocytes compared to CD4+ T lymphocytes, and also gave a higher background in the CD4−CD8− cell population. This clone requires further optimization. 7. Grinding motion, which can damage the cells, should be avoided. 8. As permeabilization with saponin is a reversible process it is important to continue to work in permeabilization buffer once the cells are permeabilized. However, cells were resuspended in SB after the last wash to allow for the pores of the membranes to close. 9. With the use of one MS column we usually purify approximately 1 × 107 CD4+ T lymphocytes.
Acknowledgments This work was supported by R01 AI067846 from National Institute of Health/NIAID to Dorina Avram. We thank Adrian Avram for the help with the illustration. References 1. Fontenot JD, Rasmussen JP, Williams LM, Dooley JL, Farr AG, Rudensky AY (2005) Regulatory T cell lineage specification by the forkhead transcription factor foxp3. Immunity 22:329–341 2. Ivanov II, McKenzie BS, Zhou L et al (2006) The orphan nuclear receptor RORgammat directs the differentiation program of proinflammatory IL-17+ T helper cells. Cell 126:1121–1133 3. Avram D, Fields A, Senawong T, ToparkNgarm A, Leid M (2002) COUP-TF (chicken ovalbumin upstream promoter transcription factor)-interacting protein 1 (CTIP1) is a sequence-specific DNA binding protein. Biochem J 368:555–563 4. Avram D, Fields A, Pretty On Top K, Nevrivy DJ, Ishmael JE, Leid M (2000) Isolation of a novel family of C(2)H(2) zinc finger proteins implicated in transcriptional repression mediated by chicken ovalbumin upstream promoter transcription factor (COUP-TF) orphan nuclear receptors. J Biol Chem 275:10315–10322
5. Arlotta P, Molyneaux BJ, Chen J, Inoue J, Kominami R, Macklis JD (2005) Neuronal subtype-specific genes that control corticospinal motor neuron development in vivo. Neuron 45:207–221 6. Chen B, Wang SS, Hattox AM, Rayburn H, Nelson SB, McConnell SK (2008) The Fezf2Ctip2 genetic pathway regulates the fate choice of subcortical projection neurons in the developing cerebral cortex. Proc Natl Acad Sci U S A 105:11382–11387 7. Leone DP, Srinivasan K, Chen B, Alcamo E, McConnell SK (2008) The determination of projection neuron identity in the developing cerebral cortex. Curr Opin Neurobiol 18:28–35 8. Arlotta P, Molyneaux BJ, Jabaudon D, Yoshida Y, Macklis JD (2008) Ctip2 controls the differentiation of medium spiny neurons and the establishment of the cellular architecture of the striatum. J Neurosci 28:622–632 9. Leid M, Ishmael JE, Avram D, Shepherd D, Fraulob V, Dolle P (2004) CTIP1 and CTIP2 are differentially expressed during mouse embryogenesis. Gene Expr Patterns 4:733–739
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10. Golonzhka O, Leid M, Indra G, Indra AK (2007) Expression of COUP-TF-interacting protein 2 (CTIP2) in mouse skin during development and in adulthood. Gene Expr Patterns 7:754–760 11. Wakabayashi Y, Watanabe H, Inoue J et al (2003) Bcl11b is required for differentiation and survival of alphabeta T lymphocytes. Nat Immunol 4:533–539 12. Cismasiu VB, Adamo K, Gecewicz J, Duque J, Lin Q, Avram D (2005) BCL11B functionally associates with the NuRD complex in T lymphocytes to repress targeted promoter. Oncogene 24:6753–6764 13. Cismasiu VB, Ghanta S, Duque J et al (2006) BCL11B participates in the activation of IL2
gene expression in CD4+ T lymphocytes. Blood 108:2695–2702 14. Albu DI, Feng D, Bhattacharya D et al (2007) BCL11B is required for positive selection and survival of double-positive thymocytes. J Exp Med 204:3003–3015 15. Jalal F, Jumarie C, Bawab W et al (1992) Polarized distribution of neutral endopeptidase 24.11 at the cell surface of cultured human intestinal epithelial Caco-2 cells. Biochem J 288:945–951 16. Zampieri S, Ghirardello A, Doria A et al (2000) The use of Tween 20 in immunoblotting assays for the detection of autoantibodies in connective tissue diseases. J Immunol Methods 239:1–11
Chapter 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 Marie-Dominique Galibert and Yorann Baron Abstract In order to determine how gene expression is regulated in response to environmental cues, it is necessary to identify the specific interaction between transcription factors and their cognate cis-regulatory DNA elements. Here we have out-lined electrophoretic mobility shift assay (EMSA) and chromatin immunoprecipitation (ChIP) protocols to define in vitro and in vivo USFs specific interacting sequences. The proposed procedures have been optimized for the USFs transcription factor family, allowing the identification of USF-specific targets. Key words: USF transcription factors, Protein/DNA interaction, EMSA, ChIP
1. Introduction Gene expression is a tightly regulated cell-process that is dependent on the loading of the Preinitiation Complex (PIC) machinery in the vicinity of the transcription start site, at the TATA-box binding-site, when present or to the initiator element (Inr) sequence (1). Subsequently, the transcription level is accurately fixed by expression-regulating complexes that include various activators and repressors. Transcription factors (also referred to trans-elements) interact specifically with their DNA-motifs (also referred to cis-elements), present in proximal and distal promoter regions and/or enhancer regions. This does not exclude possible binding downstream to the transcription start-site in exonic and/ or intronic regions (2). Cis-elements can be recognized and activated by one specific transcription factor or by members of a transcription factor family, Paul J. Higgins (ed.), Transcription Factors: Methods and Protocols, Methods in Molecular Biology, vol. 647, DOI 10.1007/978-1-60761-738-9_24, © Springer Science+Business Media, LLC 2010
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increasing the binding-specificity complexity (3, 4). Accordingly, symmetrical E-box motifs are the principle target-binding sites for the members of the conserved b-HLH and b-HLH-LZ transcription factor families (5). The canonical E-box sequence, CANNTG, is only six nucleotides long, with some potential degenerate bases, favoring a broad distribution over the genome. Albeit, only a restricted set of E-box motifs are identified as DNA control elements. Specific binding of E-box motifs is dependent on the nature of the two central nucleotides (CACGTG; CATCTG) and on the flanking ones (NNCANNTGNN). Together and in combination with the respective amount of active E-box binding proteins present in the cells at a given time, it will affect significantly the binding-affinity and selectivity of the different E-box-binding proteins (4, 6–8). The b-HLH-LZ transcription factors are characterized by the presence of a DNA-binding domain: the basic region forming specific and tight interactions with the E-box nucleotides and two dimerization domains: the HLH (Helix-Loop-Helix) and Leucine-Zipper (LZ). Dimerization is a restricted process, regulated by specific rules, and which can only occur between definite b-HLH-LZ members (9, 10). Dimerization is finally required for DNA interaction. The Upstream Stimulating Factors, USFs, are members of the b-HLH-LZ family and consist of two transcription factors, USF-1 and -2, driven by two distinct genes. The human USF-1 is located on chromosome 1 in the q22–23 region while the human USF-2 is assigned to chromosome 19 in q13. Human USF-1 and -2 proteins, which are of 43 and 44 kDa, respectively, are able to homo or heterodimerize together and interact specifically to E-box sequences (9–11). For these reasons, our efforts have focused on developing the tools required to identify accurate USF/E-box complexes participating in gene expression regulation. Before focusing on one putative E-box regulatory element within any particular gene, it is well advised to perform sequence analysis by comparing the human promoter sequence to the ones present in other species. These alignments will determine if the identified E-box element is evolutionary conserved or not. A conserved E-box, including central and flanking nucleotides, in addition to its relative position within the promoter, is generally a good candidate. Identifying specific Protein/DNA interactions requires a combination of in vitro and in vivo assays including Electro phoresis Mobility Shift Assay (EMSA) and Chromatin ImmunoPrecipitation (ChIP). EMSA is based on the observation that proteins that form specific complexes with a DNA fragment (probe) migrate through low-ionic-strength polyacrylamide gels more slowly than the uncomplexed DNA fragment (free probe) creating a band-shift. The inclusion of nonspecific competitors
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(salmon sperm DNA, calf thymus DNA) or poly(dI-dC) improves the sensitivity, allowing the identification in crude nuclear extracts, of eukaryotic proteins that recognize specific DNA sequence elements. In this technique, a radiolabeled DNA fragment is incubated with crude nuclear extract in the presence of nonspecific DNA competitors. The resulting complexes are resolved from the free probe by electrophoresis and visualized by autoradiography or with a phospho-imager (STORM), which allows the precise quantification of the protein/DNA complexes formed (Fig. 1). The sequence specificity of the detected complexes is tested by competitions using the same cold DNA fragment in the presence or absence of E-box motif mutations or cold heterologous DNA fragments of the same length. Heterologous ds-oligonucleotide or mutated E-box ds-oligonucleotides (Fig. 1c) should not affect the formation of specific protein/DNA complexes, while homologous ds-oligonucleotides should completely abolish its formation. Once the specificity of the protein/DNA complex is established, the presence of USF-members within the shifted complex can be tested by competition assays using a previously well-characterized and definite USF binding site. Specific competitions will suggest the presence of USFs members within the complex. Involvement of USF-1 and -2, within the shifted complex, is then resolved by the use of accurate antibodies, recognizing specifically USF-1 or USF-2 and other members of the b-HLH-LZ family (e.g., Mitf, Myc), as controls. In the case of USF-1/USF-2 heterodimers, both antibodies should be used in the same assay. The new antibody–protein–DNA complex formed will migrate again more slowly than the protein–DNA complex, being retained in the upper part of the gel in a “super-shifted” complex. Collectively, the EMSA experiments performed with crude nuclear extract will document the binding specificity of the nucleotide sequence and the nature of the interacting protein(s). However, these in vitro data do not estimate the protein/DNAinteraction in living cell or in the context of chromatin. Chromatin immunoprecipitation (ChIP) assays have been developed to address this question (12–14). ChIP is a powerful tool to identify proteins associated with specific regions of the genome. The initial step of ChIP-assays is the cross-linking of protein–protein and protein–DNA in live cells with formaldehyde. After cross-linking, the cells are lysed and crude extracts are sonicated to shear the DNA. Proteins together with cross-linked DNA are subsequently immunoprecipitated using, in this case, USFs antibodies. Protein– DNA cross-links in the immunoprecipitated material are then reversed, the DNA fragments are purified, and PCR is amplified. Standard PCR or Q-PCR is performed to identify the DNA sequence present in the immuno-precipitated protein/DNA complexes, using specific primers. The limiting step of this assay
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Fig. 1. E-box band shift and competition assays. (a) Free probe or unbound radiolabeled ds-nucleotide migrate at high mobility compared to the protein/DNA complexes. Binding specificity of the protein/DNA complex is evaluated by competition. A onefold excess (1×) of homologous cold competitor is diminished by 2 the protein/DNA complexes formed. Increasing the amount of cold homologous competitor leads to the loss of the protein/DNA complexes. These competitions highlight specific protein/DNA complexes, obtained between the labeled sequence and proteins present in the nuclear extract used (NE). Homologous competitors that are able to compete protein/E-box complexes efficiently are named Class I. They include the WT cold-probe but also mutated ds-oligonucleotide that does not affect the protein/E-box interactions (Panel C). (b) Competition assays performed with Class II and III competitors. Class III competitors are not able to interfere with the formation of specific protein/E-box complexes. They include all ds-oligonucleotides mutated within the core E-box motif (CACGTG). Class II competitors are able to interfere with the formation of specific protein/E-box complexes but with lesser extend than Class I. They include mutated db-oligonucleotide within the flanking E-box nucleotides; those mutations decrease the binding affinity toward a specific transcription factor. (c) Schematic representation of radiolabeledprobe and competitors used in this assay. The 32 ds-nucleotide probe is centered on the E-box sequence of the C4 promoter gene (8).
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is the antibody and its ability to immunoprecipitate the target protein in a chromatin context. In conclusion, EMSA and ChIP are complementary assays required for the study of protein/DNA interactions. EMSA focuses on a specific motif using an in vitro system with short DNA sequences (30 bp) as a bait, while ChIP assays address the question in vivo, a much more complex situation, and will document the binding of a specific transcription within a more expansive promoter region (150–300 bp). Finally, once specific protein/ DNA interaction is documented, it remains essential to prove whether or not this protein/DNA complex is transcriptionally active. Again a combination of in vitro transcription assays (Lucassay with WT or mutated recombinant transcription factor (pCMV-USF) and WT or E-box mutated promoter-region linked to the Luciferase reporter gene) and in vivo studies (e.g., qRTPCR in a knock-out cell line or in response to various stimuli) will establish the impact of the protein/DNA complex on transcription in response to environmental cues.
2. Materials 2.1. Cell Nuclear Extract Preparation
1. Cold-PBS: 8 g NaCl; 0.2 g KCl; 1.15 g Na2HPO4·7H2O; 0.2 g KH2PO4 to 1 l. 2. Cell Lysis Buffer: 10 mM Tris–Hcl (pH 7.5 at 25°C), 10 mM NaCl, 2 mM MgCl2. (store aliquots at −20°C), for 1 l. 3. Nuclear Lysis Buffer: 20 mM Hepes pH 7.9, 420 mM NaCl, 1.5 mM MgCl2, 0.2 mM EDTA pH 8, 25% Glycerol (store aliquots at −20°C). Just before using, add 0.5 mM DTT, 1× proteases inhibitors, and 1× phosphatases inhibitors. 4. 10% NP40 in H2O. 5. Protease inhibitors: Complete Protease Inhibitor Cocktail Tablets (Roche Applied Science). Dissolve one tablet in 2 ml H2O to prepare a stock solution at 25×. Aliquot and store at −20°C. 6. Phosphatase Inhibitors (1×): 1 mM NaF (Sodium Fluoride), 40 nM Na3VO4 (Sodium orthovanadate), 1 mM C3H7O6PNa2 (b Glycerophosphate). Prepare a stock solution at 25× in H2O, aliquot and store at −20°C.
2.2. Probe Preparation
1. 10× Annealing Buffer: 500 mM Tris–Hcl (pH 7.9 at 25°C), 1 M NaCl , 100 mM MgCl2, 10 mM Dithiothreitol, (corresponds to New England Biolabs® – NEBuffer 3). 2. g 32P-ATP (3,000 Ci/mmol) (be cautious!).
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3. 10× Kinase Buffer: 500 mM Tris–Hcl (pH 7.6 at 25°C), 100 mM MgCl2, 50 mM DTT, 1 mM spermidine, 1 mM EDTA. 4. T4 polynucleotide kinase (10 u/ml – Fermentas # EK0031). 5. Mini Quick Spin Oligo Columns (Roche Diagnostic # 11814 397 001). 2.3. EMSA– Competition–Supershift
1. Low ionic strength 6% polyacrylamide gel. 2. 10× Tris–borate electrophoresis buffer (TBE): Per litre mix TRIS-base 108 g, boric acid 55 g, EDTA 9.3 g. Working conditions 1×, pH must be at 8.3. 3. Running buffer (0.5× TBE). 4. 30% Acrylamide/bisacrylamide solution: 29/1 ratio. 5. 10% APS: prepare 10% solution in water, aliquot in 300 ml for single use, and immediately freeze at −20°C. 6. TEMED.
2.4. Probe Binding Reaction Reagents
1. Band shift buffer: Hepes 25 mM, KCl 150 mM, 10% glycerol. 2. poly-dIdC 1 mg/ml. 3. ssDNA 1 mg/ml. 4. DTT 10 mM. 5. Loading buffer 6×: 0.25% Bromophenol-Blue, 0.25% XyleneCyanol (XC), 30% glycerol. 6. Fix Solution: 10% acetic acid, 10% ethanol. 7. Whatman 3MM paper. 8. Saran wrap.
2.5. Competition and Supershift Assays
1. Serial dilution of cold ds-oligonucleotides.
2.6. ChIP
1. PBS: 8 g NaCl; 0.2 g KCl; 1.15 g Na2HPO4·7H2O; 0.2 g KH2PO4 to make 1 l.
2.6.1. Crosslinking Step
2. Antibodies: Anti-USF1 (Santa Cruz, # sc-229); anti-USF2 (Santa Cruz, # sc-861); anti-Tbx2 (Santa Cruz, # sc-17880), preimmune IgG (Sigma).
2. 1.5% formaldehyde in PBS. 3. 0.125 M Glycin in PBS. Filtered through a 0.2 mM unit.
2.6.2. Chromatin Preparation
1. PBS: for 1 l: 8 g NaCl; 0.2 g KCl; 1.15 g Na2HPO4·7H2O; 0.2 g KH2PO4. 2. NCP Buffer-1: EDTA 10 mM, EGTA 0.5 mM, Hepes 10 mM pH 6.5, Triton X-100 0.25%.
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3. NCP Buffer-2: EDTA 1 mM, EGTA 0.5 mM, Hepes 10 mM pH 6.5, NaCl 200 mM. 4. Lysis Buffer: EDTA 10 mM, Tris–HCl 50 mM pH 8.1, 0.5% NP40, SDS 1%. Just before using, add 1× protease inhibitors and 1× phosphatase inhibitors 2.6.3. Immunoprecipitation
1. PBS: 8 g NaCl; 0.2 g KCl; 1.15 g Na2HPO4·7H2O; 0.2 g KH2PO4 to I l. 2. Protein A/G Sepharose Beads. 3. IP-buffer: EDTA 2 mM, NaCl 150 mM, Tris–HCl 20 mM, pH 8.1, Triton X-100 1%. Just before using, add 1× protease inhibitors and 1× phosphatase inhibitors. 4. ssDNA (salmon sperm DNA): 1 mg/ml in H2O. 5. Washing Buffer 1: EDTA 2 mM, Tris–HCl 20 mM, pH 8.1, SDS 0.1%, Triton X-100 1%, NaCl 150 mM. 6. Washing Buffer 2: EDTA 2 mM, Tris–HCl 20 mM, pH 8.1, SDS 0.1%, Triton X-100 1%, NaCl 500 mM. 7. Washing Buffer 3: Tris–HCl 10 mM, pH 8.1, EDTA 1 mM, LiCl 250 mM, deoxycholate 1%, NP-40 1%. 8. Washing Buffer 4: Tris–HCl 10 mM, pH 8.1, EDTA 1 mM. 9. Extraction Buffer: SDS 1%, NaHCO3 0.1 M.
2.7. DNA Preparation
1. NucleoSpin Extract® II, Macherey Nagel, NT Buffer should be replaced by the NTB Buffer.
2.8. Real Time PCR
1. Primers (10 mM): about 20 bases, about 50% GC, amplimers of 100–150 bp. 2. Power SYBR® Green, PCR Master Mix Applied Biosystems. 3. Thermocycler used: Applied Biosystems 7900 HT Fast Real Time PCR System. 4. PCR-program: 1 × 95°C 2 min; 40× (95°C 10 min, 95°C 15 s, 60°C 1 min).
3. Methods 3.1. Cell Nuclear Extract Preparation
The following protocol has been successfully used with 15–20 × 106 501 mel cells, and 15–20 × 106 HaCaT cells. 1. Wash 80% confluent-cells twice with cold-PBS (2× 15 ml). 2. Add 1.5 ml of cold-PBS and collect cells by scraping.
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3. Spin (450 × g – 5 min – 4°C). Keep cells on ice and remove the supernatant. 4. Resuspend gently cell-pellets with 5 V of cooled-Lysis Buffer-1 by pipetting up and down several times, and leave on ice for 10 min. 5. Add 5 ml of 10% NP40 solution (detergent) per 100 ml of cell lysate. Mix gently, and incubate on ice for 5 min (see Note 1). 6. Centrifuge 5–10 min at 13,000 × g – 4°C, remove the supernatant, which corresponds to the cytoplasmic fraction. Wash the nuclei pellet with 100 ml of cooled-Lysis Buffer-1. Spin again and discard the supernatant. 7. Add 1 V of cooled-Nuclear Buffer-2, resuspend the nuclei pellet by pipetting up and down several times, and leave on ice for 10 min. 8. Centrifuge at high speed (10 min at 13,000 × g – 4°C). 9. Transfer and aliquot the supernatant to prechilled tubes. Store at −80°C, to avoid freeze and thaw cycles. 10. Proceed to protein quantification on one aliquot using the Bradford method. 3.2. Probe Selection, Annealing, and Labeling
1. The oligonucleotide sequence of about 25–30 mer should be centered on the target E-box motif. 2. Binding efficiency decreases significantly with short ds-oligonucleotides (12-mer) although binding remains possible (8). 3. The length of the oligonucleotide should be the same for the radiolabeled ds-oligonucleotides and for the cold dsoligonucleotides used as competitors. 4. If SNPs are present within the sequence, both sequences should be used even though it does not affect the cognate E-box motif, it could still modify overall DNA-binding and cooperation. 5. Single stranded (ss) complementary oligonucleotides should be hybridized at the highest concentration (use the oligonucleotide solution stock). 6. Anneal exactly the same amount of sense and antisense ss oligonucleotide by mixing 1 nMole of each ss oligonucleotides in 1× Annealing Buffer and complete with water up to 20 ml. Use a thermocycler, and set the following program: 2¢ 94°C; ramp-cool to 25°C over a period of 45 min; 10¢ 4°C. Alternatively, place tube in a standard heating block at 90–95°C. Remove the heating block from the apparatus, and allow cooling to room temperature (or at least below 30°C) on the workbench. Slow cooling to room temperature should take 45–60 min. Then, transfer to 4°C to follow cooling and store at 4°C until ready to use.
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7. Prepare a working solution at 5 pmole/ml by dilution (1/10). 8. Comparable protocol should be followed to prepare dsoligonucleotides competitors, store at −20°C. 9. 5¢ labeling of double stranded (ds)-oligonucleotides is performed by T4 polynucleotide kinase. 10. The kination reaction is performed in a final volume of 10–20 ml in the presence of g 32P-ATP. 11. Mix 15 pmole of ds-oligonucleotide (3 ml of the 5 pmole/ml working solution), with 1.5 ml of 10× Kinase Buffer, 1.5–3 ml of g 32P-ATP (3,000 Ci/mmol), complete with water up to 14 ml, and add 1 ml of T4 polynucleotide kinase (10 u/ml). 12. Incubate 30 min at 37°C. 13. Add 35 ml of H2O and proceed to the labeled-probe purification with the “Mini quick Spin Oligo Columns” (Roche Diagnostic). Apply the 50 ml labeled-probe solution to the top-center of the prepared Sephadex G-25 column bed. Elute the labeled probe (50 ml) by centrifugation (1,000 × g – 4 min – RT) and dilute it: 1/20. Alternatively, labeled probes can be recovered by phenol/chloroform extraction and ethanol precipitation. But using the spin quick column that retains free g 32P-ATP is more efficient, less time consuming, and more hazard free. 14. Labeled-probes and diluted samples are stored at −20°C or 4°C when used immediately. 15. Probe specific activity should be checked. 16. Double stranded oligonucleotides can also be labeled with Klenow, with comparable results. In this case, use protruding 5¢-end with 32P-dATP or 32P-dCTP. 3.3. EMSA
1. Use low ionic strength 6% polyacrylamide gels. 2. For a standard 20 × 20 cm/1.5 mm thick gel mix in a beaker, 10-ml of 30% acrylamide/bis solution (29:1), 2.5-ml of 10× TBE and complete with water up to 50-ml. Finally add 250 ml of 10% APS and 80 ml of TEMED. Mix again; pour immediately and insert the appropriate comb (usually 20 wells, 1.5-mm). 3. Allow acrylamide to polymerize at room temperature (RT); it should take about 30 min (see Note 2). 4. When you start the binding reaction, prerun for approximately 30 min at 190 V (about 20 mA). 5. Binding reactions are performed in 20 ml final volumes, done on ice, and started by the addition of 13 ml of Band-Shift Buffer, 1 mg of poly-dIdC, 0.5 ml of DTT (10 mM), 2–4 mg
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NE, complete with water to a final volume of 18 ml. Quick spin, shake gently, and finally add 2 ml of diluted probe (0.03 pmole) (see Note 3). 6. Shake gently and incubate on ice for 20 min; then load the binding reaction on the prerun gel. 7. Loading buffer should be added only in one lane as a control and in free lanes to allow a perfect migration, with no edge effect. 8. Electrophoresis is at 190 V for 1 h 30 min/2 h. The front dye (XC) should not run off the gel (around 2–3 cm from the bottom). Otherwise 25-mer free probes are lost. 9. At the end of the run, lift up one glass plate from a corner using a spatula. Submerge for 10 min (until the dye changes color) the gel/glass into 10% acetic acid, 10% ethanol, to fix it. Transfer the gel to a Whatman 3MM paper, cover it with saran wrap, and dry according to the dryer specification, for about 30 min at 80°C. 10. Expose over night (regular film or imager cartridge) and quantify the percentage of shifted bands. 3.4. Competition Assay
1. By serial dilution of the defined cold ds-oligonucleotides, prepare a range of concentrations of –32 to –1 fold molar of cold ds-oligonucleotides relative to the probe. 2. For the competition assay, proceed as previously described except that 1 ml of cold competitor-dilution should be added to the binding reaction. Mix and incubate on ice, before loading. Maintain an equal volume between the competition samples and the control one (no cold ds-oligonucleotides). 3. For relative affinity calculation, a range of 1×, 2×, 4×, 8×… of cold ds-oligonucleotides should be used (Fig. 1a, b). Again mix and incubate on ice, before loading. Maintain an equal volume of 20 ml between the competition samples and the control one.
3.5. Supershift
1. For supershift assay proceed as previously described except that 1 ml (0.2 unit) of antibody should be added at the binding reaction. Mix and incubate for 20 min on ice, before loading. 2. Alternatively, binding reaction can be performed first (20 min incubation on ice). Next, add the antibody to the binding reaction. Mix and incubate for an additional 20 min, before loading. Similar results, with the USFs antibodies are obtained in our hands. 3. To evaluate USF1/USF2-heterodimer and/or USF1-, USF2homodimer DNA-complexes, both anti-USF1 (a-USF1) and
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anti-USF2 (a-USF2) antibodies should be used alone and together. 4. Use an unrelated antibody for the control sample; in our case, we use rabbit-IgG or anti-Tbx2. 3.6. ChIP
3.6.1. Crosslinking Step (Day 1)
The following protocol has been adapted from Metivier and Braustein respective procedures (12, 14) and has been successfully used in experiments with 15–20 × 106 501mel cells and 15–20 × 106 HaCaT cells (about one 80% confluent 15 cm dish, which allows to perform in parallel four immunoprecipitation using four distinct antibodies). 1. Wash 80% confluent-cells twice with PBS (2× 15 ml). 2. Add 5 ml of 1.5% Formaldehyde solution to cover completely the cells, and incubate for 15–20 min at RT, under gently rotation (rotating-platform), to allow protein/protein and protein/DNA cross-linking. Crosslinking for longer periods (>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
