Chromatin Protocols (Methods in Molecular Biology)

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Author: Srikumar P. Chellappan

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METHODS

IN

M O L E C U L A R B I O L O G Y TM

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

METHODS

IN

M O L E C U L A R B I O L O G Y TM

Chromatin Protocols Second Edition

Edited by

Srikumar P. Chellappan H. Lee Moffitt Cancer Center and Research Institute, Tampa, FL, USA

Editor Srikumar P. Chellappan H. Lee Moffitt Cancer Center and Research Institute Tampa, FL, USA [email protected] Series Editor John M. Walker School of Life Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-58829-873-7 e-ISBN 978-1-59745-190-1 DOI 10.1007/978-1-59745-190-1 Library of Congress Control Number: 2008941241 # Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 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 ScienceþBusiness 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. Printed on acid-free paper springer.com

Preface Significant advancements have been made in the study of chromatin structure and function over the past 50 years, but few as spectacular as those made in the last decade. Development of novel techniques and the ability to sequence large stretches of DNA has facilitated in-depth analysis of DNA structure and modifications as well as proteins associated with it. It has been well established that histones and non-histone proteins associated with DNA in chromatin modulate many vital cellular functions, including transcription, replication, repair and gene silencing. Moreover, the modifications of the proteins associated with chromatin have been correlated with many of these functions. Development of novel molecular techniques in the past decade has provided immense momentum to detailed analysis of chromatin, which can now be carried out with remarkable precision. The second edition of Chromatin Protocols compiles many of these techniques that facilitated the explosion of information on chromatin structure and function. The first edition of Chromatin Protocols was published in 1999 and had become the staple of laboratories studying chromatin structure and function. Since its publication, the landscape of chromatin biology has changed immensely. The concept of ‘histone code’ was proposed during the earlier part of this decade and has found universal acceptance. Now it is possible not only to analyze specific histone modifications on each specific nucleosome but such analysis can also be carried out on a genome-wide scale. To a great extent, these advances in the analysis of histone modifications in relation to transcriptional regulation and gene expression were facilitated by the development of chromatin immunoprecipitation (ChIP) assays. Development and refinement of the ChIP assay have led to an understanding of molecular changes associated with histone modifications, transcription factor binding and gene expression in the cell at a level that could only be imagined a decade ago; the progress this technique and its variants have brought to the field of chromatin biology is similar to the progress electrophoretic mobility shift assays brought to transcription factor biology in the early 1980s. These are examples of techniques developed and finetuned in a handful of labs changing the landscape of an entire area of biology. The second edition of Chromatin Protocols encompasses a wide array of techniques spanning from isolation of nucleosomes, assembly of nucleosomes and study of the basic chromatin structure to detailed analysis of histone modifications and chromatin function. Techniques to prepare chromatin assembly extracts, analyze chromatin decondensation and drug-induced premature chromatin condensation, study of reconstituted chromatin, assess DNA topology in chromatin, analyze unique mononculeosomes and a variety of other protocols to study chromatin structure form the initial part of this volume.

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This section is followed by techniques to study DNA damage and repair in the context of chromatin; techniques for monitoring DNA breaks, detailed methods to study transcription-coupled repair in chromatin, techniques for analysis of H2AX phosphorylation to assess DNA damage as well as monitoring DNA breaks in chromatin are included here. A technique to study DNA damage induced by various drugs as well as radiation is included, in addition to a detailed protocol to study genomic aberrations by comparative genomic hybridization. These techniques to study DNA damage in chromatin are followed by an assay to study DNA replication in vitro using mammalian cell extracts. This technique and its variations allow the analysis of the role of different histone and non-histone proteins involved in the replication of DNA. The above sections are followed by detailed protocols to study DNA methylation as well as histone modifications including methylation, acetylation, phosphorylation and ubiquitination. In addition to the analysis of histone modifications, protocols are also provided to study the activities of many of the enzymes involved, including histone demethylases and deacetylases. An elegant technique to study histone phosphorylation by immunostaining is provided, which can be modified to study other histone modifications and chromatin-associated proteins as well. Further, detailed methods to successfully carry out chromatin immunoprecipitation assays are presented. A protocol is presented which describes how genome-wide ChIP analysis can be conducted, using a ChIP-on-chip technique; success of such studies depends to a great extent on the analysis of the data generated and methods to analyze ChIP-on-chip tiling arrays are presented. Imaginative utilization of these ChIP based techniques has immense potential to generate information on various aspects of chromatin biology. The role of telomeres in maintaining chromatin integrity and how alterations in telomerase activities lead to oncogenesis as well as other proliferative disorders have gained substantial interest in recent times. Two protocols are given in this broad context, one describing techniques to study telomerase activity and telomere length and one to study ATM-dependent chromatin modifications. These techniques, we believe, would be very useful for those working in these areas in the context of normal biology of chromatin as well as proliferative disorders associated with telomere function. We believe that these protocols, as well as their creative modifications, will facilitate in-depth molecular analysis of various aspects of chromatin structure and function including transcriptional regulation, DNA repair and replication. As can be imagined, this volume would not have been made possible without valuable contributions from a truly international panel of authors, who are all experts in their fields. My sincere thanks to them for taking the time and effort to pen down the intricate details of their favorite techniques and for their willingness to share them with the readers. In addition, my thanks to Dr. John Walker, the series editor, without whose helpful suggestions and persistence this volume would not have been completed in its present form; it was a real pleasure working with John. It is our genuine belief that the second edition of Chromatin Protocols will be a valuable tool for studying various aspects of chromatin function and would contribute significantly to the development of new techniques as well as ideas in the field of chromatin biology.

Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . v Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ix Color Plates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xiii 1

Preparation of Chromatin Assembly Extracts from Preblastoderm Drosophila Embryos . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .1 Edgar Bonte and Peter B. Becker 2 Analysis of Reconstituted Chromatin Using a Solid-Phase Approach . . . . . . . . .11 Raphael Sandaltzopoulos and Peter B. Becker 3 In Vivo Chromatin Decondensation Assays: Molecular Genetic Analysis of Chromatin Unfolding Characteristics of Selected Proteins . . . . . . . . . . . . . . .27 Sherry L. Winter, Philip Wong, and Mark G. Alexandrow 4 DNA Methyltransferase Probing of Chromatin Structure Within Populations and on Single Molecules. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .41 Carolina Pardo, Scott A. Hoose, Santhi Pondugula, and Michael P. Kladde 5 Visualization of the Expression of HMGN Nucleosomal Binding Proteins in the Developing Mouse Embryo and in Adult Mouse Tissues. . . . . . . . . . . . . .67 Takashi Furusawa and Michael Bustin 6 Drug-Induced Premature Chromosome Condensation (PCC) Protocols: Cytogenetic Approaches in Mitotic Chromosome and Interphase Chromatin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .83 Eisuke Gotoh 7 Analysis of DNA Topology in Yeast Chromatin . . . . . . . . . . . . . . . . . . . . . . . . . .93 Randall H. Morse 8 Preparation and Analysis of Uniquely Positioned Mononucleosomes . . . . . . . .109 Daria A. Gaykalova, Olga I. Kulaeva, Vladimir A. Bondarenko, and Vasily M. Studitsky 9 Monitoring DNA Breaks in Optically Highlighted Chromatin in Living Cells by Laser Scanning Confocal Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . .125 Michael J. Kruhlak, Arkady Celeste, and Andre´ Nussenzweig 10 Methods to Study Transcription-Coupled Repair in Chromatin . . . . . . . . . . . .141 He´le`ne Gaillard, Ralf Erik Wellinger, and Andre´s Aguilera 11 Cytometric Analysis of DNA Damage: Phosphorylation of Histone H2AX as a Marker of DNA Double-Strand Breaks (DSBs) . . . . . . . . . . . . . . . .161 Toshiki Tanaka, Dorota Halicka, Frank Traganos, and Zbigniew Darzynkiewicz

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Detection of DNA Damage Induced by Topoisomerase II Inhibitors, Gamma Radiation and Crosslinking Agents Using the Comet Assay . . . . . . . . .169 Lori A. Hazlehurst Analysis of Genomic Aberrations Using Comparative Genomic Hybridization of Metaphase Chromosomes . . . . . . . . . . . . . . . . . . . . . . . . . . .177 Melanie Carless In Vitro Replication Assay with Mammalian Cell Extracts . . . . . . . . . . . . . . . . .203 Wasia Rizwani and Srikumar P. Chellappan Investigation of Genomic Methylation Status Using Methylation-Specific and Bisulfite Sequencing Polymerase Chain Reaction . . . . . . . . . . . . . . . . . . . .217 Melanie Carless Biochemical Analysis of Arginine Methylation in Transcription . . . . . . . . . . . . .235 Marc Tini, Hina Naeem, and Joseph Torchia In Vitro Histone Demethylase Assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .249 Kenji Kokura and Jia Fang Histone Chaperone as Coactivator of Chromatin Transcription: Role of Acetylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .263 Shrikanth S. Gadad, Jayasha Shandilya, Venkatesh Swaminathan, and Tapas K. Kundu Histone Deacetylase Activity Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .279 Zhigang Yuan, Natalie Rezai-Zadeh, Xiaohong Zhang, and Edward Seto In Vitro and In Vivo Assays for Studying Histone Ubiquitination and Deubiquitination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .295 Ling Zhai, Heui-Yun Joo, and Hengbin Wang Detection of Histone H3 Phosphorylation in Cultured Cells and Tissue Sections by Immunostaining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .311 Jaya Padmanabhan Chromatin Immunoprecipitation Assays: Analyzing Transcription Factor Binding and Histone Modifications In Vivo . . . . . . . . . . . . . . . . . . . . . .323 Smitha Pillai, Piyali Dasgupta, and Srikumar P. Chellappan ChIP on Chip Assays: Genome-Wide Analysis of Transcription Factor Binding and Histone Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .341 Smitha Pillai and Srikumar P. Chellappan ChIP-on-Chip Analysis Methods for Affymetrix Tiling Arrays . . . . . . . . . . . . .367 Sean J. Yoder and Steven A. Enkemann Non-Radioactive Assay Methods for the Assessment of Telomerase Activity and Telomere Length . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .383 Partha P. Banerjee and Shankar Jagadeesh Detecting ATM-Dependent Chromatin Modification in DNA Damage and Heat Shock Response . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .395 Sandeep Misri, Shruti Pandita, and Tej K. Pandita

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Index. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .411

Contributors ANDRE´S AGUILERA CABIMER, Universidad de Sevilla-CSIC, Sevilla, Spain MARK G. ALEXANDROW Molecular Oncology Program, H. Lee Moffitt Cancer Center and Research Institute, Tampa, FL, USA PARTHA P. BANERJEE Department of Biochemistry and Molecular & Cellular Biology, Georgetown University Medical Center, Washington, DC, USA ¨ nchen, Germany PETER B. BECKER Molekularbiologie, Adolf-Butenandt-Institut, Mu VLADIMIR A. BONDARENKO Department of Pharmacology, Robert Wood Johnson Medical School, University of Medicine and Dentistry of New Jersey, Piscataway, NJ, USA ¨ nchen, Germany EDGAR BONTE Molekularbiologie, Adolf-Butenandt-Institut, Mu MICHAEL BUSTIN Protein Section, Laboratory of Metabolism, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA MELANIE CARLESS Department of Genetics, Southwest Foundation for Biomedical Research, San Antonio, TX, USA ARKADY CELESTE Experimental Immunology Branch, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA SRIKUMAR P. CHELLAPPAN Drug Discovery Program, H. Lee Moffitt Cancer Center and Research Institute, Tampa, FL, USA ZBIGNIEW DARZYNKIEWICZ Brander Cancer Research Institute and Department of Pathology, New York Medical College, Valhalla, NY, USA PIYALI DASGUPTA Drug Discovery Program, H. Lee Moffitt Cancer Center and Research Institute, Tampa, FL, USA STEVEN A. ENKEMANN Microarray Core Facility, H. Lee Moffitt Cancer Center and Research Institute, Tampa, FL, USA JIA FANG Molecular Oncology Program, H. Lee Moffitt Cancer Center and Research Institute, Tampa, FL, USA TAKASHI FURUSAWA Protein Section, Laboratory of Metabolism, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA SHRIKANTH S. GADAD Transcription and Disease Laboratory, Molecular Biology and Genetics Unit, Jawaharlal Nehru Centre for Advanced Scientific Research, Jakkur, Bangalore, India ´ HELE`NE GAILLARD CABIMER, Universidad de Sevilla-CSIC, Sevilla, Spain DARIA A. GAYKALOVA Department of Pharmacology, Robert Wood Johnson Medical School, University of Medicine and Dentistry of New Jersey, Piscataway, NJ, USA EISUKE GOTOH Division of Genetic Resources, National Institute of Infectious Diseases, Tokyo, Japan; Department of Radiology, Jikei University School of Medicine, Tokyo, Japan

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DOROTA HALICKA Brander Cancer Research Institute and Department of Pathology, New York Medical College, Valhalla, NY, USA LORI A. HAZLEHURST Department of Interdisciplinary Oncology, Experimental Therapeutics and Drug Discovery Programs, H. Lee Moffitt Cancer Center and Research Institute, Tampa, FL, USA SCOTT A. HOOSE Department of Biochemistry and Molecular Biology and UF Shands Cancer Center Program in Cancer Genetics, Epigenetics and Tumor Virology, University of Florida College of Medicine, Gainesville, FL, USA SHANKAR JAGADEESH Department of Biochemistry and Molecular & Cellular Biology, Georgetown University Medical Center, Washington, DC, USA HEUI-YUN JOO Department of Biochemistry and Molecular Genetics, University of Alabama at Birmingham, Birmingham, AL, USA MICHAEL P. KLADDE Department of Biochemistry and Molecular Biology and UF Shands Cancer Center Program in Cancer Genetics, Epigenetics and Tumor Virology, University of Florida College of Medicine, Gainesville, FL, USA KENJI KOKURA Molecular Oncology Program, H. Lee Moffitt Cancer Center and Research Institute, Tampa, FL, USA MICHAEL J. KRUHLAK Experimental Immunology Branch, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA OLGA I. KULAEVA Department of Pharmacology, Robert Wood Johnson Medical School, University of Medicine and Dentistry of New Jersey, Piscataway, NJ, USA TAPAS K. KUNDU Transcription and Disease Laboratory, Molecular Biology and Genetics Unit, Jawaharlal Nehru Centre for Advanced Scientific Research, Jakkur, Bangalore, India SANDEEP MISRI Washington University School of Medicine, St Louis, MO, USA RANDALL H. MORSE Wadsworth Center, NY State Department of Health and SUNY School of Public Health, Albany, NY , USA HINA NAEEM Department of Oncology, London Regional Cancer Program, Department of Biochemistry, The University of Western Ontario, London, Ontario, Canada ANDRE´ NUSSENZWEIG Experimental Immunology Branch, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA JAYA PADMANABHAN Johnnie B. Byrd, Sr. Alzheimer’s Center and Research Institute, Tampa, FL, USA SHRUTI PANDITA Washington University School of Medicine, St Louis, MO, USA TEJ K. PANDITA Department of Radiation Oncology, Washington University School of Medicine, St Louis, MO, USA CAROLINA PARDO Department of Biochemistry and Molecular Biology and UF Shands Cancer Center Program in Cancer Genetics, Epigenetics and Tumor Virology, University of Florida College of Medicine, Gainesville, FL, USA SMITHA PILLAI Drug Discovery Program, H. Lee Moffitt Cancer Center and Research Institute, Tampa, FL, USA SANTHI PONDUGULA Department of Biochemistry and Molecular Biology and UF Shands Cancer Center Program in Cancer Genetics, Epigenetics and Tumor Virology, University of Florida College of Medicine, Gainesville, FL, USA

Contributors

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NATALIE REZAI-ZADEH H. Lee Moffitt Cancer Center and Research Institute, Tampa, FL, USA WASIA RIZWANI H. Lee Moffitt Cancer Center and Research Institute, Tampa, FL, USA RAPHAEL SANDALTZOPOULOS Department of Molecular Biology and Genetics, Democritus University of Thrace, University Campus, Alexandroupolis, Greece EDWARD SETO H. Lee Moffitt Cancer Center and Research Institute, Tampa, FL, USA JAYASHA SHANDILYA Transcription and Disease Laboratory, Molecular Biology and Genetics Unit, Jawaharlal Nehru Centre for Advanced Scientific Research, Jakkur, Bangalore, India VASILY M. STUDITSKY Department of Pharmacology, Robert Wood Johnson Medical School, University of Medicine and Dentistry of New Jersey, Piscataway, NJ, USA VENKATESH SWAMINATHAN Workman Lab, Stowers Institute for Medical Research, Kansas City, MO, USA TOSHIKI TANAKA Department of Surgery and Clinical Science, Division of Chest Surgery, Graduate School of Medicine, Yamaguchi University, Yamaguchi, Japan; Brander Cancer Research Institute and Department of Pathology, New York Medical College, Valhalla, NY, USA MARC TINI Department of Physiology and Pharmacology, The University of Western Ontario, London, Ontario, Canada JOSEPH TORCHIA Department of Oncology, London Regional Cancer Program, Department of Biochemistry, The University of Western Ontario, London, Ontario, Canada FRANK TRAGANOS Brander Cancer Research Institute and Department of Pathology, New York Medical College, Valhalla, NY, USA HENGBIN WANG Department of Biochemistry and Molecular Genetics, University of Alabama at Birmingham, Birmingham, AL, USA RALF ERIK WELLINGER CABIMER, Universidad de Sevilla-CSIC, Sevilla, Spain SHERRY L. WINTER Molecular Oncology Program, H. Lee Moffitt Cancer Center and Research Institute, Tampa, FL, USA PHILIP WONG Molecular Oncology Program, H. Lee Moffitt Cancer Center and Research Institute, Tampa, FL, USA SEAN J. YODER Microarray Core Facility, H. Lee Moffitt Cancer Center and Research Institute, Tampa, FL, USA ZHIGANG YUAN H. Lee Moffitt Cancer Center and Research Institute, Tampa, FL, USA LING ZHAI Department of Biochemistry and Molecular Genetics, University of Alabama at Birmingham, Birmingham, AL, USA XIAOHONG ZHANG Department of Pathology and Cell Biology, USF College of Medicine, Tampa, FL, USA

Color Plates Color Plates following p 322 Color Plate 1

Color Plate 2

Coimmunostaining analysis of H4 cells using P-histone H3 (serine 10) and alpha-tubulin antibodies: The cells are immunostained using (A) P-histone H3 (serine 10) antibody and (B) alpha-tubulin antibody. Staining was visualized using Alexa 594 goat anti-rabbit or Alexa 488 goat anti-mouse fluorophores, respectively. (C) Represents staining of the same cells using Hoechst to analyze nuclei, and D is the composite image showing all the three stainings. The arrows indicate P-histone H3 positive cells in different phases of the cell cycle. (See discussion on p. 316) Immunohistochemical analysis of brain sections using P-histone H3 (serine 10) and NeuN antibodies: Brain sections are coimmunostained using (A) P-histone H3 (serine 10) and (B) neuron-specific NeuN antibody. Staining was visualized using Alexa 594 goat antirabbit or Alexa 488 goat anti-mouse fluorophores, respectively. (C) Represents the nuclear staining (in pyramidal layer of hippocampus) in sections using Hoechst and (D) is a composite image of all the three stainings. The arrows indicate P-histone H3 positive cells in the brain section. (See discussion on p. 319)

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Chapter 1 Preparation of Chromatin Assembly Extracts from Preblastoderm Drosophila Embryos Edgar Bonte and Peter B. Becker Abstract A rigorous biochemical analysis of chromatin structure and function requires the assembly of chromatin in vitro. A useful alternative to reconstituting nucleosomal arrays from pure or recombinant histones by salt gradient dialysis is the assembly of more complex chromatin from assembly extracts under physiological conditions. Extracts from preblastoderm embryos have proven to be particularly efficient, due to the presence of large stores of native complexes of histones, histone chaperones and ATP-dependent nucleosome spacing factors. The resulting chromatin is an excellent approximation of physiological chromatin in vivo. This chapter describes the preparation of chromatin assembly extracts and the chromatin assembly reaction. Key words: Nucleosome assembly, chromatin reconstitution, in vitro analysis, linker histone, Drosophila.

1. Introduction The preblastoderm embryo of Drosophila melanogaster is characterised by a series of extremely rapid nuclear divisions. In the absence of relevant protein synthesis replication of the genomes and their concomitant assembly into chromatin relies entirely on stockpiles of maternal histones and assembly factors. Cytoplasmic extracts from preblastoderm embryos, which contain all factors required for efficient chromatin assembly, form the basis of one of the most powerful cell-free systems for chromatin reconstitution under physiological conditions (1). Under the appropriate experimental conditions, incubation of plasmid DNA in these extracts results in the formation of long arrays of nucleosomes with regular Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_1 Springerprotocols.com

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spacing, a hallmark of bulk chromatin in nuclei (2). Added single-stranded DNA will be converted into double-stranded DNA before efficient nucleosome deposition (1). Since the crude assembly extract contain a wealth of other chromatin proteins besides histones, the reconstituted chromatin is highly complex (3). The extract is also rich in a number of nucleosome remodelling factors that are responsible for the dynamic properties of reconstituted chromatin, features that most likely reflect the inherent flexibility of chromatin structure in vivo (4–6). Preblastoderm embryos contain no linker histone H1 (7), but H1-containing chromatin can be reconstituted simply by adding an appropriate amount of purified H1 to the reconstitution reaction (1). As embryonic fly development proceeds, the maternal pools of histones are rapidly used up to assemble an exponentially growing number of nuclei. Therefore, extracts made from postblastoderm embryos contain only much less histones. Remarkably, they are as proficient in chromatin assembly as extracts from preblastoderm embryos if they are supplemented with purified histones (8). This observation can be used to assemble chromatin with variant properties from modified histones (2). A prerequisite for the preparation of chromatin assembly extract is a healthy Drosophila population, cultivated at an appropriate scale (9). About 10 g of preblastoderm embryos giving rise to about 10 mL of assembly extract can be collected comfortably during a day from a population of about 150,000–200,000 flies. Here we describe the harvesting of these embryos, the preparation of active chromatin assembly extract and two standard assays for the assessment of reconstituted chromatin. We routinely use this procedure to obtain highly active chromatin assembly extracts from both preblastoderm and postblastoderm embryos. A related procedure specifically for the preparation of extracts from postblastoderm embryos, which differs in several steps from our protocol, has been described by Kadonaga and coworkers (8). Pre- and postblastoderm extracts can be ‘reprogrammed’ with histones of choice if chromatin with defined histone status is required. To this end the endogenous histones are depleted by preincubation of the extract with immobilized DNA followed by addition of the histones of choice (10). Key chromatin assembly components have been purified from these extracts (11, 12). In this chapter we describe the harvesting of Drosophila embryos, the preparation of chromatin assembly extracts and a micrococcal nuclease digestion assay that may be used to optimize chromatin assembly (see Fig. 1.1).

Preparation of Chromatin Assembly Extracts from Preblastoderm Drosophila Embryos

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Fig. 1.1. The regular nucleosomal repeat is visualised with a micrococcal nuclease cleavage assay as described in the text. Chromatin in reactions containing increasing amounts (5–30 ml, indicated above the lanes) of Drosophila embryo extract. Reconstituted chromatin was digested with MNase for 30, 60 and 120 s as described in the text. The three corresponding DNA samples of each reaction are separated by marker lanes (M, 123 bp ladder, BRL).

2. Materials 2.1. Harvesting of the Drosophila Embryos

1. A healthy population of Drosophila melanogaster (see Note 1). 2. Apple juice agar plates. Recipe for about 200 dishes plates: melt 500 g agar in 12 L deionized water (bring to boil). When all agar has dissolved turn heater back to 80C. Add 3.5 L of apple juice, 750 mL sugar bead syrup, 420 mL 10% Nipagin (in ethanol) and mix well. Pour about 80 mL into each large, non-sterile petri dish. Stack 10 dishes each for cooling and hardening overnight. Pack stacks of 16 in the original plastic bags and store at 4C. 3. Yeast paste: 220 mL deionized H2O, 1.4 mL propionic acid (Sigma), 150 g yeast (vacuum packed) (S.I. Lesaffre). 4. Stainless steel sieves (Edelstahlpru¨fsiebe, LINKER; diameter 200 mm, height 50 mm, mesh diameter: 0.125, 0.355 and 0.710 mm) (Kurt Migge, Germany) 5. Paint brush. 6. Embryo wash (EW): 0.7% NaCl, 0.05% Triton X-100 (Sigma).

2.2. Preparation of Dechorionated Embryos

1. EW (see Section 2.1., item 5). 2. Bleach: 13% hypochloric acid (Europe) or chlorox bleach (USA) (see Note 2). 3. Test sieve (mesh diameter 0.125 mm).

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2.3. Washing and Settling of the Dechorionated Embryos

1. EW (see Section 2.1., item 5). 2. 0.7% NaCl. 3. EX-10 (extract buffer): 10 mM N-2-hydroxyethylpiperazineN0 -2-ethanesulfonic acid (HEPES–KOH, pH 7.6), 10 mM KCl, 1.5 mM MgCl2, 0.5 mM ethylene glycol-bis-(baminoethyl ether)-N,N,N 0 ,N 0 -tetraacetic acid (EGTA– KOH, pH 8.0), 10% glycerol, 10 mM b-glycerol phosphate (added freshly), 1 mM dithiothreitol (DTT) (added freshly) and 0.2 mM phenylmethylsulfonyl fluoride (PMSF) (added freshly) (see Note 3). 4. Potter–Elvehjem homogenizer (30 mL) with tight-fitting teflon pestle (Migge, Germany or equivalent).

2.4. Extract Preparation

1. Motor driven homogenizer (Heidolph). 2. 1 M MgCl2. 3. COREX centrifuge tubes. 4. HB-4 rotor ( SORVALL). 5. Polyallomer ultra centrifuge tubes (13 51 mm) (Beckman). 6. Highly liquid paraffin (Merck). 7. SW 55.1 Ti rotor (Beckman). 8. Disposable 5 cm3 syringe (Plastpak) and green needle (G21). 9. Liquid nitrogen.

2.5. Testing of the Chromatin Assembly Activity

1. Plasmid DNA (500 ng/mL). 2. 10 McNAP: 30 mM MgCl2, 10 mM DTT, 300 mM creatine phosphate (Sigma), 30 mM ATP, 1.0 mg creatine phosphokinase (Boehringer Mannheim) per 100 mL (see Note 4). 3. EX-50 buffer (see Note 3). 4. Drosophila extract. 5. Micrococcal nuclease (MNase) (50 U/mL, Sigma, see Note 5). 6. 100 mM CaCl2. 7. MNase stop solution: 2.5% N-lauroylsarcosine (Sigma), 100 mM EDTA–NaOH, pH 8.0 (Merck).

2.6. Purification and Gel Separation of the Nucleosomal "Ladder"

1. 10 mg/mL RNase A (Sigma). 2. 2% dodecylsulphate, Na salt (SDS) (Boehringer, Mannheim). 3. 10 mg/mL proteinase K. 4. 20 mg/mL glycogen (mussels) (Boehringer, Mannheim). 5. Phenol/chloroform/isoamylalcohol (25:24:1) (Amresco). 6. chloroform/isoamylalcohol (24:1). 7. 7.5 M ammonium–acetate (See Note 6).


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8. 96% ethanol. 9. 70% ethanol. 10. TE, pH 7.5: 10 mM Tris–HCl, pH 7.5, 1 mM EDTA. 11. 5 Orange-G loading dye: 50% glycerol, 5 mM EDTA, 0.3% Orange-G (Sigma). 12. 5 Tris–glycine running buffer: 0.25 M Tris, 2.0 M glycine (See Note7). 13. 123 bp DNA marker (Gibco BRL). 14. Agarose ultra PURE (Gibco BRL). 15. 1.3% agarose gel in 1 Tris/glycine running buffer. 16. 10 mg/mL ethidium bromide (Sigma) (Carcinogen !). 17. UV-lamp and gel documentation system.

3. Methods 3.1. Harvesting of the Drosophila Embryos

1. Collect embryos laid on apple juice plates with yeast paste during a 90 min time window. 2. Rinse the embryos off the plates with tap water and using the paint brush, collect the embryos in the 0.125 mm mesh sieve (see Note 8). 3. Using a squeeze bottle with EW buffer rinse the embryos from the sieve into a clean beaker. Store the suspension on ice until extract preparation. Pool the embryos of 4–6 successive collections in EX buffer on ice (see Note 9). 4. Aspirate the cold EW and replace it for fresh EW at room temperature (RT) and allow embryos to settle (see Note 10).

3.2. Collection of Dechorionated Embryos

1. Remove the supernatant and adjust the volume with fresh EW to 200 mL. 2. Add 60 mL of 13% hypochloric acid (RT) and stir vigorously for 3.0 min. 3. Pour the embryos back into the fine collection sieve (mesh diameter 0.125 mm) and rinse vigorously with cold tap water (see Note 11). 4. Transfer the dechorionated embryos to a 500 mL cylinder and wash the embryos with 500 mL EW (RT) and let the embryos settle again (see Note 12). 5. Aspirate off the EW. 6. Wash the embryos with 500 mL 0.7% NaCl (RT) and let them settle again (see Note 12).

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7. Aspirate off the 0.7% NaCl. 8. Wash the embryos with 500 mL EX-buffer (at 4C) and let them settle again. From now on keep embryos on ice and perform all manipulations at 4C with precooled buffers (see Note 12). 9. Aspirate off the EX-buffer as much as possible, without loosing embryos, add additional 100 mL 1 M DTT and 100 mL 0.2 M PMSF to the embryos (see Note 13). 10. Transfer the embryos to the Potter–Elvehjem homogenizer vessel on ice and allow them to settle for at least 15 min. 11. Aspirate off the EX-buffer as much as possible, leaving behind the packed embryos. 3.3. Extract Preparation

1. Stir up packed embryos to resuspend and homogenize them with 1 complete stroke at 3000 rpm and six strokes at 1500 rpm (see Note 14). 2. Measure the volume of the homogenate and add 1 M MgCl2 to a final concentration of 6.5 mM. Mix the MgCl2 immediately with an additional homogenization stroke at 1500 rpm (see Note 15). 3. Centrifuge the extract for 5 min at 10,000 rpm (17,000 g) in a chilled (4C) HB4 rotor, using COREX tubes. 4. Collect the turbid cytoplasmic extract (see Note 16). 5. Transfer the extract to polyallomer ultracentrifugation tubes and fill up, if necessary, with paraffin oil (see Note 17). 6. Centrifuge for 2 h in a chilled (4C) SW55.1 Ti rotor at 45,000 rpm (190,000 g). 7. Isolate the clear extract with a needle and a syringe (see Note 18). 8. Flash freeze 300–500 mL aliquots in liquid nitrogen (see Note 19) and store the aliquots at –80C.

3.4. Testing of the Chromatin Assembly Activity

1. Set up standard chromatin assembly reactions with increasing volumes of extract as follows: 1 mL (500 ng) plasmid DNA, 7 mL 10 McNAP, 10, 20, 30, 40 or 50 mL Drosophila extract, 52, 42, 32, 22 or 12 mL Ex-50 buffer. The final volume for each assembly reaction should be 70 mL. 2. Incubate for 6 h at 26C. 3. Add to each assembled chromatin sample 100 mL MNase mix: 93 mL Ex-50 buffer, 5 mL 100 mM CaCl2 and 2 mL (50 U/ mL) MNase and mix gently.


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4. After 30, 60 and 120 s remove a 55 mL aliquot from the reaction and add to 20 mL MNase stop solution. 3.5. Purification and Gel Separation of the Nucleosomal "Ladder"

1. Add to the MNase-treated aliquots 1 mL (10 mg/mL) RNase A and incubate for 30 min at 37C. 2. Add 8 mL 2% SDS and 5 mL (10 mg/mL) proteinase K and incubate over night at 37C. 3. Add 1 mL (20 mg/mL) glycogen and 10 mL TE pH 7.5. 4. Add 75 mL 7.5 M ammonium acetate. 5. Add 440 mL 96% ethanol and incubate for 10 min in the cold. 6. Centrifuge for 20 min in a tabletop centrifuge, remove supernatant. 7. Wash the pellet with 70% ethanol. 8. Air dry the pellet, do not use a SpeedVac (see Note 20). 9. Resuspend the pellet in 4 mL TE pH 7.5. 10. Add 1 mL 5 Orange-G loading dye. 11. Run the sample in a 1.3% agarose gel with 1 Tris/glycine as running buffer at 5 V/cm (see Note 21). 12. Stain the gel in running buffer containing 1 mg/100 mL ethidium bromide.

4. Notes 1. Reproducible extract preparations mainly depend on a healthy Drosophila melanogaster fly population. The conditions for the maintenance of such a fly population have been described in detail (9). Fluctuations in temperature and humidity and deviations from a regular day/night rhythm affect the embryo yields adversely. We are routinely using 20,000 flies (on average 20 g) in a 45 dm3 fly cage. The newly seeded flies are fed with yeast paste on apple juice agar plates. The embryo collection starts at day 3. 2. Incomplete dechorionation is the main reason for bad extracts. Use only fresh bleach. The bleach should be at room temperature. 3. EX-10 buffer is the extract buffer with 10 mM KCl, in a later stage of the protocol we will use EX-50 buffer, this buffer is identical to the EX-10 buffer with exception of the KCl concentration; the EX-50 buffer has 50 mM KCl. 4. The energy-supplier McNAP has to be prepared fresh every time. Some of the components are unstable even when stored at –80C. Special care should be taken with the creatine

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phosphokinase (CPK). Resuspend one vial of 20 mg lyophylized CPK in 1000 mL 100 mM imidazol (pH 6.6). Divide the solution in 20 mL aliquots and store at –80C for maximally 2 months. Prior to use add to one 20 mL aliquot 380 mL 100 mM imidazol (pH 6.6) and use of this only 1 mL in 100 mL of 10 McNAP. Do not freeze/thaw the aliquot. 5. The unit definition of micrococcal nuclease differs between the suppliers. We refer to the unit definition of Boehringer Mannheim. The lyophylized MNase is resuspended at 50 U/ mL in EX-50 buffer. To obtain a 50 U/mL solution with MNase obtained from Sigma, one should resuspend a lyophylized 500 units vial in 850 mL EX-50 buffer. 6. The 7.5 M ammonium acetate solution should be sterilized by filtering through a 0.2 mM filter. 7. There is no need to adjust the pH of the 5 Tris–glycine stock. 8. Stack the sieves according to decreasing mesh size (0.125 mm down and 0.710 mm top). The tope sieve will retain intact flies, the middle one fly parts and the lower one will retain the embryos. 9. In our hands, 10–15 mL of extract with reproducible high quality is obtained from 80 plates with embryos laid during a 90 min interval. 10. This step is to make sure that the dechorionation will take place at room temperature. Again the main reason for bad extracts is the insufficient dechorionation. Easy aspiration can be performed with, e.g., an 20 mL Pasteur pipette connected to a water vacuum pump. 11. The embryos in the lowest sieve should be rinsed extremely well (for at least 2 min) with a hard beam of tap water to remove all yeast and residual chorion parts that may still attached to embryos. Insufficient dechorionation and washes will result in "floating" embryos that do not settle in the following steps, leading to losses and poor-quality extracts. 12. Floaters will only become apparent when changing the EW buffer to the 0.7% NaCl buffer and in the later step to Ex-10 buffer. Floating embryos will prevent the tight packing of embryos in the homogenizer, which will dilute the extract. 13. The volume of the embryo suspension is roughly 50 mL. Addition of 100 ml DTT and 100 mL PMSF is sufficient to prevent possible protein degradation during the homogenization. 14. Depending on the optimal packing of the embryos the first stroke (a complete movement of the pestle up and down) will be a tough one. Take care that the extract does not heat up during this first stroke and that you do not suck in air in


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between the still-packed embryos and the pestle. This first complete stroke at 1000–1500 rpm is followed by one stroke at 2000 rpm, one at 3000 rpm and 6 strokes at 1500 rpm. 15. The final concentration (including the MgCl2 in the EX buffer is 6.5 mM (13). 16. After the low-speed centrifugation step one obtains a reasonably solid pellet, the cytoplasmic extract and a tight white lipid layer on top. Penetrate the lipid layer along the centrifuge wall with a pipette tip. Avoid the lipid upon collection of the supernatant. 17. The paraffin oil will stabilize the top layer during centrifugation. 18. After the ultracentrifugation step one obtains a solid pellet, a relatively loose, white sediment (cloud) on top of the pellet, the clear extract and a floating lipid layer. Collect the clear extract by puncturing the side of the tube with a green needle (G21) on a 5 cm3 syringe well above the sediment (cloud) but again avoid the lipid layer. Occasionally a white flocculate material is present in the otherwise clear extract. Most of this material can be removed by a 5 min centrifugation in a tabletop centrifuge. The presence of the turbid material does not affect the chromatin assembly reaction. 19. The stability of the prepared extract is fairly high. One can freeze/thaw the extract at least three times without obvious losses of the chromatin assembly activity. Storage of longer than a year should be avoided. In general the protein concentration of the extract is roughly 20–30 mg/mL. 20. Drying the pellet in a SpeedVac may denature the micrococcal nuclease-treated DNA fragments. Inefficient re-annealing may result in a continuum of fragments rather than a defined pattern representing oligonucleosomal DNA. 21. For optimal resolution and easy determination of the nucleosome repeat length (NRL) one should use combs with narrow teeth (1 mm) and a long agarose gel (20 cm). Run appropriate size markers next to the MNase-digested DNA to determine the repeat length (2, 13).

References 1. Becker, P.B., and Wu, C. (1992) Cell-free system for assembly of transcriptionally repressed chromatin from Drosophila embryos. Mol. Cell. Biol. 12, 2241–2249. 2. Blank, T.A., and Becker, P.B. (1995) Electrostatic mechanism of nucleosome spacing. J. Mol. Biol. 252, 305–313.

3. Sandaltzopoulos, R., Blank, T., and Becker, P.B. (1994) Transcriptional repression by nucleosomes but not H1 in reconstituted preblastoderm Drosophila chromatin. EMBO J. 13, 373–379. 4. Tsukiyama, T., and Wu, C. (1995) Purification and properties of an ATP-dependent

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5.

6.

7.

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nucleosome remodeling factor. Cell 83, 1011–1020. Ito, T., Bulger, M., Pazin, M.J., Kobayashi, R., and Kadonaga, J.T. (1997) ACF, an ISWI-containing and ATP-utilizing chromatin assembly and remodeling factor. Cell 90, 145–155. Varga-Weisz, P.D., Wilm, M., Bonte, E., Dumas, K., Mann, M., and Becker, P.B. (1997) Chromatin remodelling factor CHRAC contains the ATPases ISWI and toposiomerase II. Nature 388, 598–602. Ner, S.S., and Travers, A.A. (1994) HMGD, the Drosophila melanogaster homologue of HMG 1 protein, is associated with early embryonic chromatin in the absence of histone H1. EMBO J. 13, 1817–1822. Kamakaka, R.T., Bulger, M., and Kadonaga, J.T. (1993) Potentiation of RNA polymerase II transcription by Gal4-VP16 during but not after DNA replication. Genes Dev. 7, 1779–1795.

9. Shaffer, C., Wuller, J.M., and Elgin, S.C.R., ed. Raising large quantities of Drosophila for biochemical experiments. Methods in Cell Biology, ed. L.S.B. Goldstein and E.A. Fyrberg. Vol. 44. 1994, Academic Press: San Diego. 99–110. 10. Nightingale, K.P., and Becker, P.B. (1998) Structural and functional analysis of chromatin assembled from defined histones. Methods 15, 343–353. 11. Fyodorov, D.V., and Kadonaga, J.T. (2003) Chromatin assembly in vitro with purified recombinant ACF and NAP-1. Methods Enzymol. 371, 499–515. 12. Varga-Weisz, P., Wilm, M. Bonte, E., Dumas, K., Mann, M., and Becker, P.B. (1997) Chromatin-remodelling factor CHRAC contains the ATPases ISWI and topoisomerase II. Nature 388, 598–602. 13. Blank, T.A., Sandaltzopoulos, R. and Becker, P.B. (1997) Biochemical Analysis of chromatin structure and function using Drosophila embryo extracts. Methods 12, 28–35.

Chapter 2 Analysis of Reconstituted Chromatin Using a Solid-Phase Approach Raphael Sandaltzopoulos and Peter B. Becker Abstract Complex experimental strategies involving in vitro reconstituted chromatin or simple chromatin interaction studies are much facilitated by immobilizing the nucleosomal arrays to paramagnetic beads. Chromatincontaining beads can be retrieved from a reaction mix solution on a magnet fast and quantitatively, effectively separating bound, loosely attached and unbound components efficiently. This chapter details a convenient strategy for immobilization of linear plasmid DNA on streptavidin-coated beads, the reconstitution of chromatin on such beads and some fundamental handling procedures. Key words: Biotin-streptavidin, Dynabeads, solid phase, chromatin assembly, chromatin constituents.

1. Introduction The value of a solid support was recognised early in history by the ancient Greek engineer Archimedes who, amazed by the power of the leverage machines that he invented, exclaimed that he could even move the entire planet had he only a suitable solid support to rely on. In biochemistry, sophisticated multi-step experimental procedures require that a substrate is purified and processed through a sequence of reactions under different optimal conditions. Solid-phase techniques are invaluable because they allow instant and quantitative purification of reaction intermediates and readjustment of new reaction conditions. Here we describe a method for chromatin reconstitution on a solid support and present how solid-phase chromatin can be analysed or prepared as a substrate in subsequent reactions. Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_2 Springerprotocols.com

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Chromatin reconstitution in crude extracts from Xenopus oocytes or eggs or Drosophila embryos provides a powerful means to study structure/function relationships in chromatin organisation (1–5). For many of those analyses, e.g. the evaluation of the transcriptional potential of a chromatin template, the chromatin must be purified from the complex reconstitution reaction. The most common method for chromatin purification is its centrifugation through a sucrose gradient. Although efficient, this method is time consuming, does not allow parallel processing of many samples and many

Fig. 2.1. DNA immobilisation strategies (see Note 1).

Analysis of Reconstituted Chromatin Using a Solid-Phase Approach

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loosely associated chromatin components that may be of pivotal importance, e.g. for chromatin dynamics, may be lost during the long centrifugation. By contrast, the solid-phase approach enables the rapid, nondisruptive and quantitative purification of chromatin. A linear fragment of DNA that bears the sequences of interest (i.e. enhancer/promoter and gene coding regions) is biotinylated at one end and then immobilised on streptavidin-coated superparamagnetic beads (Section 3.1 and Fig. 2.1). The bead-coupled template is then subjected to chromatin assembly (Section 3.2.1). Chromatin assembled on immobilised DNA resembles that of soluble DNA with respect to the optimal reconstitution conditions, the kinetics of chromatin assembly, the nucleosomal repeat length, the histone stoichiometry, the association of histone H1, the inhibition of transcription and the association of many non-histone proteins. The immobilised chromatin can then be purified in a magnetic field, washed as desired and then used to purify and analyse chromatin-associated proteins (Section 3.3.1) or to develop assays for putative chromatin-binding proteins. The rapid isolation of immobilised chromatin in a magnetic field facilitates quick buffer exchanges and the efficient removal of soluble components, such as nucleotides or unbound proteins. Therefore, multi-step reconstitutions are facilitated, i.e. reactions in which the chromatin reconstitution must be separated from other steps, such as the interaction of transcription factors prior to chromatin assembly or subsequent chromatin ‘‘remodelling’’ reactions [for examples of applications, see refs. (6–13)]. We also describe how nucleosomes can be reconstituted from pure histones by a salt gradient dialysis procedure on immobilised DNA (Section 3.2.2).

2. Materials 2.1. Immobilisation of DNA

1. Streptavidin-coated paramagnetic beads (Dynabeads M-280, Invitrogen-Dynal). 2. Magnetic particle concentrator (MPC-6, Invitrogen-Dynal). 3. 0.5 mM or 10 mM biotin-21-dUTP in 50 mM Tris–HCl pH 7.5 (Clontech) or 0.4 mM biotin-14-dATP in 100 mM Tris–HCl pH 7.5, 0.1 mM EDTA (Invitrogen-Gibco). Biotin-dCTP is available from Invitrogen-Gibco, but we have not tested it yet. 4. 10 mM a-thio-deoxyribonucleotides, pH 8.0.

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5. Restriction enzymes and 10 digestion buffers (according to the supplier’s recommendation). 6. Klenow (exo–) polymerase. 7. 10 polymerase buffer: 0.1 M Tris–HCl pH 7.5, 50 mM MgCl2, 75 mM DTT). 8. Vent (Exo–) DNA polymerase (New England Biolabs). 9. 10 mg/ml glycogen (Roche Applied Science). 10. Ethanol. 11. Quick Spin columns TE Sephadex G-50 fine (Roche Applied Science) or ChromaSpin column 50-TE (Clontech) or equivalent home-made spin columns. 12. PBS-BSA-NP40: 1.7 mM KH2PO4, 5 mM Na2HPO4, 150 mM NaCl pH 7.4, 0.05% (w/v) BSA, 0.05% v/v nonidet P40 (IGEPAL CA-630). 13. Wash and binding buffer (WB buffer): 2 M NaCl, 50 mM Tris–HCl, pH 8.0, 1 mM EDTA. 14. Lambda DNA (Promega). 15. Kilobase binder reagent (Invitrogen-Dynal). 16. Shaker with regulated speed and temperature (e.g. thermomixer, Eppendorf). 17. TE: 10 mM Tris–HCl pH 7.5, 0.1 mM EDTA. 18. Rotating wheel with regulatable speed. 2.2. Chromatin Reconstitution on Immobilised DNA

2.2.1. Reconstitution in Drosophila Embryo Extracts

1. Chromatin assembly extract (4) 2. 0.5 M MgCl2. 3. McNAP mix (4): 100 mM creatine phosphate, 30 mM ATP pH 8.0, 10 mM DTT, 100 mg/ml creatine phosphokinase. 4. Extract buffer (EX): 10 mM Hepes–KOH pH 7.6, 1.5 mM MgCl2, 0.5 mM EGTA, 10% v/v glycerol, 10 mM b-glycerophosphate, 1 mM dithiothreitol, 0.2 mM AEBSF (4-(2-aminoethyl)benzenesulfonyl fluoride, Sigma-Aldrich). 5. Temperature-regulated chamber with integrated rotating wheel (e.g. hybridization oven).

2.2.2. Nucleosome Reconstitution by Salt Gradient Dialysis

1. Core histones. 2. Two peristaltic pumps. 3. Magnetic stirrer and stirrer bars. 4. DB-1: 2 M NaCl, 10 mM Tris–HCl pH 7.5, 1 mM EDTA, 0.05% nonidet P40, 1 mM b-mercaptoethanol (freshly added). 5. DB-2: 50 mM NaCl, 10 mM Tris–HCl pH 7.5, 1 mM EDTA, 0.05% nonidet P 40, 1 mM b-mercaptoethanol (freshly added). 6. Dialysis tubing Spectra/Por 2, MWCO 12-14,000, 2 ml/cm.


2.3. Analysis of Reconstituted Chromatin

2.3.1. Analysis of Chromatin Proteins

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1. EX-S, (EX, see Section 2.2.1), where S represents the concentration of KCl in mM. 2. EX-50-NP40: extract buffer supplemented with 50 mM KCl and 0.05% v/v nonidet P40 (IGEPAL CA-630). 3. 4 SDS-loading buffer: 200 mM Tris–HCl pH 6.8, 40% (v/v) glycerol, 400 mM b-mercaptoethanol, 4% (w/v) SDS, 0.002% (w/v) bromophenol blue. 4. Equipment for PAGE.

2.3.2. Micrococcus Nuclease Digestion

1. Micrococcal nuclease (Roche Applied Science), 50 units/ml in EX buffer). 2. 5 nuclease stop mix: 2.5% v/v sarkosyl, 100 mM EDTA pH 8.0. 3. TE: 10 mM Tris–HCl, 1 mM EDTA pH 8.0. 4. 5 Orange loading buffer: 50% (v/v) glycerol, 5 mM EDTA pH 8.0, 0.3% w/v Orange G. 5. Equipment for agarose gel electrophoresis.

3. Methods 3.1. Immobilisation of DNA

3.1.1. Digestion of DNA

1. Cleave plasmid DNA with an appropriate pair of restriction enzymes (example is given for ClaI and EcoRI; see Note 1 and Fig. 2.1) as follows: Mix 40 ml of supercoiled plasmid (1 mg/ml = 40 mg), 5 ml 10 digestion buffer, and 5 ml ClaI (10 u/ml). Incubate for 3 h at 37C. 2. Assure complete linearisation by analysing 0.2 ml of the digest by electrophoresis on an 0.8% agarose gel and staining with ethidium bromide (see Note 2). 3. Add 50 ml H2O, 11 ml 10 digestion buffer, and 10 ml EcoRI (10 u/ml). Adjust the final volume to 160 ml with H2O. Incubate for 3 h at 37C. 4. Precipitate DNA: add 16 ml 3 M sodium acetate pH 5.3. Mix. Add 480 ml ethanol, mix. Incubate for 10 min on ice. 5. Spin 15 min at top speed in a tabletop centrifuge. Discard supernatant. 6. Wash pellet with 800 ml 80% ethanol. Dry pellet 2 min in speed vac without heating. 7. Dissolve pellet thoroughly in 40 ml TE.

3.1.2. Biotinylation of DNA

1. Add 7.5 ml 0.4 mM biotin-14-dATP, 1.2 ml each of 10 mM a-thio-dTTP, 10 mM a-thio-dCTP and 10 mM a-thio-dGTP

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(see Note 3), 6.0 ml 10 polymerase buffer and 3.5 ml of 5 units/ml Klenow (exo–). 2. Incubate for 1 h at 37C (see Note 4). 3.1.3. Removal of Free Biotin (see Note 5).

1. Resuspend the matrix of a Quick spin sephadex G-50 TE spin column. Uncap the top then the bottom of the column. 2. Place in a reaction tube provided (without lid) and let drain in a vertical position (about 5 min). 3. Empty the reaction tube and put the column (together with the reaction tube) in a 15 ml Falcon tube. 4. Spin for 1 min at 1100 g. 5. Discard flow-through and spin at 1100 g for 2 min. 6. Replace the collection tube by a fresh one. Apply the biotinylation reaction slowly at the centre of the resin without touching the resin. 7. Spin for 2 min at 1100 g and collect flow-through. The volume of your sample should stay constant (approx. 60 ml). Measure optical density at 260 nm to define DNA concentration in order to account for the losses during gel filtration. Usually the losses are between 10 and 30%. 8. Add 200 ml 2 WB buffer and 140 ml H2O. This is the coupling mix that is ready to be added to the beads. The final NaCl concentration must be 2 M. Save 1 ml for testing immobilisation efficiency.

3.1.4. Coupling of DNA to Dynabeads

1. Resuspend beads well. 2. Remove appropriate amount of bead suspension from the vial. About 1 mg of beads (100 ml) is required for the immobilisation of 1 pmol of DNA (see Note 6). For example, since 1 pmol of a 5 Kb DNA fragment is 3.3 mg, 1.210 ml (40/3.3 times 100) of bead suspension is needed to immobilise 40 mg of fragment. 3. Place tube on MPC (magnetic particle concentrator) for 1 min. 4. Discard supernatant. 5. Wash beads in 300 ml PBS-BSA-NP40. 6. Wash beads twice with 300 ml WB. 7. Resuspend beads in coupling mix (see Section 3.1.3, step 8). 8. Rotate at room temperature for at least 3 h or overnight. 9. Concentrate beads and remove supernatant. 10. Check 10 ml of supernatant (equivalent to originally 300 ng) on 0.8% agarose gel alongside the uncoupled aliquot (see Note 7).


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11. Resuspend DNA beads in WB buffer at a concentration of 30 ng of immobilised DNA per ml of buffer and store at 4C (under these conditions, they can be stored for several months). 3.1.5. Efficient Immobilisation of Very-Long DNA (see Note 6).

1. Mix: 300 ml lambda DNA (100 mg, 40 ml 10 Vent polymerase buffer, 8 ml 10 mM a-thio dGTP, 8 ml a-thio 10 mM dCTP, 8 ml 10 mM a-thio dATP, 4 ml 10 mM biotin-21dUTP, 5 ml Vent (exo–) DNA Polymerase (2 units/ml) and 27 ml H2O (total volume is 400 ml). 2. Incubate for 30 min at 76C 3. Add 40 ml 3 M sodium acetate, pH 5.3 and mix gently. 4. Add 1100 ml absolute ethanol. Mix and incubate for 5 min on ice. 5. Spin for 10 min at top speed in a tabletop centrifuge. 6. Wash pellet twice with 70% ethanol. 7. Dry and resuspend in 300 ml H2O (approximately 1 pmol/ 100 ml) (see Note 8). 8. Add an equal volume of 2 WB buffer and transfer to equilibrated beads (steps 1–7 of 3.1.4). Then add 1/4 of this volume of kilobase binder reagent (see Note 9). Mix gently. 9. Rotate at room temperature for overnight. 10. Check immobilisation efficiency and store DNA beads as in Section 3.1.4, steps 9–11 (see Note 8).

3.2. Chromatin Reconstitution on Immobilised DNA

3.2.1. Chromatin Reconstitution Using Drosophila Embryo Extracts

1. Resuspend stock of immobilised template. Pipette out appropriate amount of bead-DNA. About 900 ng of DNA is sufficient for a MNase assay or analysis of bound histones by silver staining. 2. Concentrate on the MPC. Remove supernatant and wash once with 300 ml of PBS-BSA-NP40. 3. Wash again with 300 ml EX-NP40. 4. Prepare chromatin assembly reaction by mixing 70 ml chromatin assembly extract, 12 ml McNAP and 38 ml EX buffer for each 900 ng of DNA (see Note 10). 5. Concentrate bead DNA, remove supernatant and resuspend beads in complete chromatin assembly reaction. 6. Transfer to 250 ml micro test tubes (see Note 11). 7. Rotate at 26C for up to 6 h, the rotation axis being perpendicular to the longitudinal axis of the tube. 8. Check occasionally for aggregation of beads. If necessary disperse clumps by gently tapping the tube. Some clumping

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may occur during the first 1–2 h of the assembly reaction. If beads are redispersed once, they usually do not clump again. 3.2.2. Chromatin Reconstitution Using Purified Histones

1. During a salt gradient dialysis reconstitution, histones and DNA are first dialyzed into high salt buffer DB-1 (see Note 12). During overnight dialysis the salt concentration is reduced by diluting the dialysis buffer with low salt buffer while keeping the volume of the dialysis constant. 2. Set up a beaker with 600 ml DB-1 buffer at 4C and prepare a larger container with 3 L DB-2. Use two peristaltic pumps and appropriate tubing to pump DB-2 at a rate of 3 ml/min into the dialysis beaker containing DB-1 while at the same time pumping the equivalent volume out of the diluted dialysis buffer into a waste container. This set-up assures that the volume of the dialysis reaction remains constant while the salt concentration is reduced. 3. Prepare the samples. Mix 60 ml 5 M NaC1, 82.25 ml TE, 3.75 ml 20 mg/ml BSA, and 4 ml (0.375 mg/ml) purified core histones (see Note 13). 4. Prepare the mini-dialysis chamber (Fig. 2.2). Detach the cap of a 1.5-ml reaction tube by cutting the connecting hinge with a pair of scissors. Cut the remaining tube at the 1.5 ml mark. The cut-off ring will serve as membrane clamp. Cut dialysis membrane (12-14,000 MWCO) to 2 cm 3 cm pieces. Equilibrate membrane pieces to DB-1 for 30 min. 5. Pipet 50 ml of bead-DNA suspension (30 ng DNA/ml) into a reaction tube. Concentrate the beads on an MPC. Discard supernatant and wash once with 200 ml PBS-BSA-NP40.

Fig. 2.2. The preparation of a mini-dialysis chamber (see Section 3.2.2). The entire dialysis chamber containing the nucleosome reconstitution is thrown into the beaker with dialysis buffer. The tumbling of the chamber due to the vigorous stirring will assure that the beads remain suspended during the 16 h dialysis. We acknowledge the help of Udo Ringeisen in preparing this figure.


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Wash again with 200 ml of WB buffer (see Section 2.1). Concentrate beads, discard supernatant, and resuspend beads into the reaction mix prepared at step 2. 6. Transfer the suspension into empty, inverted Eppendorf tube cap. Place a piece of dialysis membrane on top and clamp membrane with the tube ring (see Fig. 2.2). A reaction volume of 150 ml should essentially fill the cap. Avoid trapping air in the cap which will interfere with dialysis. 7. Throw the closed dialysis cap(s) into the dialysis container and start stirring very fast. Vigorous stirring is essential for maintaining beads in suspension. 8. Dialyze for 45 min before you turn on the pumps to dilute the salt concentration. Dialyze until most of DB-2 has been pumped into the dialysis container (15–16 h). 9. Recover bead suspension by puncturing the membrane with a pipet tip. Transfer to a fresh reaction tube and process as desired. 3.3. Analysis of Reconstituted Chromatin

Reconstituted, immobilised chromatin can be purified from the reaction mix and analysed in various ways. Chromatin proteins can be separated by SDS-PAGE and visualised by Western blotting, silver staining, or even Coomassie staining, depending on the scale of the reaction. The quality of the reconstituted chromatin can be tested by visualisation of the correct histone stoichiometry and a regular nucleosomal array by micrococcal nuclease digestion. Chromatin association of proteins of interest can be tested and the correct stoichiometry of core histones verified.

3.3.1. Analysis of Chromatin Proteins (see Note 14)

1. If a smaller tube was used for chromatin assembly reaction transfer all liquid to Eppendorf tubes that fit into the MPC. The small tube may be rinsed with 100 ml EX-50-NP40 to ensure complete recovery. Concentrate on an MPC for 1 min (not longer!) and remove supernatant completely. Be careful not to touch the pellet with the tip. This may lead to losses. 2. Wash twice with 200 ml EX-50-NP40. Resuspend well each time by gently tapping the tubes. Do not pipette to resuspend. Remove all supernatant each time. If droplets are dispersed on the tube walls spin for 15 s at 1000 rpm in a benchtop minifuge if necessary. 3. Suspend beads in 7.5 ml of EX-Y1-NP40 for elution (see Note 15). Concentrate beads and save supernatant. Repeat and pool supernatant for PAGE (total volume 15 ml). 4. Wash beads with 200 ml of EX-Y2-NP40. Discard wash. 5. Proceed to the next salt concentration. Each elution is done by extracting twice in 7.5 ml (save for gel) and a large 200 ml wash (for completeness).

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6. Place all samples for PAGE on the MPC for 2–3 min to remove any trapped beads. Recover supernatant into new tubes containing 5 ml 4 SDS loading buffer. 7. Resuspend beads in 20 ml 1 SDS-loading buffer. Incubate for 10 min at 37C. Do not boil (see Note 16). Concentrate the beads and save supernatant. This sample represents the proteins that are not eluted even with the most stringent wash applied. 8. Denature all samples for 5 min at 95C, separate by 15% SDS-PAGE. 9. Stain gel with silver or transfer to membrane for Western blotting. 3.3.2. Micrococcal Nuclease Treatment (see Note 17)

1. Assemble 900 ng of immobilised DNA into chromatin as described in Section 3.2.1. Concentrate chromatin on MPC and remove supernatant. 2. Wash chromatin twice with 100 ml of EX-Y-NP40 (see Note 15). 3. Wash beads with 50 ml EX-50-NP40. 4. Resuspend in 120 ml EX-50-NP40 containing 5 mM MgCl2, prewarmed at 26C. 5. Add 180 ml of MNase premix (168 ml EX-50, 9 ml CaCl2, 3 ml MNase (5 u/ml) prewarmed at 26 C (see Note 18). 6. After 30 s, 1 min and 8 min recover 100 ml into a tube containing 25 ml of nuclease stop mix and vortex briefly. 7. When all samples are processed, add 1 ml RNase (10 mg/ml) and incubate for 5 min at 37C. 8. Add 2 ml 20% SDS and 5 ml proteinase K (10 mg/ml) and digest overnight at 37C. 9. Concentrate beads on MPC and recover supernatant. 10. Add 90 ml 7.5 M ammonium acetate, pH 5.3 and 0.5 ml glycogen 20 mg/ml. Mix and add two volumes ethanol. 11. Leave on ice for 5 min and spin for 15 min at top speed in a benchtop centrifuge at 4C. 12. Wash pellet carefully with 800 ml of 75% ethanol and air dry on the bench. Do not dry pellet in the speed vac as this may cause DNA denaturation! 13. Take pellet up in 8 ml TE and add 2 ml Orange loading buffer (5 ). 14. Electrophorese on a 1.3% agarose gel in Tris–glycine buffer ((4); see Note 19).


21

4. Notes 1. In order to immobilise a plasmid two restriction enzymes must be selected as follows (see also Fig. 2.1): The plasmid must be linearised with an enzyme leaving a 50 overhang that can be filled in with biotin-21-dUTP or biotin-14-dATP with Klenow polymerase. In order to prevent the coupling via both ends (which may result in the shearing of the DNA) the linearised DNA must be restricted with a second enzyme leaving a site where no biotin will be incorporated during the fill-in reaction (e.g. blunt ends, 30 overhangs or 50 overhangs with GC-rich sequences). If the secondary cut results in two large fragments, a mixture of both fragments will be immobilised. If the secondary enzyme is chosen such that one large fragment and one very small fragment are produced, this fragment may be removed during the subsequent gel filtration step (Fig. 2.1). Ideally, the biotinylated residue should not be the last nucleotide to be incorporated during the fill-in reaction so that it can be protected against exonuclease activity by sealing the ends with a-thio-dNTPs (see Note 3). Some enzymes that we have used to create an end suitable for biotinylation are EcoR I, Spe I, Afl II, Hind III, and Sal I. Not I and Cla I can be used for the other end. These enzymes produce 50 overhang sequences lacking A or T residues which are not filled in with biotin-14-dATP or biotin-21-dUTP. 2. Incomplete restriction enzyme digestion may lead to low coupling efficiency. We routinely check completeness of digestion at each step. Therefore, even when two compatible restriction enzymes are utilised, we prefer to perform the digestions in two steps rather than in one step, in order to monitor digestion efficiency. Digest first with the enzyme that creates the end that will not be biotinylated and assure complete linearisation. 3. In order to protect the ends from exonuclease invasion that may occur in some experimental systems, we use a-thio-dNTPs in addition to the biotinylated dNTP to fill in the ends which increase the half-life of the ends in crude exonuclease-containing extracts considerably. Ideally, the biotinylated dNTPs should be shielded by 1–2 a-thiodNTPs. In principle, other modified nucleotides (e.g. aminoallyl–dNTPs) which are easier to find could also be used instead of -thio-dNTPs but we have not compared their efficiency. 4. Poor filling-in by Klenow DNA polymerase affects immobilisation. Avoid using ammonium acetate for DNA precipitation as it may inhibit the polymerase. Klenow Exo– is better

22


suited for this application than ordinary Klenow DNA polymerase. 5. Incomplete removal of unincorporated biotin is a common reason for inefficient coupling. Biotin reacts with streptavidin readily and may outcompete the immobilisation of DNA. Spin columns from different suppliers have diverse specifications which should be followed precisely. 6. Coupling efficiency drops drastically with increasing length of DNA to be immobilised. For some applications (6) long chromatin templates may be particularly useful. We describe here a protocol for efficient immobilisation of lambda DNA (50 kb) using the kilobase binder reagent from Dynal. Approximately 1 pmol of lambda DNA can be immobilized per 100 ml of Dynabeads. 7. If coupling was efficient the supernatant from the coupling reaction should be free of DNA (missing band test). In the case of incomplete immobilisation, comparison of band intensities serves to accurately estimate the percentage of immobilised template. Efficiencies higher than 95% are routinely obtained. 8. It is essential to dissolve the pellet completely at this step. Do not vortex to avoid shear. Allow a long time, if possible overnight, to dissolve DNA pellet. In general, minimise manipulations such as extensive pipetting that may shear the concentrated, viscous lambda DNA. We cut the end of the pipette tips with scissors to widen the tip opening. Avoid pipetting up and down in order to resuspend lDNA after its precipitation. 9. See Dynal’s instructions for up-to-date effective concentration. 10. The amount of chromatin assembly extract to be added has to be determined empirically on soluble plasmid DNA. For each amount of extract used, chromatin assembly efficiency is monitored by MNase digestion and agarose gel electrophoresis in order to define the optimum (4). In general 50–90 ml of extract is required for 900 ng of template in a 120 ml reaction. Once the optimal conditions are determined scaling up or down is feasible. If a small amount of immobilised DNA is to be assembled into chromatin it is advisable to fill the reaction up with soluble carrier DNA to keep reaction volume conveniently high rather than scaling down. 11. Reaction tubes of different sizes are used in order to match the volume of chromatin assembly reactions. If there is too much empty space in the tube, the reaction mixture spreads all over the surface of the rotating tube. When possible scale up the chromatin assembly reaction to fill up most of the tube. A small air bubble trapped in the tube will help to


23

maintain the beads dispersed in suspension. Since the magnetic field is much stronger close to the base of the tube, we avoid using relatively big volumes (greater than 600–700 ml) per tube as this would increase the duration of the concentration (in a viscous milieu this can lead to incomplete recovery). Thus when it is necessary to concentrate a greater volume (e.g. when conditioning a great volume of bead suspension for coupling reaction) split the reaction into aliquots and concentrate them successively. After the first aliquot of beads is concentrated and the supernatant discarded, the second aliquot is added to the tube and so on. If the reaction volume is very low, use small (250 ml), elongated tubes. In this case apply the reaction mixture to the bottom of the tube avoiding contact with its walls. The droplet of the reaction mixture will remain at the bottom of the tube due to surface tension. 12. The nucleosome assembly by salt dialysis is a modification of the one described by Neugebauer and Hörz (14). For further descriptions of salt gradient dialysis procedures see (15, 16). Here we concentrate on those modifications to the procedure required when working with immobilised DNA. 13. A ratio of purified core histones to DNA of 1:1 reproducibly results in efficient nucleosome assembly. However, an empirical titration of core histones using soluble DNA may be required. As an internal control in the assembly reactions a short, radioactively labelled and gel purified PCR fragment may be added in the same dialysis chamber with the immobilised DNA. This will serve to determine the efficiency of nucleosome assembly by a band shift assay. Complete nucleosome assembly results in a shift of the probe from free to mononucleosome band. 14. A background of proteins sticking non-specifically to the bead matrix itself is anticipated. Consider the following parameters to optimise the signal-to-noise ratio. First, maximise the amount of DNA per bead by adding an excess of biotinylated DNA in the coupling reaction. Second, pre-adsorb the beads by washing them a couple of times in a buffer containing 0.01% w/v BSA and 0.05% v/v nonidet P 40. This decreases background and also enables easier handling of the beads by reducing their stickiness. Third, different suppliers provide beads with different matrix characteristics. In our experience, Dynabeads gave a low background when used with Drosophila embryo extracts. 15. You have the option to elute proteins sequentially with washes of increasing salt to determine how tightly a protein interacts with chromatin. In general, a buffer containing 400 mM KCl strips off most of the chromatin-associated

24


proteins while core histones require high salt (2 M) for their elution. In the following protocol substitute salt concentration (Y1= salt 1 in mM) in the buffers according to your application. 16. Many proteins that interact with the bead matrix per se are not eluted in SDS-loading buffer unless the beads are boiled. By contrast, chromatin proteins (including histones) are stripped from DNA without boiling. Therefore it is very important to omit boiling of the beads. 17. MNase digestion can be performed with or without prior purification of the template. Here we describe a protocol for MNase digestion of purified chromatin that has been washed. In the case of nuclease treatment without isolation of the DNA from the assembly reaction, approximately ten times more MNase units are required. Conversely, the more stringent the washings of chromatin are the less the nuclease is needed. 18. Upon addition the MNase mix, pipette up and down a couple of times to suspend beads. During longer incubation times resuspend beads once by tapping the tube. Alternatively use the Eppendorf Thermomixer at setting 10. 19. The appearance of the characteristic, ladder-like pattern of DNA fragments generated by MNase analysis and subsequent agarose gel electrophoresis is slightly compromised because only those fragments that are cleaved off the beads (by a double-stranded cut) are recovered for electrophoresis. Since underdigestion may result in only very little DNA on the gel, fine-tuning of the MNase digestion may be required. The nucleosome repeat length of immobilised chromatin assembled in Drosophila extracts is a bit shorter compared to chromatin assembled on plasmid template under identical conditions. This difference is not due to the immobilisation, but rather reflects a difference between linear and supercoiled DNA (17 ).

References 1. Almouzni, G. and Mećhali, M. (1988) Assembly of spaced chromatin. Involvement of ATP and DNA topoisomerase activity. EMBO J. 7, 4355–4365. 2. Becker, P.B. and Wu, C. (1992) Cell-free system for assembly of transcriptionally repressed chromatin from Drosophila embryos. Mol. Cell. Biol. 12, 2241–2249.

3. Tremethick, D. (1999) Preparation of chromatin assembly extracts from Xenopus oocytes. In Methods in Molecular Biology, Vol. 119 Chromatin Protocols, Ed. P.B. Becker, Humana Press, Chapter 12, 175–186. 4. Bonte, E. and Becker, P.B. (1999) Preparation of chromatin assembly extracts from


5.

6.

7.

8.

9.

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11.

preblastoderm Drosophila embryos. In Methods in Molecular Biology, Vol. 119 Chromatin Protocols, Ed. P.B. Becker, Humana Press, Ch. 13, 187–194. Gaillard, P.H. and Roche, D. (1999) Nucleotide excision repair coupled to chromatin assembly. In Methods in Molecular Biology, Vol. 119 Chromatin Protocols, Ed. P.B. Becker, Humana Press, Ch. 17, 231–244. Heald, H., Tournebize, R., Blank, T.A., Sandaltzopoulos, R., Becker, P.B., Hyman, A. and Karsenti, K. (1996) Self organization of microtubules into bipolar spindles around artificial chromosomes in Xenopus egg extracts. Nature 382, 420–425. Sandaltzopoulos, R. and Becker, P.B. (1995) Analysis of protein-DNA interaction by solid-phase footprinting. Meth. in Mol. and Cell Biol. 5, 176–181. Sandaltzopoulos, R., Blank, T., and Becker, P.B. (1994) Transcriptional repression by nucleosomes but not H1 in reconstituted preblastoderm Drosophila chromatin. EMBO J. 13, 373–379. Sandaltzopoulos, R. and Becker, P.B. (1994) Solid phase footprinting: Quick and versatile. Nucleic Acids Res. 22, 1511–1512. Sandaltzopoulos, R. and Becker, P.B. (1998). Heat shock factor increases the reinitiation rate from potentiated chromatin templates. Mol. Cell. Biol. 18, 361–367. Sandaltzopoulos, R. and Becker, P.B. (2003). Analysis of activator-dependent

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transcription reinitiation in vitro. Meth. Enzymol. 370, 487–501. Fletcher, T.M., Ryu, B.-W., Baumann, C.T., Warren, B.S., Fragoso, G., John, S. and Hager, G.L. (2000). Structure and dynamic properties of a glucocorticoid receptorinduced chromatin transition. Mol. Cell. Biol. 20, 6466–6475. Acevedo, M.L. and Kraus, W.L. (2003) Mediator and p300/CBP-steroid receptor coactivator complexes have distinct roles, but function synergistically, during estrogen receptor -dependent transcription with chromatin templates. Mol. Cell. Biol. 23, 335–348. Neubauer, B. and Hörz, W. (1989) Analysis of nucleosome positioning by in vitro reconstitution. Meth. Enzymol. 170, 630–644. Luger, K., Rechsteiner, T. and Richmond T. (1999) Expression and purification of recombinant histones and nucleosome reconstitution. In Methods in Molecular Biology, Vol. 119 Chromatin Protocols, Ed. P.B. Becker, Humana Press, Ch. 1, 1–16. Flaus, A. and Richmond, T. (1999) Base-pair resolution mapping of nucleosomes in vitro. In Methods in Molecular Biology, Vol. 119 Chromatin Protocols, Ed. P.B. Becker, Humana Press, Ch. 4, 45–60. Blank, T.A. and Becker, P.B. (1996) The effect of nucleosome phasing sequences and DNA topology on nucleosome spacing. J. Mol. Biol. 260, 1–8.

Chapter 3 In Vivo Chromatin Decondensation Assays: Molecular Genetic Analysis of Chromatin Unfolding Characteristics of Selected Proteins Sherry L. Winter, Philip Wong, and Mark G. Alexandrow Abstract Of critical importance to many of the events underlying transcriptional control of gene expression are modifications to core and linker histones that regulate the accessibility of trans-acting factors to the DNA substrate within the context of chromatin. Likewise, control over the initiation of DNA replication, as well as the ability of the replication machinery to proceed during elongation through the multiple levels of chromatin condensation that are likely to be encountered, is almost certain to involve the creation of chromatin accessibility. In the latter case in particular, chromatin access will likely need to be a transient event so as to prevent total genomic unraveling of the chromatin that would be deleterious to cells. While there are many molecular and biochemical approaches in use to study histone changes and their relationship to transcription and chromatin accessibility, few techniques exist that allow a molecular dissection of the events underlying DNA replication control as it pertains to chromatin changes and accessibility. In this review, we outline a novel experimental strategy for addressing the ability of specific proteins to induce large-scale chromatin unfolding (decondensation) in vivo upon site-specific targeting to an engineered locus. We have used this system successfully to directly address the ability of DNA replication proteins to create chromatin accessibility and have incorporated modifications to the basic approach that allow for a molecular genetic analysis of the players involved in causing chromatin decondensation by a protein of interest. Here, we briefly describe the nature of the experimental system, its history, and a basic protocol for using the system. Alternative approaches involving co-transfections, concurrent drug treatments, and analysis of co-localizing histone modifications are also addressed, which are useful for extending basic findings to physiological mechanisms. Key words: Acetylation, methylation, chromatin, decondensation, histone modification, Lac operator, LacI DNA-binding domain, protein targeting, heterochromatin.

Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_3 Springerprotocols.com

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28 Winter, Wong, and Alexandrow

1. Introduction The analysis of chromatin decondensation by selected proteins in vivo is achieved through the use of a cell-biological experimental system developed by Andrew Belmont and colleagues (1–3). This system tests the ability of specific proteins to promote large-scale decondensation of compacted chromatin when targeted via a fused LacI DNA-binding domain to a unique and identifiable chromosomal locus. To generate this system, multiple copies of a lac operator (lacO)/dihydrofolate reductase (DHFR) vector were integrated into a Chinese hamster ovary (CHO) cell line, followed by amplification with methotrexate (2). The resulting cell line (A03_1) contains a 90-Mb homogenously staining region (HSR) of 60 amplicons, each consisting of 400 kb of tandemly integrated 14-kb lacO/DHFR vectors and 1000-kb of flanking co-amplified genomic DNA. Significantly, this system allows observation of LacI-targeted proteins with conventional light microscopy and is further capable of allowing visualization of protein-induced chromatin changes in vivo. The HSR can be directly visualized in fixed or living A03_1 cells by expressing and targeting the LacI protein, LacI-GFP derivatives, or proteins of interest fused in-frame to LacI. Under microscopic examination, the HSR normally assumes a condensed dot-like structure and is heterochromatic (2, 3). Binding of LacI-GFP to the HSR in living cells causes no alteration in the morphology of the locus (2). However, the expression and targeting of certain LacI-fusion proteins (e.g., LacI-VP16 and LacI-Cdc45) induce a dramatic decondensation of the HSR (4), likely by recruitment of chromatin remodeling enzymes to the LacI-fused protein (3–5). For example, LacI-Cdc45 targeting causes dramatic chromatin unfolding (4) that is mediated by recruitment of Cdk2 and phosphorylation of histone H1, creating access to higher order chromatin (e.g., in heterochromatin) during replication fork progression, which is usually high for H1 content and typically condensed, unless H1 is phosphorylated (4, 6–11). This remodeling system, although artificially constructed to allow visualization of something that normally cannot be seen, is quite specific in terms of which proteins can remodel, and in terms of how those proteins achieve the remodeling at the mechanistic level. Several proteins are incapable of remodeling chromatin in this system, including Cdc6 (4), geminin (unpublished results), LacI (3), Cdc45–CT (unpublished results; contrasts with wtCdc45 below), and several mutant alleles of BRCA1 (5). Different histone modifications contribute to the decondensation observed with proteins that are effective: VP16-, E2F1-, and p53-decondensed chromatin correlates with core histone hyperacetylation or

In Vivo Chromatin Decondensation Assays 29

hypermethylation and recruitment of HATs (3–5), BRCA1 alleles that can cause decondensation have been found associated with phospho-H2AX and the possible recruitment of COBRA1 (5). Cdc45 decondensation involves recruitment of Cdk2 and H1-P and is sensitive to Cdk2-inhibitory drugs or co-transfection of the Cdk inhibitor p21, but other core histone modifications do not co-localize (4). Consistent with this, and showing the clear specificity of the system, certain Cdc45 mutants lacking the C-terminus (Cdc45–CT) are unable to open the HSR (unpublished results). Importantly, it is in the C-terminus of Cdc45 where the Cdk2 enzyme mediating the remodeling is predicted to bind to RXL cyclin-binding motifs in Cdc45 (12). Finally, it has recently been shown that Cdc45, which is targeted by a Chk1-dependent pathway following BPDE-dependent DNA damage, is less capable of remodeling chromatin in this system when cells are co-treated with BPDE or Chk1 is co-transfected (13). Thus, the mechanisms that cause remodeling in this system are distinctly different for each protein tested and, importantly, derive from physiologically relevant events related to chromatin remodeling that are apparently specific for each targeted protein. For these reasons, this system proves very valuable at allowing investigation of chromatin unfolding by a protein of choice in vivo, as well as investigation of how the particular protein actually brings about the unfolding that occurs. Thus, while the system is cell biological in nature, it has the propensity to allow for a molecular biological dissection of the mechanisms involved in the decondensation events.

2. Materials 1. Phosphate-buffered saline (PBS): 150 mM NaCl, 5 mM Na2HPO4; 1.7 mM KH2PO4 pH to 7.4 and autoclave, Make 1 l of 10 concentrated in water (above is for 1 solution). 2. PBS/NDS: in 1 PBS (made by diluting 10 PBS with water), add normal donkey serum (NDS; Jackson ImmunoResearch; cat#017-000-121) to 1% final concentration (vol/ vol). Normal donkey serum stock (10 ml lyophilized on receipt) is resuspended in 10 ml water, aliquoted, and stored at –20C. 3. Methotrexate solution (MTX) (Calbiochem/EMD) is made in serum-free medium at 1 mM, filter sterilized, and stored at 20C. Final working concentration is 0.3 mM.


4. Formaldehyde solution: 37% formaldehyde liquid solution, Fluka (cat#47608). 5. DAPI solution: 4’-6-diamidino-2-phenylindole (DAPI) is from Sigma (cat#D-9542). Stock is made in water at 5 mg/ ml and stored at –20C. Final working concentration is 0.2–0.5 mg/ml. 6. Coverslips/slides: corning number 2, 22 mm square (cat#2875-22). Pre-cleaned, plain slides are frosted on one side, are 25 75 1 mm, and are from VWR (cat#48300-025). 7. HCl solution: 1.5 N HCl diluted in water and stored at room temperature. As low as 1 N HCl has been used successfully in our hands to allow BrdU visualization. 8. Anti-LacI antibodies: Stratagene rabbit polyclonal anti-LacI (cat#217449-51) and Upstate mouse monoclonal anti-LacI (cat#05-503; clone 9A5) are both necessary. 9. A03_1 cells and expression vectors: Originally generated by the Belmont laboratory (2, 3) and used by other labs including the Alexandrow (4) and Li labs (5). The pRcLac vector was designed by the Li lab and used by Li and colleagues and the Alexandrow lab (see also Support Protocols). Other cell lines designed by Spector and colleagues may be used for these analyses (14, 15), but the system in these lines is different from that described here. 10. Permeabilization buffer: 1% NDS, 0.2% Triton X-100; add to 1 PBS. Make fresh for each experiment as necessary; 2 ml per 35 mm plate. 11. Pre-extraction buffer I: 0.1% Triton X-100, 5 mM MgCl2, 0.1 mM EDTA (ethylenediaminetetraacetic acid) pH 8, made in water as necessary for number of plates used; 2 ml per 35 mm plate. 12. Pre-extraction buffer II: 10 mM HEPES (1-Piperazineethane sulfonic acid, 4-(2hydroxyethyl)-monosodium salt), pH 7.9 with potassium hydroxide (KOH); 10 mM KCl, 1.5 mM MgCl2, 0.5 mM dithiothreitol (DTT), 0.1% Triton X-100, made in water as necessary for number of plates used; 2 ml per 35 mm plate. 13. Mounting medium/kit: Molecular Probes ProLong AntiFade kit (cat#P-7481). Make fresh for each experiment. See also Support Protocols, Section 3.2.2, and Alternate Protocols, Section 3.3.1. 14. Tools required: Forceps and scalpel with handle to place coverslips into 35 mm plates and remove coverslips, respectively. Forceps are sterilized over a Bunsen burner flame prior to gripping coverslips for transfer to plates.


15. Secondary antibodies: All from Jackson ImmunoResearch. Texas Red-conjugated donkey anti-rabbit (H+L) (cat#711-075-152); FITC-conjugated donkey anti-mouse (H+L) (cat#715-095-150); FITCconjugated donkey anti-goat (H+L) (cat#705-095-147). All are resuspended in water as directed and then an equal amount of sterile 100% glycerol is added such that the final glycerol concentration is 50%, and secondaries are stored at –20C. Working concentration is 2 ml per 50 ml solution on each coverslip. 16. Transfection reagent: FuGene 6 reagent from Roche is preferable, but Lipofectamine (or derivatives) from Invitrogen is also suitable. 17. Bromodeoxyuridine (BrdU) (Sigma): dissolved in water as 10 mM stock and stored at –20C. Final working concentration is 10–15 mM. 18. Anti-BrdU antibodies: Roche anti-BrdU mouse monoclonal antibody (Cat# 1170376) is 50 mg/500 ml, stored at –20C and used at 1:20 dilution. Molecular Probes/Invitrogen anti-BrdUAlexa-594 conjugate (Cat# A-21304) is also used at 1:20 dilution and stored at –20C. 19. DNAse I: 10,000 units per ml (Amersham/Pharmacia).

3. Methods 3.1. In Vivo Chromatin Decondensation/ Remodeling Assays

3.1.1. Cell Plating, Transfection Conditions, and Fixation

1. A03_1 cells are plated 2 days before the transfections, since often they display non-uniform adhesion to the plates the next day and many cells have not fully spread out in 24 h. A03_1 cells require culture in the presence of 0.3 mM (final) of methotrexate (MTX) to maintain the presence of the LacO repeats within the HSR. Therefore, cells are routinely cultured in minimal essential medium (MEM; from Cellgro) plus MTX and serum from Hyclone (Fetal Clone I or II work equally well). Cells are plated in 35 mm tissue culture plates containing a sterile square glass coverslip that has been previously baked (see Support Protocols, Section 3.2.3). Cells in a stock 10 cm tissue culture plate at high density (90–100%) are split at a ratio of 1:30 (10 cm plate:35 mm plates), which produces approximately 60–70% confluency on the 35 mm plates after 2 more days of culturing and is a suitable density for transfections and future microscopic analysis. 2. Transfections with LacI-fusion expressing plasmids are performed using a ratio of 5 ml FuGene 6 reagent to 2 mg of


purified DNA. If plasmids are co-transfected, then the ratio will change (see manufacturer for details; Roche). As high as 6–7 mg of total plasmid DNA with 10 ml FuGene 6 reagent per 35 mm plate transfection has been used with success. Transient transfections last 24 h. We also include control transfections in many experiments (see Notes 1 and 2). Protein expression can be verified using immunoblotting with anti-LacI antibodies (see Support Protocols, Section 3.2.4). If one wishes to perform concurrent BrdU labeling analyses, see Alternate Protocols, Section 3.3.2. 3. Cells are washed one time with 2 ml PBS at room temperature for 15 min. One may choose to pre-extract the cells prior to fixation as an alternate approach (see Alternate Protocols, Section 3.3.3). Pre-extraction serves to remove loosely bound proteins from the nuclei prior to permanent fixation with formaldehyde. This is often referred to as the ’chromatinbound’ fraction that remains after pre-extraction and can alter the outcome of a given experiment or suggest new means of interpreting the data. 4. Cells are then fixed on the coverslips in the 35 mm plates for 15 min at room temperature with 2 ml per plate of PBS/2% formaldehyde (stock is made by adding 540 ml 37% formaldehyde to 10 ml PBS). Other fixation methods are useful as well (see Alternate Protocols, Section 3.3.4). 5. Cells are finally washed two times (15 min each time) at room temperature with 2 ml PBS per plate. Plates can be stored at 4C for a week or two under PBS if desired. 3.1.2. Processing Cells for Immunofluorescence

1. Wash fixed cells on coverslips in 35 mm plates two times with 2 ml PBS per plate for 15 min at room temperature. Use a gentle rocking motion, preferably on an orbital shaker. 2. Permeabilize the fixed cells with 2 ml per plate of permeabilization buffer for 5 min at room temperature. Rock gently on an orbital shaker. (see Section 4, Note 3). Keep 1% NDS in all solutions as a competitor/blocking agent from this step until after staining with the secondary antibodies. 3. Wash cells in 35 mm plates two times with 2 ml PBS/1% NDS per plate for 15 min at room temperature. Use a gentle rocking motion on an orbital shaker. 4. Prepare primary antibodies in PBS/1% NDS at the appropriate dilution for each antibody to be used (see Section 4, Note 4). One will need 50 ml of diluted antibodies for each coverslip to be probed. If one is analyzing the LacI-targeted protein together with analysis of co-localization of an endogenous protein, such as a histone or enzyme, then the primary antibody to that protein must be included as well. If


desired, 0.1% Triton X 100 can be included in the dilutions to achieve a higher level of stringency. To analyze two primary monoclonal antibody probings simultaneously, refer to Alternate Protocols, Section 3.3.5. 5. Label frosted slides for each coverslip to be probed. The antibody dilution (50 ml) is placed as a drop onto the appropriate slide. Using the scalpel, carefully pry up the coverslip from the 35 mm plate and use the other hand to grasp the coverslip on its sides. Carefully transfer the coverslip to the frosted slide, turn over, and place face down (cell-side down) onto the drop of diluted antibodies. If the coverslip is accidentally dropped, then use the scalpel to gently ’slice’ a line through one side of the coverslip. If a line appears, the ’cellside’ of the coverslip has been ’sliced’ and should be placed face down. 6. Place slides/coverslips into the humidified chamber (see Support Protocols, Section 3.2.5) in a flat manner. Cover the chamber and incubate for 1–2 h at room temperature. 7. Remove coverslips from the slides and carefully transfer coverslips back to the same 35 mm plates from where they originated. Add 2 ml per plate of PBS/1% NDS and wash two times at room temperature for 15 min on an orbital shaker. 8. Prepare secondary-conjugated antibodies as described above for the primary antibodies (Section 3.1.2, step 4). Secondary antibodies are directed against the species of the primary antibody used above in Section 3.1.2, step 4 (e.g., anti-rabbit, anti-mouse, or anti-goat). Combine Texas Red- and FITC-conjugated antibodies into the same mixture in PBS/1% NDS. A 1:50 dilution of secondaries from Jackson ImmunoResearch has been used. Use the same approaches as described above to place 50 ml antibody drops onto the slides, remove and turn over coverslips, and incubate for 1–2 h at room temperature in the humidified chamber (Section 3.1.2, steps 5 and 6). Keep the slides in the dark from this step forward to reduce background fluorescence. 9. Remove slides/coverslips and carefully transfer coverslip back to their proper 35 mm plate from where they originally came. Add 2 ml per plate of PBS (no need for NDS from this step forward) and wash two times at room temperature for 15 min on an orbital shaker. 10. Stain with DAPI solution for 3–5 min at room temperature in PBS. Use a 1:10,000 dilution of the 5 mg/ml stock of DAPI (final is 0.5 mg/ml, but can go as low as 0.2 mg/ml). Rock on an orbital shaker.


11. Wash two times at room temperature with PBS for 10 min each wash. 12. Mount coverslips on slides using 50 ml of Prolong Anti-fade reagent (see Support Protocols, Section 3.2.2) that is dropped onto each slide, and onto which the coverslips are placed cell-side down as a described above (Section 3.1.2, step 5). An alternate mounting medium can be used if desired (see Alternate Protocols, Section 3.3.1). 13. Place slides/coverslips on a flat surface in the dark overnight at room temperature to dry. 14. Seal the edges of the coverslips on the slides the next morning, or after up to 3 days, with clear nail polish. Slides can be stored at 4C for a short term (up to a week), or at –20C for longer periods prior to viewing by microscopy (see Section 3.4.1 to 3.4.3). 3.2. Support Protocols

3.2.1. Subcloning into pRcLac

3.2.2. Preparation of Mounting Medium

The pRcLac vector was engineered by Li and colleagues (5) and has within it one AscI restriction enzyme site useful for subcloning cDNAs into the vector for in-line expression with an aminoterminal LacI DNA-binding domain tag/targeting domain. To subclone, one of two approaches should be used. First, polymerase chain reaction (PCR) can be used to add AscI sites to both ends of the cDNA, followed by direct ligation into pRcLac. Alternatively, a vector we have generated (pcAscIx2; vector and map available from us upon request) can be used as a shuttle vector to add AscI sites to both ends of the cDNA. For transferring cDNAs into the vector, BamHI and EcoRI restriction sites have been incorporated into pcAscIx2 such that cDNAs will be in the proper reading frame with LacI in the final pRcLac construct. This necessitates that the user already have, or add (e.g., using PCR methods), such restriction sites in-frame with their cDNAs prior to this subcloning. (see also Section 4, Note 5). Mounting medium from Molecular Probes/Invitrogen consists of two components, A and B. Component A is a chemical pellet at the bottom of a small brown tube; Component B is a viscous solution that is largely solid when stored frozen. Component B must be thawed at least an hour prior to use, but we have found that 5–10 s in a microwave oven, with the lid off, is suitable for quick thawing. Dropwise add 1 ml of Component B to a clear microcentrifuge tube. Then transfer the 1 ml solution to the Component A tube. Wait for 15 min at room temperature. Use a micropipette tip to stir and dislodge the pellet. Vortex briefly. Spin out bubbles at full speed in a microcentrifuge. Fifty microliters are used to mount each coverslip, but we have found that one tube will only be suitable for 17–18 slides (rather than 20).


3.2.3. Coverslip Preparation and Transfer

Coverslips are placed into a glass petri dish, covered with aluminum foil, and baked at 80C overnight for 16 h to sterilize. To transfer into 35 mm tissue culture plates, forceps that have been sterilized just prior with a Bunsen burner flame are used to carefully pick up and transfer only one coverslip. Accidental transfer of two coverslips will yield mounted slides in the end that are unreadable with the microscope. Coverslips from Corning prepared in this way do not need to be acid treated.

3.2.4. Simplified Immunoblotting Analysis

To detect LacI-fusion protein expression, transfected cells are washed on 35 mm plates with excess PBS two times, then collected into microcentrifuge tubes with a cell scraper. After a quick spin in a microcentrifuge to pellet the cells (1000 RPM for 20 s), PBS is aspirated and 100 ml of 1 Laemmli protein loading dye (16) is added to each pellet and boiled for 10 min. Five microliters of each sample are loaded onto gels, transferred, and analyzed by immunoblotting with 1:20,000 dilution of the anti-LacI antibodies.

3.2.5. Humidified Chamber Preparation

A plastic dark room developer tray (8 10 inches, 1.5–2 inches deep), or similar, is used to prepare a humidified chamber. Paper towels are wetted with water, then laid in the tray as smooth as possible. Plastic wrap (e.g., Saran Wrap) is used to cover the chamber, and cardboard plates, such are used to sandwich autoradiography cassettes, are laid on top. The chamber is allowed to evaporate water for about 2 h prior to use to create vapor pressure that prevents slides from drying out.

3.3. Alternate Protocols

The ProLong Anti-fade mounting reagent from Molecular Probes/Invitrogen is preferred for durability of mounting and prevention of ultraviolet bleaching of samples. However, an alternative mount can be made as follows: Mix 8.5 ml glycerol (Polysciences cat#00084) with 750 ml PBS. Add 10 mg 1,4phenylenediamine (Aldrich Chemical cat#27,515-8), wrap in aluminum foil, and stir for half a day to overnight. Check that the pH is 6 (using pH paper). Add 20–30 drops of carbonate–bicarbonate buffer to give a final of pH 8. Store in aliquots at 20C wrapped in foil. The carbonate–bicarbonate buffer is itself made by making 0.2 M solution of anhydrous sodium carbonate and 0.2 M solution of sodium bicarbonate, then combining 4 ml of 0.2 M solution of anhydrous sodium carbonate with 46 ml of 0.2 M solution of sodium bicarbonate, and bringing to a final volume of 200 ml with water. The pH of the mixed solutions will be 9.2.

3.3.1. Alternate Mounting Medium Preparation

3.3.2. Concurrent Bromodeoxyuridine (BrdU) Analysis

To analyze DNA replication concurrently with chromatin unfolding, cells are pulse labeled with 10–20 mM BrdU for 30 min prior to fixation. The BrdU solution is added directly to the tissue culture medium. To process cells for BrdU incorporation,


proceed through Section 3.1.2, step 3, and keep non-BrdUlabeled cells rocking in PBS/1% NDS. BrdU-labeled cells are treated with 1.5 N HCl (in water) for 30 min at room temperature on an orbital shaker. Alternatively, use DNAse I to create access for anti-BrdU antibodies (see Alternate Protocols, Section 3.3.6). Cells are washed three times with PBS/1% NDS at room temperature for 10 min each time. The protocol then resumes at Section 3.1.2, step 4. When diluting anti-BrdU antibodies, we use a 1:20 dilution of the Roche or Molecular Probes/Invitrogen antibodies onto the slides/coverslips in PBS/1% NDS (see also Section 4, Note 4). Samples treated with 1.5 N HCl still retain the ability to bind DAPI effectively (at Section 3.1.2, step 10). 3.3.3. Pre-extracting with Detergents Prior to Fixation

Treat non-fixed, PBS-washed cells (from Section 3.1.1, step 3) for 10 min at 4C with (cold) pre-extraction buffer I or II prior to fixation. Then resume with formaldehyde fixation in Section 3.1.1, step 4.

3.3.4. Alternate Fixation Methods

MeOH (100%, cold) or a 1:1 mixture of MeOH:acetone (cold) can be used to fix cells instead of formaldehyde. A 10 min incubation at –20C in either solution will fix cells. Following fixation, dry for 15–30 min at room temperature. Formaldehyde fixation can block certain epitopes for antibody recognition, and these other approaches may alleviate this problem.

3.3.5. Using Two Primary Monoclonal Antibodies Simultaneously

It is not possible to use two primary monoclonal antibodies simultaneously followed by secondary antibody probings that can differentiate between the two primary antibodies. However, if using two monoclonals as primaries is the only way to proceed, then at least one of the primary monoclonal antibodies must be pre-conjugated to a fluorophore. For example, see Section 4, Note 4. To perform the analysis in this manner, proceed through the immunofluorescence assays from Section 3.1.2, steps 1 through 9, using the first non-conjugated primary antibody and secondary antibody probings. Then, fix the coverslips/cells again with formaldehyde as in Section 3.1.1, step 4. Continue through the procedure from Section 3.1.1, step 4, until Section 3.1.2, step 4, at which time add diluted primary antibody of the second monoclonal that is pre-conjugated to a fluorophore. Follow the procedure from Section 3.1.2, step 4 through Section 3.1.2, step 6. There is no need to probe with another secondary, since the second primary antibody is pre-conjugated. Therefore, finish the assays by resuming at Section 3.1.2, step 9. In the end, the procedure will have been performed essentially 1.5 times to achieve analysis of two monoclonals simultaneously.


3.3.6. DNAse I Treatment for BrdU Analysis

DNAse I treatment can be used rather than HCl to denature DNA for anti-BrdU access and recognition. For this, use 50–100 ml of 10 Units/ml DNAse I in PBS/1% NDS on each coverslip turned over onto a slide face down at 37C for 1–2 h in a humidified tissue culture incubator (to prevent drying out of the slides). Then proceed with anti-BrdU staining as described above (see Alternate Protocols, Section 3.3.2). We must caution, however, that DNAse I treatment/accessibility has proven unsuccessful in our hands. For more information on this approach, the reader is referred to a Kennedy and Harlow paper (17).

3.4. Interpretations

Microscopic examination is used to detect the chromatin structures (via fluorescence of the anti-LacI antibody + secondary conjugate that marks the region where the HSR is located and the targeted protein is concentrated). This may entail using a traditional wide-field ultraviolet microscope, or a more sensitive confocal laser scanning microscope system. The A03_1 cells often display two HSRs, with some examples displaying one closed HSR and one open HSR in the same cell. The importance of this is that this chromatin unfolding system derives from a complex cell biological environment. There are many variables influencing the ability of a targeted protein to unfold chromatin (or not). Simple targeting of any protein will not open chromatin, and even proteins that do usually open chromatin may not necessarily lead to a decondensation event at all targeted HSRs for reasons that are unknown.

3.4.1. Data Acquisition

3.4.2. Criteria for Chromatin Decondensation/NondDecondensation

This experimental approach is admittedly a subjective cell biological assay for the chromatin unfolding capability of a protein of interest. However, we use an analysis method that is aimed at mitigating the subjectivity of the assay and incorporates a degree of objectivity into the results. Rather than assign an ‘open’ definition as being larger than 2% of the area of the nucleus (2, 3), which is a measurement that even some arguably closed chromatin structures would fall into based on our studies, we have assigned ‘open’ to structures that are unambiguously open and rather large in shape. Such open structures are easily seen by almost any observer and are generally greater than 10% of the nuclear area. The ‘closed’ definition we use to describe chromatin structures that are clearly still dots and condensed to almost any observer. The ‘closed’ structures tend to comprise 2–5% of the nuclear area in our analyses (4). In many experiments, approximately 4–12% of the total samples show chromatin structures that are not clearly open or closed and are referred to as ‘indeterminate’ structures. The size of such ‘indeterminates’ often comprises 5–10% of the nuclear area, usually toward the smaller side. Although these structures are counted, they are generally excluded from our interpretations, leaving us with strictly open or closed structures


that are clear to most observers. In this manner, our approach at analysis lends a more objective interpretation to the ability (or lack thereof) of a tested protein to induce chromatin unfolding in vivo. 3.4.3. Important Considerations

While this chromatin unfolding analysis system seems capable of further investigating the control of DNA replication or transcription at the HSRs after targeting proteins of interest, it is important to keep in mind that this system was never designed for this purpose. It is strictly a targeting system that allows one to test whether a protein of interest can remodel chromatin or not. However, using co-transfections, drug treatments, or other combinatorial approaches, the initial inroads into deciphering how a protein may be remodeling, and whether this remodeling is physiologically relevant due to supporting evidence gained from the combinatorial approaches, can be made. Common sense dictates that fusion of a large DNA-binding domain (LacI; 40 kilodaltons) will likely interfere in some manner with normal function of the proteins fused to LacI, since this is not the normal structure of the proteins. Nonetheless, this system clearly has tremendous advantages that allow one to make initial discoveries into the nature and possible mechanisms whereby a specific protein decondenses chromatin. It is up to the investigator to find means of relaying these findings to the physiological significance and relevance of the tested protein’s role in normal cell processes. In the end, much of the novelty of this system is dependent on the creativity of the person using the system.

4. Notes 1. Controls may include a LacI-VP16 vector (pNYE4-GFPVP16; see ref. (3) and/or an ‘empty’ vector that only expresses the LacI DNA-binding domain alone (pRcLac alone). We have used the LacI-VP16 vector as a comparison when performing co-localization analyses with antibodies against modified histones (e.g., acetylated or methylated histones). This aids in verifying that the antibodies against endogenous modified histones are working as expected and that the microscopic settings are correct for visualizing co-localization. 2. Co-transfection of other plasmids encoding proteins that modulate the activity of the LacI-targeted protein, or the activity of proposed mediators recruited by the LacI-tagged protein, can be performed. In addition, the transfected cells can be treated with various drugs that modulate the enzymatic activities of proteins thought to be mediators of the chromatin unfolding. In either case, the timing of co-transfection or drug


treatment can also be varied to suit the experimental requirements. Both of these approaches offer a novel means to elaborate on mechanisms that may be involved in the chromatin unfolding mediated by a particular LacI-fused protein being analyzed. 3. It is preferable to use the same species of blocking normal serum as the species of secondary antibody conjugate used in the immunofluorescence assays. This reduces background fluorescence (16). We use secondaries raised in donkey and normal donkey serum throughout. 4. The anti-LacI antibodies used may need titration. The rabbit and mouse anti-LacI antibodies that we use require 1:20,000 dilutions into PBS/1% NDS. In addition, an alternative to using primary antibodies followed by secondary antibody staining is to use pre-conjugated primary antibodies that have a fluorophore already chemically attached. For example, antiBrdU conjugated to Alexa-594 (Molecular Probes/Invitrogen) can be used. 5. An internal in-frame green fluorescent protein (GFP) in the constructs can be used as a control or to aid in visualizing the targeted proteins without antibody staining procedures. Likewise, to analyze fragments of proteins for domains that can remodel, it may be necessary to include a nuclear localization sequence in the final construct to guarantee nuclear import during the transfection.

Acknowledgments We are extremely grateful to Andrew Belmont (University of Illinois, Urbana-Champaign) for providing us with the A03_1 cell line, the LacI-VP16 vector, and helpful suggestions. Work in the Alexandrow lab is supported by funds from the Florida Department of Health, the James and Esther King Foundation (Award 06-NIR-01), and the National Cancer Institute (Lung SPORE, Section 3; PSO-CA11997). References 1. Belmont, A. S., Li, G., Sudlow, G., and Robinett, C. (1999) Visualization of large-scale chromatin structure and dynamics using the lac operator/lac repressor reporter system. Methods Cell Biol 58, 203–222. 2. Li, G., Sudlow, G., and Belmont, A. S. (1998) Interphase cell cycle dynamics of a latereplicating, heterochromatic homogeneously

staining region: Precise choreography of condensation/decondensation and nuclear positioning. J Cell Biol 140, 975–989. 3. Tumbar, T., Sudlow, G., and Belmont, A. S. (1999) Large-scale chromatin unfolding and remodeling induced by VP16 acidic activation domain. J Cell Biol 145, 1341–1354.


4. Alexandrow, M. G., and Hamlin, J. L. (2005) Chromatin decondensation in Sphase involves recruitment of Cdk2 by Cdc45 and histone H1 phosphorylation. J Cell Biol 168, 875–886. 5. Ye, Q., Hu, Y.-F., Zhong, H., Nye, A. C., Belmont, A. S., and Li, R. (2001) BRCA1induced large-scale chromatin unfolding and allele-specific effects of cancer-predisposing mutations. J Cell Biol 155, 911–921. 6. Wolffe, A. P. (1997) Histones, nucleosomes and the roles of chromatin structure in transcriptional control. Biochem Soc Trans 25, 354–358. 7. Wolffe, A. P., Khochbin, S., and Dimitrov, S. (1997) What do linker histones do in chromatin? Bioessays 19, 249–255. 8. Dou, Y., Bowen, J., Liu, Y., and Gorovsky, M. A. (2002) Phosphorylation and an ATP-dependent process increase the dynamic exchange of H1 in chromatin. J Cell Biol 158, 1161–1170. 9. Dou, Y., and Gorovsky, M. A. (2000) Phosphorylation of linker histone H1 regulates gene expression in vivo by creating a charge patch. Mol Cell 6, 225–231. 10. Dou, Y., Mizzen, C. A., Abrams, M., Allis, C. D., and Gorovsky, M. A. (1999) Phosphorylation of linker histone H1 regulates gene expression in vivo by mimicking H1 removal. Mol Cell 4, 641–647.

11. Shen, X., Yu, L., Weir, J. W., and Gorovsky, M. A. (1995) Linker histones are not essential and affect chromatin condensation in vivo. Cell 82, 47–56. 12. Takeda, D. Y., Wohlschlegel, J. A., and Dutta, A. (2001) A bipartite substrate recognition motif for cyclin-dependent kinases. J Biol Chem 276, 1993–1997. 13. Liu, P., Barkley, L. R., Day, T., Bi, X., Slater, D. M., Alexandrow, M. G., Nasheuer, H. P., and Vaziri, C. (2006) The Chk1-mediated S-phase checkpoint targets initiation factor Cdc45 via a Cdc25a/Cdk2-independent mechanism. J Biol Chem, 281(41), 30631–30644. 14. Tsukamoto, T., Hashiguchi, N., Janicki, S. M., Tumbar, T., Belmont, A. S., and Spector, D. L. (2000) Visualization of gene activity in living cells. Nat Cell Biol 2, 871–878. 15. Janicki, S. M., Tsukamoto, T., Salghetti, S. E., Tansey, W. P., Sachidanandam, R., Prasanth, K. V., Ried, T., Shav-Tal, Y., Bertrand, E., Singer, R. H., and Spector, D. L. (2004) From silencing to gene expression: real-time analysis in single cells. Cell 116, 683–698. 16. Harlow, E., and Lane, D. (1988) Antibodies: A Laboratory Manual, Cold Spring Harbor Laboratory Press. 17. Kennedy, B. K., Barbie, D. A., Classon, M., Dyson, N., and Harlow, E. (2000) Nuclear organization of DNA replication in primary mammalian cells. Genes Dev 14, 2855–2868.

Chapter 4 DNA Methyltransferase Probing of Chromatin Structure Within Populations and on Single Molecules Carolina Pardo, Scott A. Hoose, Santhi Pondugula, and Michael P. Kladde Abstract Non-invasive methods for mapping chromatin structure are necessary for creating an accurate view of genome function and dynamics in vivo. Ectopic induction of cytosine-5 DNA methyltransferases (C5 MTases) in Saccharomyces cerevisiae is a powerful technique for probing chromatin structure with minimal disruption to yeast physiology. Accessibility of MTases to their cognate sites is impaired based on the strength and span of the protein–DNA interaction to be probed. Methylated cytosines that resist chemical deamination are detected positively by the PCR-based technique of bisulfite genomic sequencing. PCR amplicons can be sequenced directly yielding an average m5C frequency or accessibility of each target site within the population, a technique termed methyltransferase accessibility protocol (MAP). More recently, the sequencing of cloned molecules in MAP for individual templates (MAPit) enables assignment of the methylation status of each target site along a continuous DNA strand from a single cell. The unique capability to score methylation at multiple sites in single molecules permits detection of inherent structural variability in chromatin. Here, MAPit analysis of the repressed and induced PHO5 promoter of budding yeast, using a C5 MTase with dinucleotide recognition specificity, reveals considerable cell-to-cell heterogeneity in chromatin structure. Substantial variation is observed in the extent to which the MTase gains entry to each of the nucleosomes positioned at PHO5, suggesting differences in their intrinsic thermodynamic stability in vivo. MAPit should be readily adaptable to the analysis of chromatin structure and non-histone protein–DNA interactions in a variety of model systems. Key words: Chromatin, DNA methyltransferases, footprinting, nucleosomes, single-molecule probing, transcription, yeast.

1. Introduction Site-specific protein–DNA interactions and chromatin structure have emerged as central aspects of chromosome biology in all eukaryotic organisms. Efficient techniques are needed to precisely Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_4 Springerprotocols.com

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locate these interactions and thus unravel the complexities of genome function in living cells. Previously, we devised a method for interrogating chromosome structure in bulk populations of cells, termed methyltransferase accessibility protocol (MAP) (1–6). As conveyed by its acronym, MAP uses a DNA methyltransferase (MTase) as the probing agent of DNA accessibility in chromatin. In the majority of our studies, MTases have been expressed in vivo under the control of an ectopic regulatory system, which minimizes disruption to physiology and maintains the true native state of chromatin. In vivo expression of MTases circumvents cell disruption in that the only required cofactor, the methyl donor S-adenosyl methionine, is abundant in cells. However, purified enzymes have also been used as probes in binding reactions containing purified factors or nuclear extracts, reconstituted chromatin and intact nuclei, which are supplemented with S-adenosyl methionine (2, 6–12). Best results are achieved when multiple doses of enzyme or times of methylation are employed. The association of histones and site-specific DNA-binding factors with DNA hinders MTase binding based on the affinity of the complexes (1–6). After probing the samples of interest, protein–DNA interactions are located by determining the pattern of accessible and inaccessible MTase target sites in each purified DNA sample. The premise is identical to probing with conventional nucleases, except that ‘footprints’ arise from protection against DNA methylation rather than scission. Cytosine-5 (C5) MTases with dinucleotide specificity are the most useful probes (13–15), as the frequency of their target sites in natural sequences yields sufficient resolution to footprint most nucleosomes and many protein–DNA interactions (1–12). Moreover, detection of 5-methylcytosine (m5C) in DNA is performed at nucleotide resolution using the PCR-based technique of bisulfite genomic sequencing (16, 17). In this procedure (Fig. 4.1), cytosines that are unmethylated because of either limiting MTase activity or occlusion of the probe by DNA-associated factors are chemically converted to uracil by bisulfite ion. In contrast, m5C is retained in the sample because it is resistant to hydrolytic chemical conversion. During PCR of the region of interest, m5C is amplified as C, whereas U base pairs with A in the first PCR cycle and is subsequently replaced by T (i.e., there is a net transition of unmethylated C to T). Therefore, PCR products resulting from amplification of bisulfite-converted DNA have a mixture of C and T at each target site for the ectopic MTase probe and/or endogenous MTases. The method for assaying the relative amounts of C and T and thus the frequency of methylation at each MTase target site depends largely on the goals of the experiment. To obtain the populationaveraged positions of nucleosomes and fractional occupancies of specific elements in the region of interest, the PCR product amplified

Single-Molecule Probing of Chromatin Structure with DNA MTases

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Fig. 4.1. Detection of methylated cytosine bases by bisulfite genomic sequencing. M.CviPI sites (GC) are marked by filled circles. A DNA-bound protein is shown as a dashed partial ellipse. Unmodified cytosines are deaminated to uracil. Cytosines that have been methylated (m) resist chemical deamination. Note that the a0 and b0 DNA strands are no longer complementary following deamination, and thus can and must be amplified separately with dedicated primer pairs (a1/a2 or b1/b2). PCR products are then either directly sequenced using ddGTP (a1 or b1 primer) or ddCTP (a2 or b2 primer) terminators for population-ensemble MAP analysis or cloned and subjected to conventional sequencing in single-molecule MAPit analysis.

from bisulfite-converted DNA is directly subjected to thermal cycle sequencing with an end-labeled primer. In the absence of added dGTP, efficient incorporation of 20 ,30 -ddGTP terminator across from C residues in the PCR product template provides a positive display of C that reflects in vivo m5C. Therefore, on a sequencing gel, band intensities are directly proportional to the percentage of molecules that were methylated in the original sample DNA, that is, prior to bisulfite treatment. Signal intensities of each band on the gel are compared with their respective bands generated from an appropriate control sample (e.g., methylated protein–free DNA) to reveal the locations of nucleosomes and DNA-bound site-specific factors (1–10). In general, smaller regions of protection against methylation correspond to non-histone regulatory proteins and larger ones to positioned nucleosomes. However, as nucleosomes ‘breathe’ by a process termed site exposure (18–21), methylation may increase at nucleosome entry–exit sites and thereby shorten nucleosomal footprints (1, 2, 22). A universal limitation of this strategy and all populationensemble techniques for mapping chromatin structure is that only the first event proximal to the location of primer or probe hybridization is detected on each molecule in the probed sample (e.g., first C encountered by the extending primer). In other words, multiple ‘hits’ cannot be detected on the same molecule.

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In practice, excessive methylation, or cutting in the case of nuclease probing, is avoided by achieving conditions of single-hit kinetics. According to Poisson statistics, criteria for single-hit kinetics are met when methylation is random and each molecule contains, on average, 0.5 modifications. Under these conditions, at least 60% of the molecules are not methylated, which collectively yield a full-length product band at the top of each lane (1, 2). In cases where there is reason to expect modification of many sites on individual molecules, pyrosequencing offers a more quantitative approach for determining m5C frequencies in a PCR amplicon (23, 24). An examination of sequencing technologies is beyond the scope of this chapter. Many situations are likely to occur in chromosome biology in which the pattern of probe accessibility may deviate from randomness and thus adherence to Poisson single-hit kinetics is precluded. For example, after shifting a culture of budding yeast from glucose to glucoseþ galactose, a binary mode of induction of GAL1 occurs in which a fraction of the cells activate transcription and some do not (25). Considering the close correlation of chromatin remodeling and gene activation at the GAL1 locus, it is possible, if not probable, that the chromatin architecture under these conditions is also bimodal. While this hypothesis needs to be tested experimentally, it illustrates that a population-averaged view of chromatin structure would obscure differences between individual gene copies. Therefore, the MAP assay has been modified to capture the inherent heterogeneity of chromatin structure between single molecules comprising a population, a procedure we refer to as MAP for individual templates (MAPit) (4, 6, 11, 12). In this protocol, individual molecules from PCR amplicons derived from bisulfite-treated DNA are first cloned and then sequenced, as originally described for bisulfite genomic sequencing (16, 17). Thus, the pattern of methylation in each clone represents the chromatin state of a continuous region from a single cell. Warnecke et al. (26) should be consulted for help in avoiding and resolving artifacts associated with bisulfite genomic sequencing. A key advantage of MAPit is that the methylation status of each site is determined independently from other sites on individual DNA strands, enabling m5C at multiple sites on the same molecule to be scored. This eliminates concerns regarding single-hit kinetic levels of methylation and absolute rather than relative methylation frequencies are obtained. Therefore, analysis of sets of cloned molecules yields a quantitative view of chromatin structure and simultaneously discerns variation between molecules from different cells. MAPit has been used to demonstrate heterogeneity in chromatin remodeling at the induced PHO5 promoter of budding yeast (4). Variable numbers of nucleosomes were disrupted in individual cells with preferential targeting of nucleosomes in the vicinity of promoter-associated activators (Fig. 4.2). Moreover,


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Fig. 4.2. Single-molecule MAPit analysis of S. cerevisiae PHO5 promoter chromatin. The PHO5 gene, coding for the major yeast acid phosphatase, is induced when environmental phosphate becomes limiting. Under these conditions, the transactivator Pho4 is imported into the nucleus and binds to upstream activation sequence (UAS) elements UASp1 and UASp2, leading to remodeling of several positioned nucleosomes, designated 5 to þ1 (only 3 to þ1 are shown). In this experiment, chromatin structure at the PHO5 promoter was probed in wild-type (PHO4) or PHO4 deletion (pho4) strains containing M.CviPI on integrated plasmid pSH1052. Strains were grown in CSM-glutamine phosphate-free media for 14 h, which achieves high-level induction of the PHO5 promoter. After collecting a 0 min time point, the cells were maintained in phosphate-free medium and supplemented with 100 nM 17b-estradiol (E2)

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frequent disruption of adjacent nucleosomes suggested a mechanism for chromatin remodeling that involves spreading. Similarly, single-molecule MTase footprinting of isolated mammalian nuclei has been used to show sequential co-occupancy by site-specific activators at an induced promoter (12). The utility of such single-molecule studies is evident and warrants their extension to additional model systems. Many common yeast plasmid vectors can be used for cloning the appropriate coding and regulatory sequences for expression of MTases in vivo in yeast. Such vectors should include an antibiotic resistance gene (such as the -lactamase gene, bla, conferring ampr) for Escherichia coli cloning purposes and an appropriate yeast selectable marker (such as LYS2) for selection of yeast transformants. We have constructed a series of yeast integrating plasmids (YIp) for use in MTase expression, comprising estrogen-, doxycycline-, and galactose-inducible systems, with a variety of C5 MTases including M.SssI (m5CG) (13), M.CviPI (Gm5C) (14) and M.HhaI (Gm5CGC) (27). In addition, fusions of M.CviPI and M.HhaI have been constructed which confer additional functional and regulatory characteristics, such as a Pho4M.CviPI fusion expressed from the endogenous PHO4 promoter (28). The Materials and Methods sections address the preparation and execution of an experiment using pSH1052, a LYS2-marked HO-targeted YIp which codes for all necessary ectopic components of an estrogen-inducible free M.CviPI expression system. Herein we have focused on MAPit singlemolecule probing of protein–DNA interactions. Previous chapters can be consulted for a full discussion of MAP analysis of populations (3, 5, 6).

Fig 4.2. (continued) for 80 or 120 min at 30C to induce M.CviPI synthesis in vivo. DNA was then rapidly isolated and treated with bisulfite. The deaminated DNA was amplified with b1/b2 primers, and PCR products were cloned and sequenced as described in the text. Independent clones are shown by gray lines that indicate the methylation status of each M.CviPI target site, numbered 1 through 33 (filled circles, Gm5C; open circles, GC). The histograms below each set of single molecules show the cumulative m5C at each GC site. In the pho4 strain, which cannot activate PHO5 transcription, note that m5C accumulates at the highest rate in the accessible-histone-free UASp1 region and also in the edges of nucleosomes 3, 1, and þ1. By contrast, no m5C is observed in pho4 cells at nucleosome 2, even at two GC sites located very close to its downstream edge, suggesting that this nucleosome has the highest thermodynamic stability (18–21). In the PHO4 strain, each of the positioned nucleosomes has been disrupted to different extents as evidenced by the different levels of m5C present at sites located at their pseudodyad centers. Other symbols are full and partial ellipses, positioned nucleosomes; filled rectangles, UASp1, UASp2 and TATA, as indicated; bent arrow, major transcription start site; partial rectangle, partial PHO5 coding region. All features are drawn to scale, except that the GC sites are oversized by twofold.


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2. Materials 2.1. Equipment

1. Micropipets. 2. 30 and 37C incubators. 3. Incubator shakers. 4. Water baths. 5. Hot-stir plate. 6. Heat block. 7. Multi-tube vortexer. 8. Benchtop vortexer. 9. Refrigerated superspeed centrifuge. 10. Swinging-bucket rotor with tube holders. 11. Microcentrifuge. 12. Spectrophotometer. 13. Thermocycler with a dual 30/48 Alpha UnitTM Block Assembly with heated lids (MJ Research PTC-200 or similar). 14. pH meter. 15. Vac-Man vacuum manifold (Promega). 16. Mini-gel boxes. 17. UV transilluminator. 18. ABI 3130 DNA sequencer. 19. Comparable equipment may be substituted.

2.2. Sterile Consumables

1. 1250 mL micropipet tips. 2. 200 mL micropipet tips. 3. 50 mL conical centrifuge tubes. 4. 15 mL conical centrifuge tubes. 5. 1.7 mL microcentrifuge tubes. 6. 1.5 mL screw-cap microcentrifuge tubes. 7. 0.65 mL microcentrifuge tubes. 8. 0.5 mL thin-wall PCR tubes. 9. 1.2 mL cryogenic storage vials. 10. 10 mL pipets. 11. 5 mL pipets. 12. 0.45 mm bottle-top filters. 13. 96-MicroWell plates. 14. AirPoreTM Tape Sheets (Qiagen 19571). 15. Aluminum tape.

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16. Parafilm1. 17. Wood applicator sticks (Puritan 807). 18. 1 mL slip-tip syringe barrels. 19. 0.45 mm syringe filters. 20. Non-sterile disposable cuvettes. 21. Comparable consumables may be substituted for any of the above items. Unless otherwise specified, solutions are prepared using water that has been purified to a resistance of 18.2 M -cm (dH2O). Caution: A number of hazardous compounds are used in these protocols, including but not limited to 17-estradiol, chloroform, phenol, sodium metabisulfite, sodium hydroxide, hydroquinone, and guanidine thiocyanate. Personnel should be trained in the use of the materials listed herein and should take appropriate precautions as set forth by the manufacturer and institutional policy. 2.3. Construction of MTase-Expressing Yeast Strains

2.3.1. Bacterial Strain Transformation, Storage, and Plasmid Isolation

1. Chemically competent or electro-competent E. coli cells that are deficient for restriction of methylated DNA (Mrr Hsd Mcr). We have used competent DH10BTM cells (Invitrogen) with success. 2. Luria–Bertani Broth þ ampicillin medium (LB þ amp): 25 g/ mL Miller Luria–Bertani Broth (DifcoTM 244610), 15 g/L bacteriological agar (US Biological) for plates. Autoclave to sterilize. Add sterile-filtered 100 mg/mL ampicillin sodium salt in dH2O to a final concentration of 100 mg/mL after autoclaving and cooling the medium. LB liquid medium may be stored at room temperature (RT); plates containing amp must be stored at 4C and should not be prepared more than 1 month prior to use. Add ampicillin to liquid medium immediately before use. Store amp stock at 20C for up to 6 months. 3. pSH1052. 4. Sterile 50% glycerol: dilute with dH2O. Filter sterilize. Store at 4C. 5. Materials for plasmid miniprep procedure. 6. Sterile 0.1 TE, pH 8.0: 1 mM Tris–HCl, pH 8.0 (JT Baker 4109-06), 0.1 mM EDTA, pH 8.0. Autoclave to sterilize.

2.3.2. Yeast Transformation, Selection, and Storage

1. pSH1052. 2. R.AatII (New England Biolabs R0117S) and R.AscI (New England Biolabs R0558S), with accompanying 10 reaction buffers and supplements. 3. A yeast strain with a lys2 auxotrophic marker. 4. Yeast extract peptone dextrose medium (YPD): 20 g/L dextrose, 20 g/L peptone (BD BactoTM), 10 g/L yeast extract


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(BD BactoTM), 20 g/L bacteriological agar (US Biological A0930) for plates. Autoclave 30 min to sterilize. 5. Sterile dH2O. Autoclave to sterilize. 6. Sterile 0.1 M lithium acetate (LiOAc). Autoclave to sterilize. 7. Sterile 50% (w/v) poly(ethylene glycol) average molecular weight 3350 (PEG4000): dissolve PEG4000 in dH2O. Filter sterilize. 8. Sterile 1.0 M LiOAc. Autoclave to sterilize. 9. 5 mg/mL carrier DNA: dissolve 0.05 g salmon testes DNA sodium salt (Sigma D1626) in 10 mL sterile 1 TE, pH 8.0 (10 mM Tris–HCl, pH 8.0, 1 mM EDTA, pH 8.0) with vigorous stirring or agitation. Aliquot 200 mL into 0.65 mL tubes and store at 20C. The aliquots must be placed in a boiling water bath for 5 min and quick chilled on ice prior to first use. 10. Complete supplement mixture minus lysine selective medium (CSM-Lys): 20 g/L dextrose, 6.7 g/L yeast nitrogen base (US Biological), 0.74 g/L CSM-Lys drop-out mix (BIO101 4510-622), 20 g/L bacteriological agar (US Biological) for plates. Autoclave 30 min to sterilize. 11. Sterile 50% glycerol. 2.3.3. Screening for Yeast Strains with Functional M.CviPI Expression Systems

1. YPD. 2. 100 mM 17-estradiol (E2): dissolve 13.62 mg E2 (Sigma E8875) in 10 mL 95% ethanol to yield a 5 mM solution. Mix 1470 mL 95% EtOH with 30 mL 5 mM E2 to dilute to 100 mM. Store at 20C.

2.3.3.1. Rapid Screening for M.CviPI Activity by Culture Density 2.3.3.2. Total Yeast DNA Isolation

1. 1 TE, pH 8.0: 10 mM Tris–HCl, pH 8.0, 1 mM EDTA, pH 8.0. 2. 425–600 mm diameter acid-washed glass beads (Sigma G8772). 3. ‘Smash buffer’: 10 mM Tris–HCl, pH 8.0 (JT Baker 410906), 1 mM EDTA, pH 8.0, 2% Triton X-100 (JT Baker X19805), 1% sodium dodecyl sulfate, 100 mM NaCl. 4. 24:1 chloroform:isoamyl alcohol (24:1 CHCl3:IAA): add 20.8 mL IAA to a new 500 mL bottle of CHCl3 and mix thoroughly. 5. Equilibrated phenol: to a 400 mL bottle of saturated phenol add the entire contents of the accompanying small bottle of equilibration buffer to raise the pH from 6.6 to 7.9. Mix the two phases thoroughly and allow them to separate. The aqueous buffer will lie on top of the equilibrated phenol. Store at 4C. 6. 10 M ammonium acetate (NH4OAc, EMD AX1220-1). 7. Isopropanol (Fisher A416-500).

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8. 7:1 EtOH:1 TE, pH 8.0: aliquot 13 mL 1 TE, pH 8.0 to a 50 mL centrifuge tube and qs to 50 mL with 95% EtOH and mix thoroughly. The final EtOH concentration is 70% (v/v). 9. 0.1 TE, pH 8.0. 2.3.3.3. R.McrBC Digestion of Isolated DNA

1. R.McrBC (New England Biolabs M0272S) with accompanying 10 NEB2 buffer and 100 BSA/GTP supplements. 2. 50% glycerol. 3. Electrophoretic gel: 1% (w/v) agarose (BioRad 162-0138), 1 Tris–acetate–EDTA (TAE), 0.5 mg/mL ethidium bromide (EtBr). 4. Electrophoretic buffer: 1 TAE, 0.5 mg/mL EtBr. 5. 10 loading buffer for agarose gel electrophoresis.

2.4. MTase Mapping of DNA–Protein Interactions

2.4.1. Growth of Strain(s) Under Experimental Conditions 2.4.2. Bisulfite Deamination of Isolated Yeast DNA

See above for media formulation appropriate for the experiment to be performed.

1. 1 TE. 2. 20 mL glass scintillation vial. 3. Wizard1 Minicolumns (Promega A7211). 4. 3 mL Luer-lock syringe barrels. 5. Unopened vial containing 5 g aliquot of sodium metabisulfite (Sigma 255556 or similar) (see Note 1). 6. 3 mg/mL sheared calf thymus DNA (sh.CTDNA): dilute from stock of calf thymus DNA (Sigma D8661; lot concentration is given on packaging). Shear by passing 15 times through a 21-gauge needle/syringe. 7. Degassed distilled water (dg.dH2O): boil >200 mL dH2O for 20 min in a glass beaker. After 20 min, carefully pour the boiling water into a 125 mL bottle until it is completely full (above the lip) and screw the cap on tightly. Cool overnight on the benchtop. 8. 3 N NaOH: make fresh shortly before use. Weigh out approximately 0.4 g NaOH pellets into a 15 mL centrifuge tube. Add the appropriate volume of dg.dH2O (8.333 g of NaOH = mL of dg.dH2O). Dissolve the pellets by gently rocking the tube rather than by vortexing, in order to minimize aeration. 9. 100 mM hydroquinone (HQ): make fresh shortly before use. Weigh out approximately 0.04 g HQ (Sigma) into a 15 mL centrifuge tube. Add the appropriate volume of dg.dH2O


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(90.827 g of HQ = mL of dg.dH2O). Dissolve the HQ by gently rocking the tube rather than by vortexing, in order to minimize aeration. 10. 0.5 M EDTA, pH 8.0. 11. Sample denaturation buffer (SDB): make fresh shortly before use. Mix buffer components in the following ratios (10 mL total volume per sample to be deaminated, plus a little excess): 5.8 mL dg.dH2O, 3.0 mL 3 N NaOH, 0.7 mL 3 mg/mL sh.CTDNA, 0.5 mL 0.5 M EDTA, pH 8.0. Use the 3 N NaOH made in step 8. 12. Saturated sodium metabisulfite solution (SMBS): make fresh shortly before use. a. Put a small stir bar in a clean 20 mL scintillation vial and place the vial in the center of a stir plate next to the pH meter. b. Take all reagents to be used (100 mM HQ, an unopened 5 g vial of sodium metabisulfite, dg.dH2O, and 3 N NaOH) to the bench. c. Take P100 and P1000 micropipets, tips, and a 5 mL pipet and pipet-aid to the bench. d. Calibrate and prepare the pH meter. e. Prepare appropriate volumes of reagents prior to beginning preparation of SMBS as follows. f. Draw 7 mL dg.dH2O into the pipet and set aside. g. Draw 1 mL 3 N NaOH using the P1000 and set aside. h. Draw 100 mL 100 mM HQ using the P100 and add to the scintillation vial. i. Open the 5 g vial of sodium metabisulfite and quickly dump the entire contents into the scintillation vial. j. Immediately and in rapid succession add the 7 mL dg.dH2O, begin stirring, and add the 1 mL 3 N NaOH. k. Begin taking the pH of the solution. Adjust the pH to 5.0 using 3 N NaOH (it usually requires an additional 200–300 mL). Record the final pH. l. Since this is designed to be a saturated solution at RT, there will probably be some undissolved sodium metabisulfite remaining. 13. 0.1 TE. 14. Desulfonation solution: make fresh shortly before use. Mix buffer components in the following ratios (8 mL total volume per sample to be desulfonated, plus a little excess): 7.0 mL 3 N NaOH, 1.0 mL 3 mg/mL sh.CTDNA. Use the 3 N NaOH made in step 8. 15. Wizard1 PCR Preps DNA Purification Resin (Promega A7181). 16. 80% (v/v) isopropanol. 17. 10 M NH4OAc.

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18. 95% EtOH. 19. 7:1 EtOH:1 TE, pH 8.0 (70% EtOH). 2.4.3. PCR Amplification of Deaminated Template DNA

1. JumpStartTM Taq DNA polymerase (Sigma D4184) or equivalent with accompanying 10 reaction buffer and 25 mM MgCl2. 2. 2.5 mM deoxynucleotide triphosphates (dNTPs): the solution is prepared by mixing 360 mL dH2O with 10 mL of each 100 mM stock of dNTP (Amersham Biosciences) to yield a 2.5 mM solution with respect to individual nucleotides. Store at 20C. 3. 20 mM DNA deamination PCR primers (b1/b2 or a1/a2 pair) (see Note 2). 4. Electrophoretic gels: 1% (w/v) agarose, 1 Tris–acetate–EDTA (TAE), 0.5 mg/mL ethidium bromide (EtBr). 5. Electrophoretic buffer: 1 TAE, 0.5 mg/mL EtBr. 6. 10 loading buffer for agarose gel electrophoresis. 7. Wizard1 PCR Preps DNA Purification System (Promega A7170).

2.4.4. Cloning PCR Amplicons Amplified from Deaminated Template DNA

1. Chemically or electro-competent strain of E. coli cells. We have used competent DH10BTM cells (Invitrogen) with success. 2. Appropriate cloning vector. We typically use pBlueScript SK() (Stratagene) that contains ampicillin resistance marker and uses blue/white screening (see Notes 3 and 4). 3. Appropriate restriction endonucleases, with accompanying 10 reaction buffers and supplements for cloning in the selected vector. 4. T4 DNA ligase (New England Biolabs). 5. LB þ amp plates.

2.4.5. SequencingCloned DNA Molecules

1. LB þ amp liquid media. 2. 50% glycerol.

3. Methods 3.1. Construction of MTase-Expressing Yeast Strains

1. Transform a chemically or electro-competent strain of E. coli that is deficient for restriction of methylated DNA (Mrr Hsd Mcr) with pSH1052. Select transformants on a LB þ ampicillin plate with incubation for approx. 12–16 h at 37C (see Note 5).

3.1.1. Bacterial Strain Transformation, Storage, and Plasmid Isolation

2. Inoculate approx. 5–7 mL LB þ amp with a single colony. Incubate the cultures for 8–14 h at 37C with shaking at 250 rpm (see Note 6).


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3. Prepare a frozen stock of the transformed strain by mixing 350 mL sterile-filtered 50% (v/v) glycerol with 700 mL of culture in a cryogenic storage vial or 1.7 mL microcentrifuge tube. Store at 80C. Future cultures can be prepared by directly inoculating media with a small amount of cells that are sterilely scraped from the surface of the frozen stock. If difficulty is encountered in obtaining unrearranged plasmid, then inoculate cultures with single colonies from an LB þ amp plate. 4. Isolate plasmid DNA using any of the many common techniques or commercially available kits. Dissolve/elute the DNA in 30–50 mL 0.1 TE. Store the plasmid solution at 20C.

3.1.2. Yeast Transformation, Selection, and Storage

1. Digest approx. 100 ng pSH1052 with R.AatII and R.AscI in a double digest according to the directions of enzyme supplier (see Note 7). 2. Yeast cells are prepared for transformation by early log-phase culture growth for approx. 3–6 h in YPD at 30C with shaking at 250 rpm. For our purposes, early log is defined as logphase growth at an optical density at 600 nm (OD600) of 1.0. About 5 U of cells are used per transformation (1.0 U = 1.0 mL of culture at an OD600 of 1.0 = 2.0 mL of culture at an OD600 of 0.5, and so on). A negative control transformation with no plasmid DNA, if performed, counts as one transformation (see Note 8). 3. Pellet the cells 5 min at 3300 g at 25 to 30C in a centrifuge. Aseptically decant the media. Wash the cells by resuspending in 1.0 mL sterile distilled water (st.dH2O), transferring to a 1.7 mL microcentrifuge tube and vortexing briefly. Centrifuge briefly at 16,000 g in a microcentrifuge to pellet the cells (tap spin). Draw off and discard the wash. 4. Resuspend the cells in 100 mL 0.1 M sterile LiOAc per transformation; if multiple transformations or negative controls are to be done, aliquot equal volumes to new 1.7 mL microcentrifuge tubes. Tap spin to pellet the cells. Draw off and discard the LiOAc. 5. Resuspend the cells in the following solutions in the order listed, vortexing well after the addition of each: 240 mL 50% (w/v) sterile PEG4000 (see Note 9), 36 mL 1.0 M sterile LiOAc, 5 mL 5 mg/mL carrier DNA, and 75 mL approx. 100 ng pSH1052 þ st.dH2O (i.e., plasmid digest, qs to 75 mL with st.dH2O), for a total of 356 mL. 6. Incubate 30 min in a 30C water bath. Transfer to a 42C water bath, incubate an additional 45 min. 7. Tap spin. Draw off and discard the transformation solution. Wash the cells by resuspending in approx. 500 mL st.dH2O

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and vortexing briefly. Tap spin. Draw off and discard the wash. Resuspend in 100 mL st.dH2O and plate on CSM-Lys. 8. Incubate the plates at 30C. Colony growth generally requires 2–3 days. Upon obtaining colonies, purify four independent transformants by picking a single colony and streaking the cells out to single colonies on quadrants of a second CSM-Lys plate. 9. When distinct single colonies arise, inoculate approx. 5 mL YPD with a single colony. Inoculate multiple cultures from different quadrants (i.e., independent transformants) to increase the odds of obtaining a strain that will express active M.CviPI. Incubate at 30C with shaking at 250 rpm for 12–16 h (see Note 10). 10. Prepare frozen stocks of the clonal transformed strains by mixing 350 mL sterile-filtered 50% glycerol with 700 mL culture in a cryogenic storage vial or 1.7 mL microcentrifuge tube. Store at 80C. The strain can then be prepared for future applications by patching a small amount of cells sterilely scraped from the surface of the frozen stock onto rich or selective media, incubating the plate at 30C, and inoculating liquid media from the patch once it has grown up (see Note 11). 3.1.3. Screening for Yeast Strains with Functional M.CviPI Expression Systems (see Note 12) 3.1.3.1. Rapid Screening for M.CviPI ACTIVITY by Culture Density

1. From a plate, inoculate approx. 5 mL YPD with the strains to be screened to an OD600 of approx. 0.5. Incubate at 30C with shaking at 250 rpm for several minutes to achieve uniformity. 2. Determine the A600 and reseed a 10.0 mL YPD culture at an OD600 = 0.1 in a 50 -mL conical centrifuge tube. Incubate as above to achieve uniformity. From this culture aliquot 5.0 mL into a new tube. Add 5.0 mL 100 mM 17-estradiol (E2) to this 5 mL culture to a final concentration of 100 nM and incubate both tubes 12–16 h (overnight) at 30C with shaking at 250 rpm. 3. At the conclusion of the incubation, compare the densities of the E2þ and E2 cultures. Strains with a functional M.CviPI expression system are consistently significantly less dense in the E2þ vs. the E2 cultures (an OD600 1.0–2.0 vs. OD600 4.0–7.0 is typical for our strains) (see Note 13).

3.1.3.2. Total Yeast DNA Isolation

1. Grow a culture under inducing conditions for approx. 12–16 h (the E2þ culture from Section 3.1.3.1 can be used). 2. Pellet cells 5 min at 3300 g at 4–30C. Decant the supernatant, resuspend cells in 700 mL 1 TE, pH 8.0, and transfer to a 1.5 mL screw-cap microcentrifuge tube (see Note 14).


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3. Add approx. 0.3 g acid-washed 400–625 mm diameter glass beads. Pellet the cells by tap spinning. Draw off and discard the supernatant and add the following in the order indicated: 200 mL 1 TE, pH 8.0, 200 mL smash buffer, 200 mL 24:1 CHCl3:IAA, and 200 mL equilibrated phenol (see Note 15). 4. Cap the tubes tightly and vortex at maximum speed for 8 min. Centrifuge 5 min at 16,000 g at RT. Transfer the upper (aqueous) phase to a 1.7 mL microcentrifuge tube, being careful not to transfer any of the debris at the phase interface. Add 200 mL 10 M NH4OAc to the supernatant and vortex. Incubate the tube on ice 2 h to overnight. 5. Centrifuge 5 min at 16,000 g at RT. Transfer the supernatant to a new 1.7 mL tube. Add 600 mL isopropanol to the supernatant and invert the tube multiple times until mixed. Incubate at RT for 5 min. 6. Centrifuge 5 min at 16,000 g at RT to pellet the nucleic acids. Draw off and discard the supernatant. Tap spin, then carefully draw off and discard the remainder of the supernatant. 7. Add 400 mL 7:1 EtOH:1 TE, pH 8.0 (70% EtOH) and vortex briefly to wash the pellet and tube. Centrifuge 2 min at 16,000 g at RT. Carefully draw off and discard the wash. Tap spin, then carefully draw off and discard the remainder of the supernatant. Dry the pellet briefly by centrifuging the open tube 1 min at 400 g in an open microcentrifuge rotor. Add 45 mL 0.1 TE to the nucleic acid pellet. Incubate 1–24 h at 4C and then mix the DNA solution gently by pipetting to ensure homogeneity. Store the DNA at 20C. 3.1.3.3. R.McrBC Digestion of Isolated DNA

1. Make a reaction mixture of the following composition: 13.2 mL dH2O, 2.4 mL 10 NEB2, (supplied by manufacturer), 2.4 mL 10 BSA (supplied as 100 by manufacturer), 2.4 mL 10 GTP (supplied as 100 by manufacturer), and 2.4 mL isolated DNA solution, for a total of 22.8 mL (see Note 16). 2. Aliquot 9.5 mL to two 1.7 mL microcentrifuge tubes. To one, add 0.5 mL (5 U) R.McrBC. To the other, add 0.5 mL 50% (v/v) glycerol. Incubate both tubes for 1–4 h at 37C. 3. Tap spin both tubes. Add 1.1 mL 10 loading buffer to each and mix thoroughly. Load 10 mL of each sample in adjacent lanes on a 1% agarose–TAE–EtBr mini-gel and electrophorese 30 min at 100 V. Visualize by UV transillumination (see Note 17).

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3.2. MTase Mapping of DNA–Protein Interactions

3.2.1. Growth of Strain(s) Under Experimental Conditions

3.2.2. Bisulfite Deamination of Isolated Yeast DNA

The nature of the experiment will determine the details of this portion of the protocol. Variations may include growth of a single MTase-expressing strain under several different conditions or growth of MTase-expressing experimental and control strains under a single condition. Depending upon the method of regulation, induction of the MTase may be accomplished at high levels in a short pulse at the desired time, at lower levels over a longer period, or constitutively as in the case of a native DNA-binding factor-MTase fusion expressed from an endogenous constitutive promoter. At the completion of the experiment, total yeast DNA is rapidly isolated and subsequently subjected to selective deamination of unmethylated cytosines by bisulfite ion. 1. Prepare the degassed water the evening prior to execution of subsequent steps. 2. Prepare the 3 N NaOH and 100 mM hydroquinone solutions. Prepare the sample denaturation buffer (SDB). Aliquot 10 mL of SDB to a 0.65 mL microfuge tube for each sample. Add 20 mL yeast DNA solution to the SDB and mix by pipetting. The samples should be left at RT while preparing subsequent reagents. 3. Prepare the sodium metabisulfite solution (SMBS) and prewarm it to 50C. It is not necessary to confirm the temperature prior to use. 4. While the SMBS is warming, denature the samples in a thermocycler for 5 min at 98C. a. At about the 4 min point, bring the SMBS to the thermocycler, uncap it and stir briefly. b. At 5 min, open the thermocycler while maintaining block temperature at 98C and, working rapidly, open the first tube in the block, add 200 mL SMBS, cap and remove the tube, vortex it immediately and place it in a rack or float. c. Proceed likewise for all the samples, using a new tip for each sample. d. After all samples have been prepared, incubate them and the remaining SMBS at 50C for 6 h in the dark (see Note 18). 5. During the incubation, label two sets of 1.7 mL microcentrifuge tubes. Attach one labeled Promega minicolumn to a 3 mL syringe barrel for each sample. 6. Near the conclusion of the incubation, attach the minicolumn assemblies to the vacuum manifold. Heat the 0.1 TE to 95C. Prepare the desulfonation solution (DSS) and aliquot 8 mL to each 1.7 mL microcentrifuge tube of the first labeled set from step 5. Read and record the final pH of the unused SMBS (see Note 19).


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7. At the conclusion of the incubation, transfer the sample solutions to the second set of 1.7 mL microcentrifuge tubes from step 5. a. Add 1 mL Promega Wizard1 PCR Preps resin to each sample and vortex. b. Transfer the resin to the syringe barrels on the vacuum manifold. c. Apply the vacuum to draw the resin into the minicolumns. d. Close individual stopcocks as the syringe barrels empty completely. e. Reopen the stopcocks, add 2 mL 80% (v/v) isopropanol to each syringe barrel, and apply the vacuum to draw the wash through the columns. f. Repeat the wash with 1 mL 80% (v/v) isopropanol. g. After the washes have completely passed through all columns, reopen the stopcocks and apply the vacuum for 30 s to begin drying them. h. Detach the minicolumns from the syringe barrels and press them into 1.7 mL microcentrifuge tubes (use a set of reusable tubes with lids removed that are designated for this purpose, not labeled tubes from step 5). i. Centrifuge at 16,000 g for 2 min to remove residual wash. 8. Transfer the minicolumns to the first set of labeled tubes. Add 52 mL 0.1 TE (preheated to 95C) to each minicolumn and let sit at RT for 5 min. Centrifuge for 20 s at 16,000 g to elute the DNA. Mix the eluate and DSS by vortexing. Tap spin and incubate in a 37C water bath for 15 min (see Note 20). 9. Add 18 mL 10 M NH4OAc and mix by pipetting. Add 200 mL 95% EtOH and vortex. Incubate overnight at 20C. 10. Centrifuge the tubes at 16,000 g for 20 min at RT to pellet the DNA. Draw off and discard the supernatant. Add 400 mL 7:1 EtOH:1 TE, pH 8.0 (70% EtOH), and vortex to wash. Centrifuge at 16,000 g for 2 min, carefully draw off and discard the supernatant, centrifuge again and carefully draw off and discard the remainder of the wash (see Note 21). 11. Dry the pellet briefly by centrifuging the open tube 1 min at 400 g in an open microcentrifuge rotor. Before resuspending the pellet, check to be sure there is no residual EtOH wash left in the tube. Resuspend/dissolve the pellet in 25 mL 0.1 TE. Incubate 1–24 h at 4C. Mix by pipetting and centrifuge prior to use. Store the deaminated DNA at 20C.

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3.2.3. PCR Amplification of Deaminated DNA Template

1. Use 2 –4 mL deaminated DNA as template in the following 50 mL PCR reaction: dH2O to volume, 1 Sigma JumpStartTM PCR buffer, 2.25 mM MgCl2, 0.2 mM dNTPs, 0.8 mM b1 (or a1) primer, 0.8 mM b2 (or a2) primer, 1.25 U Sigma JumpStartTM Taq DNA polymerase. Thermocycle as follows: 1 cycle of 5 min at 94C, 30 cycles of 45 s at 94C, 45 s at 5C below the calculated Tm of the primers, and 30 cycles of 1 min/kb at 72C, and 1 cycle of 5 min at 72C (see Notes 22 and 23). 2. Check for successful amplification of PCR product by electrophoresis of 1 mL of the unpurified reaction on a 1% agarose–TAE–EtBr mini-gel for 30 min at 100 V. Visualize by UV transillumination. The PCR product is purified with Promega Wizard1 PCR Preps DNA Purification System according to manufacturer’s directions. Elute the PCR product from the minicolumn in 30 mL st.dH2O. Store at 20C. 3. In preparation for cloning of the individual templates, quantify the relative product concentration by electrophoresing 2 mL of each product to be cloned on a 1% agarose–TAE–EtBr mini-gel for 30 min at 100 V followed by UV imaging. Obtain a linear digital image of the gel (i.e., not saturated due to overexposure) under UV transillumination. Using image analysis software, calculate the relative intensities of the product bands. Store samples at 20C (see Note 24).

3.2.4. Cloning PCR Amplicons Amplified from Deaminated Template DNA

1. Digest approx. 100 ng PCR product (insert) with the selected restriction endonucleases according to the directions of the enzyme supplier. We have used HindIII and XhoI (New England Biolabs) with good results. 2. Ligate the digested inserts to 30–50 ng of cloning vector (digested with the same set of restriction endonucleases as the insert) at 1:1–1:3 molar ratio of vector to insert. Ligate the insert and vector at room temperature for 1–3 h in a ligation reaction containing 1 T4 ligation buffer and 200 U T4 DNA ligase according to the manufacturer’s recommendations (see Note 25). 3. Transform chemically or electro-competent of E. coli strain. Select transformants on LB plates with the corresponding selection for the chosen plasmid (see Note 26). Incubate for 12–16 h at 37C. 4. Transformation efficiency can be determined by blue/white screening, colony PCR, or plasmid isolation followed by screening with restriction enzymes (see Note 27). 5. Sequence positive transformants.


3.2.5. SequencingCloned DNA Molecules

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At this point, the cloned inserts can be sequenced by several standard sequencing technologies (see Note 28). For MAPit, we are currently using rolling circle amplification (see Note 29) and conventional automated BigDye sequencing (Applied Biosystems). For this protocol, the plasmid from the frozen bacterial cultures is sequenced at the core facility in the Interdisciplinary Center for Biotechnology Research (ICBR) at the University of Florida. The steps to prepare these bacterial cultures are outlined below. 1. In a 96-well plate inoculate a single positive transformant colony per well containing 100 mL of LB þ amp. Cover the plate with AirPoreTM Tape Sheets (Qiagen). Incubate at 37C for 12–16 h (see Note 30). 2. Add 16 mL 50% glycerol to obtain 8% (v/v) final concentration of glycerol. Cover and seal the plate with aluminum tape (we purchase standard aluminum insulation tape at a home supply center). Close the plate with the corresponding lid and seal it with ParafilmTM. 3. Incubate at 80C for at least 30 min (see Note 31).

4. Notes 1. We purchase 0.5 kg and re-package approx. 5 g amounts of sodium metabisulfite in glass 5 g vials in an H2O- and oxygenfree safety hood. The vials are stored tightly capped in the dark in a sealed bottle with Drierite1 desiccant to protect against oxidation. Artifacts due to poor deamination efficiency have been widely reported (26), but we have found that measures taken to protect the reagents from oxidation during storage and use (29) eliminate such artifacts or greatly reduce them to nondetectable levels. As a simpler alternative, commercially available bisulfite deamination kits such as the EZ DNA Methylation-DirectTM Kit (Zymo Research D5020) or EpiTect1 Bisulfite Kit (Qiagen 59104) achieve comparable results. 2. Deamination primers are designated as a1/a2 or b1/b2 pairs according to the convention of Frommer et al. (16). Primers must be designed to account for deamination of unmethylated residues. For best results, primers should discriminate between deaminated and undeaminated template and should contain as many C (a1/b1) or G (a2/b2) residues as possible while avoiding potential methylation sites. If C residues within MTase target sites cannot be avoided, the appropriate degeneracy (C/T or G/A) should be inserted at this position. We usually design primers to have a Tm of 55–57C at 0.8 mM

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primer in 0.1 M NaCl. It becomes difficult to obtain high yield for amplicons longer than 1 kb because the DNA is partially degraded by the bisulfite conversion procedure. 3. We have observed high numbers of incorrect recombinants with TA cloning. Therefore, for MAPit, we increase the efficiency of obtaining plasmids with the correct inserts through directional cloning. For this, we incorporate different restriction sites at the 50 end of each deamination PCR primer for cloning of the individual templates in the vector of choice. 4. Blue/white screening is used to enrich for insert-positive recombinant plasmids, however, a significant number of false negatives occur when cloning PCR amplicons generated from bisulfite-treated DNA. For this reason, we recommend the verification of cloned inserts by a different method, such as colony PCR or plasmid isolation, before inoculating transformants for sequencing. 5. Owing to potential toxicity of foreign DNA methylation, incubation times both on plates and in liquid media should be minimized in order to avoid selection for hosts carrying plasmids with mutations in the MTase-coding sequence and/ or regulatory sequences. To help mitigate these effects, we have cloned the strong constitutive E. coli promoter conII in the inverse orientation immediately downstream of the M.CviPI open reading frame in pSH1052, which leads to high antisense transcription through the MTase-coding sequence in E. coli. 6. If the plasmid used in the transformation has not yet been tested for activity, inoculate multiple cultures in order to increase the odds of obtaining a functional expression plasmid. 7. When transforming with a YIp, plasmid DNA is prepared for transformation by restriction with enzyme(s) chosen to place the region(s) of homology at the ends of the linear fragment that is to be integrated. Yeast episomal (YEp) and centromeric (YCp) plasmids require no additional preparation prior to transformation. 8. Culture A600 readings are linear within the range of approx. 0.1–0.5 on most spectrophotometers. Therefore, in order to determine the actual density of cultures that are too dense to read accurately, a dilution of a small portion must be done using growth medium to bring the read density into the linear range. The actual culture optical density (OD600) is calculated by multiplying the resulting A600 value by the dilution factor. 9. It is very difficult to accomplish the initial resuspension in 50% PEG4000 by pipetting or vortexing, due to the viscosity of this solution. For this reason it works best to add the


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PEG4000 to the tube and then use the pipet tip to directly disrupt and stir the pellet. This results in an uneven suspension which can then be vortexed to uniformity. Make sure that all of the cell pellet and resuspension is off the tip before discarding it and be sure that no residual cell pellet remains in the bottom of the tube before proceeding. 10. Success rates of approx. 50% are common. 11. Due to potential toxicity of foreign DNA methylation, it is advisable to work with strains that have been freshly patched (20%) than mitotic index (usually 1–2%, but will be often much lower) that facilitates the observation and analysis of chromosome much easier (1). Therefore, drug-induced PCC has been becoming popular and many articles using the technique have been published (8, 29–42). In addition, the drug-induced PCC has an outstanding merit that

Drug-Induced Premature Chromosome Condensation

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allows the interphase chromatin (i.e., G1-, S-, and G2-phase chromatin) to be visualized as condensed form of chromosome structure (1, 24). The drug-induced PCC method is, thus, not only a useful alternative method for mitotic chromosome analysis, but also will be a new analytical tool for interphase chromatin cytogenetics (1). In this chapter, the detailed protocol of druginduced PCC will be described. The protocol is very simple and can be carried out by those not well versed in chromosome preparation and it can be imagined that the drug-induced PCC technique will be used in a wide range of studies in cytogenetics and related fields.

2. Materials 1. Calyculin A (Wako Chemicals) (see Note 2): Reconstitution of calyculin A for 100 mM stock solution. For 100 mg of calyculin A (Wako Chemicals cat. No. 032-14451) lyophilized in a vial supplied from the supplier, dissolve with 1 mL of solvent (DMSO or 100% ethanol, calyculin a does not dissolve in water) will make 100 mM stock solution (M.W. of calyculin A is 1009.17). Aliquot in 100 mL in a microcentrifuge tube and stored at –20C and stable at least 3 months (see Notes 3 and 4). For longer storage, re-lyophilization is recommended. 2. 0.075 M KCl: dissolve 7.45 g KCl (M.W. 74.55) in 100 mL MilliQ water to make a 1 M KCl stock solution. Take 7.5 mL of the 1 M stock solution and make it up to 100 mL with MilliQ water to obtain a 0.075 M KCl working solution. 3. Carnoy’s fixative (methanol:glacial acetic acid = 3 part:1 part): Take three part of methanol and one part of glacial acetic acid (for example, 150 mL of methanol and 50 mL of acetic acid for make 200 mL fixative) and mix together well. The temperature of mixture will be increased according to exothermic reaction. Preferable to prepare just before use, however the fixative can be stored at –20C for a month. After stored in longer time, esterification will proceed resulting in production of methyl acetate and water. Consequently, fixative smells different, and when dropped on a glass slide, fixative does not spread well circularly and water will be left over after evaporation. In this case, new fixative should be prepared. 4. 10 mM phosphate buffer (pH 6.8 at 25C): make as two solutions and combine as follows. a. Solution A: dissolve 136.09 g KH2PO4 in 1 L MilliQ water to make 1 M stock solution.

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b. Solution B: dissolve 174.18 g K2HPO4 in MilliQ water to make it up to 1 L for 1 M stock solution. c. Mix 50.3 mL solution A with 49.7 mL solution B and make it up to 1 L with MilliQ water to make 10 mM working solution. If desired, adjust the pH before the final dilution up to 1 L. d. Gurr Buffer tablet (one tablet for 1 L solution, pH 6.8, available from Invitrogen, cat. No. 10582013) or phosphate buffer powder (for 1 L solution, 1/15 M, pH 6.8, available from Wako Chemicals, cat. No. 163-16471) are can also be used as convenient substitutes for in-house phosphate buffer. 5. Giemsa dye solution (Merck KGaA): dilute with 10 mM phosphate buffer (pH 6.8 at 25C) to make 5% (4–6%) solution (freshly prepare just before use) (see Note 8). 6. Glass slides and cover slips: glass slides are soaked in methanol until use. Wipe out methanol with KIMWIPES.

3. Methods 3.1. Drug-Induced Premature Chromosome Condensation (PCC) for Adherent Cells

3.1.1. Cell Culture, Premature Chromosome Condensation and Harvest

3.1.2. Chromosome Preparation

Plating the cells in culture dish 1–2 days before the experiment in order to reach in 70–80% confluent and growing exponentially at the induction of PCC will be necessary to achieve a high PCC index (see Note 5). One 35 mm culture dish of cells is sufficient to obtain a substantial numbers of PCCs for analyzing the one data point. Add 1/2000 volume of 100 mM calyculin A (50 nM final concentration) to the medium, then incubate for another 30 min at 37C. After incubation, gentle pipetting with the medium using a 1000 mL Pipetman with a blue tip, will allow cells to easily detach and suspend in the medium. Neither trypsinization nor the use of scrapers are needed for detaching cells, as done following colcemid treatment, because the cells become round and attach very loosely on the dishes after calyculin A exposure. If the cells do not become round and detach from the dish, it suggests the loss of calyculin activity. Replace new aliquot calculin A. Transfer the cell-suspended medium into a centrifuge tube, centrifuge at 250 g for 5 min, and discard the supernatant by inverting the tube (see Note 7). Loosen the cell pellet by shaking the tube and add gently prewarmed (37C) 1.5 mL 0.075 M KCl to suspend the cells for hypotonic treatment. Incubate at 37C for 20 min. Fix cells by slowly adding the same volume of cold Carnoy’s fixative to the


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medium and mix well gently. Centrifuge at 250 g for 5 min, discard the supernatant, loose the cell pellet, add the fixative, and centrifuge. Repeat three to four times these steps for complete fixation. After fixation, cells are finally suspended in adequate volume of fixative (200–300 ml for 35 mm dish culture of cells). The prepared chromosome suspension ( 15 mL using a Pipetman using a yellow tip) was dropped on a glass slide and air dried. To make sure that the cells are certainly condensed, observe the chromosome spreads under a microscope at low magnification before staining. Normally, many chromosome spreads are seen, and 10–20% PCC index will be achieved. Stain with 5% Giemsa dye solution, cover with a coverslip and mount with sealant for observation (see Note 8). Usually, one sample slide may give a substantial number of PCCs for karyotyping or chromosome aberration analysis. Unstained chromosome specimens will be stored in a desiccated container and will be subjected to several special banding procedures (G-, Q-, or R-banding), in situ hybridization including FISH or immunostaining. 3.2. Drug-Induced PCC for Suspension Cells

3.2.1. Cell Culture, Premature Chromosome Condensation and Harvest

3.2.2. Chromosome Preparation

The protocol for suspension cells such as peripheral blood lymphocytes is basically the same as for adherent cells. For suspension cells, okadaic acid also works well as calyculin A (see Note 4). Prepare the suspension cell culture in an appropriate cell concentration (i.e., 105 – 106 cells/mL, depends on cell types used) to maintain exponentially growing at the time of PCC induction (see Note 5). Prior to the PCC induction, it is convenient to transfer the cells in a centrifuge tube. One milliliter of cell culture is sufficient to obtain a substantial numbers of PCCs for analyzing the one data point. Add 50 nM final concentration of calyculin A to the medium, process as same as the protocol for attached cells. After 30 min incubation, the cells are harvested by centrifuging and subjected to hypotonic treatment, cell fixation and spreading on the glass slid as the same as adherent cells (see Section 3.1.2).

4. Notes As easily recognized, the drug-induced PCC protocol is a very simple, substituting use of calyculin A instead of colcemid in established chromosome preparation protocols. However, some points need to be attended to in order to achieve high PCC index and good-quality PCCs. Following are the vital points that the authors have found to be important:

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1. Premature chromosome condensation (PCC) usually means the phenomenon or method, and the condensed interphase chromatin is called as prematurely condensed chromosomes (PCCs). They give the similar abbreviations that are usually used interchangeable. 2. Wako Chemicals is in Japan (Osaka, Japan). Calyculin A is now available from many other chemical suppliers (Section 2.1). 3. It is hard to see and weigh out the lyophilized calyculin A powder in the vial. So, pour the desired solvent (ethanol or DMSO) into the vial using a Pipetman, gently slant and swirl the vial to dissolve the whole chemicals, and transfer the solution to the microcentrifuge tube (e.g., Eppendorf tube). If dissolved in ethanol, the use a screw-cap tube with air-tight cap is preferable (Section 2.1), otherwise ethanol will escape in vapor and the concentration of calyculin A will change. Accordingly, DMSO is more preferable for dissolve calyculin A. 4. Okadaic acid, its salts or derivatives, or other inhibitors endothal or chantharidine can also be used as PCC inducers (25–27), but these works only in suspension type cells (for example, peripheral blood lymphocytes) but not in adherent cells. The reason is unclear and presumably due to the different cell sensitivities to these chemicals. Therefore, the use of calyculin A is recommended for PCC induction both in suspension and in adherent types of cell. If okadaic acid is used in suspension cells, 100 nM final concentration rather than 50 nM will work better (Sections 2.1, 3.1.1, 3.2.1). 5. The PCC efficiency is highly dependent on cell types, cell growing, dose, or incubation time of calyculin A. Usually, 50 nM final concentration of calyculin A works very well in many kinds of cells and a sufficient numbers of PCCs will be obtained following 30 min of incubation. However, some cells show the resistance to calyculin A but others are hypersensitive to calyculin A. In such cases, start the experiment at first with 50 nM concentration and for 30 min incubation time, then increase or decrease the dose and/or the incubation time depending on the cell response and resulting PCC index. Keep in mind that the cells should be growing in exponentially at the induction of PCC otherwise PCC index will drop down. In the case of adherent types of cells, cells should be re-plated 1–2 day(s) before the experiment to reach in 70–80% confluent at the time of PCC induction. Avoid re-plating the cells on the day of the experiment. The higher the dose and the longer the incubation time, the PCC index is higher but the resulting PCCs become hyper-condensed. Therefore, the appropriate dose and incubation time should be taken on balance depend on the experiment design. PCC starts 5 min after beginning of


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calyculin exposure, and 10 min of incubation will induce a substantial numbers of PCCs (8, 28, 33) (Sections 3, 3.1.2, 3.2.2). 6. Usually, calyculin A works efficiently and gives a high PCC index. Combined use of colcemid with calyculin A will be sometimes effective in obtaining the higher PCC index (31). 7. A round bottomed centrifuge tube (for example, Beckton & Dickinson 5 mL polystyrene centrifuge tube with snap cap, cat. No. 352058) is preferable for chromosome preparation, so the cells are less packed after centrifugation as to avoid the cells aggregation (Sections 2, 3.1.1, 3.2.1). 8. The tint of Giemsa staining varies as the pH of phosphate buffer shifts; color turns from reddish purple (pH 6.8) to bluish purple (pH 7.2). Which pH of phosphate buffer used is dependent on the individual favorite, but reddish purple color seems to be better for observation and taking images under a microscope (Sections 2.4, 2.5, 3.1.2, 3.2.2). 9. Calyculin A, okadaic acid, or other phosphatase inhibitors induce PCC predominantly in G2- and S-phase but only few in G1- and G0-phase (1). Presumably drug-induced PCC exploits the activation of intracellular MPF by phosphatase inhibitors, but amount of intracellular maturation promoting factor (MPF, also known as p34cdc2/cyclinB complex) (43–47) is very low in G1- or G0-phase (1, 48). In the cases that PCC analysis is requested in G0- or G1-phase of cells, the conventional fusion PCC is still worthwhile to do. Another approach of drug-induced PCC in G0-phase peripheral blood lymphocytes was reported using adenosine triphosphate (ATP) and p34cdc2/cyclinB with okadaic acid (49). The protocol of fusion PCC or okadaic acid:p34cdc2/cyclinB:ATP induced PCC has been described previously (2, 12, 14, 49, 50).

Acknowledgment I wish to express my thanks to Prof. Srikumar Chellappan for recommending me to contribute this chapter.

References 1. Gotoh, E., and Durante, M. (2006) Chromosome condensation outside of mitosis: Mechanisms and new tools J. Cell. Physiol. 209, 297–304 and Cover page. 2. Johnson, R. T., and Rao, P. N. (1970) Mammalian cell fusion: induction of premature

chromosome condensation in interphase nuclei Nature 226, 717–722. 3. Sperling, K., and Rao, P. N. (1974) The phenomenon of premature chromosome condensation: its relevance to basic and applied research Humangenetik 23, 235–258.

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4. Sperling, K., and Rao, P. N. (1974) Mammalian cell fusion. V. Replication behaviour of heterochromatin as observed by premature chromosome condensation Chromosoma 45, 121–131. 5. Rao, P. N., Wilson, B., and Puck, T. T. (1977) Premature chromosome condensation and cell cycle analysis J. Cell. Physiol. 91, 131–141. 6. Mullinger, A. M., and Johnson, R. T. (1983) Units of chromosome replication and packing J. Cell Sci. 64, 179–193. 7. Ito, S., Gotoh, E., Ozawa, S., and Yanagi, K. (2002) Epstein-Barr virus nuclear antigen-1 is highly colocalized with interphase chromatin and its newly replicated regions in particular J. Gen. Virol. 83, 2377–2383. 8. Gotoh, E. (2007) Visualizing the dynamics of chromosome structure formation coupled with DNA replication Chromosoma. 116(5), 453–462. 9. Cornforth, M. N., and Bedford, J. S. (1983) X-ray – induced breakage and rejoining of human interphase chromosomes Science 222, 1141–1143. 10. Cornforth, M. N., and Bedford, J. S. (1983) High-resolution measurement of breaks in prematurely condensed chromosomes by differential staining Chromosoma 88, 315–318. 11. Pantelias, G. E., and Maillie, H. D. (1984) The use of peripheral blood mononuclear cell prematurely condensed chromosomes for biological dosimetry Radiat. Res. 99, 140–150. 12. Durante, M., George, K., Wu, H., and Yang, T. C. (1996) Rejoining and misrejoining of radiation-induced chromatin breaks. I. experiments with human lymphocytes Radiat. Res. 145, 274–280. 13. Hittelman, W. N. (1990) Direct measurement of chromosome repair by premature chromosome condensation Prog. Clin. Biol. Res. 340B, 337–346. 14. Pantelias, G. E., and Maillie, H. D. (1983) A simple method for premature chromosome condensation induction in primary human and rodent cells using polyethylene glycol Somatic Cell Genet. 9, 533–547. 15. Rao, P. N., and Johnson, R. T. (1970) Mammalian cell fusion: studies on the regulation of DNA synthesis and mitosis Nature 225, 159–164. 16. Rao, P. N., and Johnson, R. T. (1971) Mammalian cell fusion. IV. Regulation of

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condensation (PCC) technique. EPO 95305645.4 UK, Germany, Italy, France, Netherlands, Sweden Gotoh, E. Agents and a method of chromosome preparation using protein phosphatase inhibitors induced premature chromosome condensation. Canadian Patent 2,155,813 Canada Bryant, P. E., and Mozdarani, H. (2007) A comparison of G2 phase radiation-induced chromatid break kinetics using calyculin-PCC with those obtained using colcemid block Mutagenesis. 22(5), 359–362. Gotoh, E., and Asakawa, Y. (1996) Detection and evaluation of chromosomal aberrations induced by high doses of gamma-irradiation using immunogold-silver painting of prematurely condensed chromosomes Int. J. Radiat. Biol. 70, 517–520. Asakawa, Y., and Gotoh, E. (1997) A method for detecting sister chromatid exchanges using prematurely condensed chromosomes and immunogold-silver staining Mutagenesis 12, 175–177. Durante, M., Furusawa, Y., and Gotoh, E. (1998) A simple method for simultaneous interphase-metaphase chromosome analysis in biodosimetry Int. J. Radiat. Biol. 74, 457–462. Johnson, R. T., Gotoh, E., Mullinger, A. M., Ryan, A. J., Shiloh, Y., Ziv, Y., and Squires, S. (1999) Targeting double-strand breaks to replicating DNA identifies a subpathway of DSB repair that is defective in ataxiatelangiectasia cells Biochem. Biophys. Res. Commun. 261, 317–325. Gotoh, E., Kawata, T., and Durante, M. (1999) Chromatid break rejoining and exchange aberration formation following gamma-ray exposure: analysis in G2 human fibroblasts by chemically induced premature chromosome condensation Int. J. Radiat. Biol. 75, 1129–1135. IAEA (2001) Cytogenetic analysis for radiation dose assessment. A manual, International Atomic Energy Agency, Vienna. Bezrookove, V., Smits, R., Moeslein, G., Fodde, R., Tanke, H. J., Raap, A. K., and Darroudi, F. (2003) Premature chromosome condensation revisited: a novel chemical approach permits efficient cytogenetic analysis of cancers Genes Chromosomes Cancer 38, 177–186. El Achkar, E., Gerbault-Seureau, M., Muleris, M., Dutrillaux, B., and Debatisse, M.

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(2005) Premature condensation induces breaks at the interface of early and late replicating chromosome bands bearing common fragile sites Proc. Natl. Acad. Sci. USA 102, 18069–18074. Srebniak, M. I., Trapp, G. G., Wawrzkiewicz, A. K., Kazmierczak, W., and Wiczkowski, A. K. (2005) The usefulness of calyculin A for cytogenetic prenatal diagnosis J. Histochem. Cytochem. 53, 391–394. Gotoh, E., and Tanno, Y. (2005) Simple biodosimetry method for cases of high-dose radiation exposure using the ratio of the longest/shortest length of Giemsa-stained drug-induced prematurely condensed chromosomes (PCC) Int. J. Radiat. Biol. 81, 379–385. Gotoh, E., Tanno, Y., and Takakura, K. (2005) Simple biodosimetry method for use in cases of high-dose radiation exposure that scores the chromosome number of Giemsastained drug-induced prematurely condensed chromosomes (PCC) Int. J. Radiat. Biol. 81, 33–40. Mochida, A., Gotoh, E., Senpuku, H., Harada, S., Kitamura, R., Takahashi, T., and Yanagi, K. (2005) Telomere size and telomerase activity in Epstein–Barr virus (EBV)-positive and EBV-negative Burkitt’s lymphoma cell lines Arch. Virol. 150, 2139–2150. Deckbar, D., Birraux, J., Krempler, A., Tchouandong, L., Beucher, A., Walker, S., Stiff, T., Jeggo, P., and Lobrich, M. (2007) Chromosome breakage after G2 checkpoint release J. Cell Biol. 176, 749–755. Kanda, T., Kamiya, M., Maruo, S., Iwakiri, D., and Takada, K. (2007) Symmetrical localization of extrachromosomally replicating viral genomes on sister chromatids J. Cell Sci. 120, 1529–1539. Masui, Y. (1974) A cytostatic factor in amphibian oocytes: its extraction and partial characterization J. Exp. Zool. 187, 141–147. Masui, Y. (2001) From oocyte maturation to the in vitro cell cycle: the history of discoveries of Maturation-Promoting Factor (MPF) and Cytostatic Factor (CSF) Differentiation 69, 1–17. Dunphy, W. G., Brizuela, L., Beach, D., and Newport, J. (1988) The Xenopus cdc2 protein is a component of MPF, a cytoplasmic regulator of mitosis Cell 54, 423–431.

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46. Gautier, J., Norbury, C., Lohka, M., Nurse, P., and Maller, J. (1988) Purified maturationpromoting factor contains the product of a Xenopus homolog of the fission yeast cell cycle control gene cdc2+ Cell 54, 433–439. 47. Maller, J., Gautier, J., Langan, T. A., Lohka, M. J., Shenoy, S., Shalloway, D., and Nurse, P. (1989) Maturation-promoting factor and the regulation of the cell cycle J. Cell Sci. Suppl. 12, 53–63. 48. Doree, M., and Galas, S. (1994) The cyclindependent protein kinases and the control of cell division FASEB J. 8, 1114–1121. 49. Prasanna, P. G., Escalada, N. D., and Blakely, W. F. (2000) Induction of

premature chromosome condensation by a phosphatase inhibitor and a protein kinase in unstimulated human peripheral blood lymphocytes: a simple and rapid technique to study chromosome aberrations using specific whole-chromosome DNA hybridization probes for biological dosimetry Mutat. Res. 466, 131–141. 50. Prasanna, P. G., and Blakely, W. F. (2005) Premature chromosome condensation in human resting peripheral blood lymphocytes for chromosome aberration analysis using specific whole-chromosome DNA hybridization probes Methods Mol. Biol. 291, 49–57.

Chapter 7 Analysis of DNA Topology in Yeast Chromatin Randall H. Morse Abstract Topology of closed circular DNA is affected by its packaging into nucleosomes and potentially by alteration of nucleosome structure. Changes in topology that reflect alterations in chromatin structure can be measured and quantified using closed circular plasmids from living yeast. Here we describe detailed protocols for measuring DNA topology in yeast chromatin. Key words: Chromatin, yeast, nucleosomes, topology.

1. Introduction Topological measurements have been used to investigate chromatin structure since the discovery that closed circular DNA molecules differing only in the number of supercoils that they contain can be resolved by gel electrophoresis (1) and the recognition that nucleosomes, the fundamental units of chromatin, confer defined changes in topology on closed circular DNA templates (1, 2). The number of supercoils introduced into a closed circular DNA molecule per nucleosome can be calculated by comparing the linking number of unassembled and nucleosome-assembled DNA. Nucleosomes can be counted by electron microscopy, and the total number of supercoils analyzed by gel electrophoretic comparison of naked and nucleosome-assembled DNA relaxed by topoisomerase (the latter after stripping off the histones). This kind of analysis has revealed that one nucleosome confers almost exactly one negative supercoil into DNA both in vitro (2–4) and in vivo, in yeast as well as in higher eukaryotes (5–7). This change is somewhat smaller than expected, since DNA wraps around the Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_7 Springerprotocols.com

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histone octamer about 1.75 turns, leading to a calculated change of –1.65 in linking number (8). Some of the excess change is accounted for by a compensating alteration in the twist of nucleosomal DNA, but a part of it remains unaccounted for (9). Nevertheless, it is an empirical fact and allows the number of nucleosomes on a closed circular DNA molecule to be inferred by comparing the linking number of relaxed plasmid chromatin to that of relaxed naked DNA. Since each nucleosome confers one negative supercoil to DNA, loss of nucleosomes or alteration of nucleosome structure is expected to change DNA topology. This property has been exploited in numerous studies of chromatin structure and function (for a thorough discussion of studies on DNA topology in chromatin structure and gene expression done prior to 1992, see ref. (10)). For example, a variety of mutations in the histone proteins have been shown to alter topology of plasmid chromatin in the yeast Saccharomyces cerevisiae (11–13), as has histone depletion (14). Acetylation of the histone amino termini has been reported to alter nucleosome topology in vitro (4, 15). Removal of the same amino termini by trypsinization was reported not to alter linking number (16); whether these two observations are compatible or not has not been determined. Conflicting in vivo experiments on the effect of histone acetylation on topology has been reported (17, 18). Chromatin topology has been studied in plasmids containing binding sites for transcriptional activators in frog oocytes and in yeast; in both systems, loss of negative supercoiling consistent with nucleosome loss or alteration was observed in the presence of the transcriptional activator, and in both cases required an activation domain (19, 20). More recently, topological studies have been combined with chromatin immunoprecipitation and density centrifugation to demonstrate loss of nucleosomes by histone eviction from the activated PHO5 promoter in yeast (21, 22). Topological studies have also been used in structural investigations of chromatin. Naked DNA undergoes thermal untwisting; this small helical unwinding that occurs with increasing temperature (about 0.011 radial degrees/C/base pair) results in increased negative supercoiling for molecules relaxed at increasing temperatures (23, 24). This effect is suppressed by incorporation of DNA into nucleosomes, such that each nucleosome prevents about 160–180 bp of DNA from thermally untwisting (16, 25–27). In the yeast Saccharomyces cerevisiae, however, this suppression is reduced, such that the equivalent of only about 60 bp of DNA per nucleosome is prevented from thermally untwisting (7, 28). These effects of nucleosomal packaging on thermal untwisting of DNA have also been interpreted as arising from changes in nucleosome shape (29–31). This explanation, however, requires opposite shape changes for yeast and higher

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eukaryotic nucleosomes in their responses to temperature and also requires the extent of deformation of yeast nucleosomes to be proportional to the change in temperature, which seems not entirely plausible. Regardless of the explanation, it is important to be aware that altering the temperature at which plasmid chromatin is prepared from yeast cells will affect DNA topology (see Note 1). One considerable advantage that topological analysis has over other methods of analyzing chromatin structure is that small samples can be used and isolation of material is done rapidly. These features have allowed kinetic analysis of minichromosome topology which showed that although topoisomerase relaxes torsional stress in yeast minichromosomes extremely rapidly at room temperature and above (1–5 min), no change in topology is observed due to transcription of the same templates, leading to the inference that nucleosomes recover their normal structure quickly following polymerase passage (32). Kinetic analysis was also used to show that the change in topology caused by activator binding to a yeast episome mentioned above is complete within 45 min, indicating that DNA replication is not required for the alteration of chromatin structure (19). 1.1. Theory

Experiments predating the separation of individual DNA topoisomers by gel electrophoresis demonstrated that the free energy associated with DNA supercoiling is proportional to the square of the extent of supercoiling (equation [1]) (33, 34). Gsc ¼ K ðLkÞ2

[1]

This behavior is in accordance with Hooke’s Law, suggesting that the energy associated with supercoiling arises from elastic deformation of structure. Together with classical thermodynamic considerations, equation [1] implies that the concentration of individual topoisomers at equilibrium (i.e., after being relaxed by DNA topoisomerase) will conform to a Gaussian distribution (23, 24, 35). Specifically, ðRT=Lki Þ lnðIi =ImaxÞ ¼ KðLki þ 2 wTÞ

[2]

where R ¼ 1.99 cal deg1 mol1 (the gas constant), T is the temperature, Lki is the difference in linking number between topoisomer i having relative concentration Ii and the maximally abundant topoisomer having relative concentration Imax, and wT is the number of non-integral turns by which the precise center of the topoisomer distribution differs from Lkmax, the linking number of the maximally abundant topoisomer. Plotting (1/Lki) ln (Ii/Imax) versus Lki therefore yields a straight line having slope m and intercept 2 m wT; dividing the intercept by two times the slope yields wT, which then allows determination of the

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precise center of the topoisomer distribution. Since analysis of plasmid chromatin from living cells has revealed that minichromosome topoisomers also conform to a Gaussian distribution, the same analysis can be applied to closed circular DNA packaged as chromatin (26, 27, 36).

2. Materials 1. Media for growing yeast 2. Acid-washed glass beads (425–600 m) 3. Solution of 5 mg/mL proteinase K, 5% SDS (this may be stored indefinitely in 1 ml aliquots at –20C and thawed by warming in a 37C water bath) 4. TE solution (10 mM Tris–HCl pH 8.0, 0.5 mM EDTA); ice. 5. Phenol 6. Chloroform 7. 4 M ammonium acetate 8. Ethanol 9. TE solution (10 mM Tris–HCl pH 8.0, 0.5 mM EDTA) 10. 10 mg/ml RNase A. 11. Agarose (high-quality molecular biology grade) 12. 60 mg/mL solution of chloroquine diphosphate (chloroquine is toxic and should be handled with caution) in deionized water (this can be stored indefinitely in the dark at 4C; we make this up in a 50 mL conical tube and wrap the tube in aluminum foil) 13. 50 TAE buffer: 2 M Tris–acetate, 50 mM EDTA (242 g Tris base, 57.1 mL glacial acetic acid, 100 mL 0.5 M EDTA, pH 8.0 per liter of solution (37) 14. Loading dye: 18% Ficoll 400 plus 0.3% xylene cyanol (see Note 2). 15. Nylon membrane for blotting. 16. 0.5 M HCl (concentrated HCl is 12 M) 17. 0.5 M NaOH (dilute from 10 M NaOH solution) 18. 3 mm paper 19. 20 SSC (35): 3 M NaCl, 0.3 M sodium citrate, pH 7.0 (175 g NaCl, 88.2 g sodium citrate dihydrate dissolved in 800 mL H2O; pH adjusted with HCl and volume adjusted to 1 L)


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20. 6 SSC 21. 0.9 M Tris, 1.5 M HCl (add 12.5 mL concentrated HCl to 87.5 mL 1 M Tris–HCl, pH 8.0) 22. Paper towels 23. Plastic wrap 24. UV crosslinker 25. Sealable plastic bag 26. Heat sealer. 27. Radioactively labeled probe specific to the DNA you wish to analyze (use any of the commercially available kits for random prime labeling of DNA) 28. 1 M NaPi: 134 g Na2HPO4.7H2O (71 g anhydrous) plus 4 mL phosphoric acid per liter (pH will be 7.2) (add sodium phosphate to water; if you add water to the sodium phosphate it becomes very difficult to dissolve) 29. Hybridization buffer: 52.5 mL 1 M NaPi pH 7.2, 12 mL H2O, 35 mL 20% SDS, 0.2 mL 0.5 M EDTA, pH 8, 1 g bovine serum albumin (dissolve by 5–10 min incubation in 65C water bath; this may form some precipitate on standing but can be redissolved at 65C) 30. Church Gilbert wash #1: 350 mL H2O, 20 mL 1 M NaPi, 125 mL 20% SDS, 1 mL 0.5 M EDTA (pH 8) 31. Church Gilbert wash #2: 910 mL H2O, 40 mL 1 M NaPi, 50 mL 20% SDS, 2 mL 0.5 M EDTA (pH 8). 32. Church Gilbert wash #1 and #2: 0.1% SDS/0.2 SSC; all at 65C. 33. X-ray film or phosphorimager screen. 34. Scanning densitometer or phosphorimager. 35. Spreadsheet computer program.

3. Methods 3.1. Comparison of Topology Among Species Having Relatively Small Differences in Linking Number

This protocol is suitable for many kinds of topological analysis, for example, examining effects of mutant histones, activator binding, and thermal untwisting on topology (see Section 1). Modifications which allow comparison of species having larger differences in linking number, for example, to count nucleosomes by comparing linking number of plasmid chromatin and relaxed naked DNA, are given in a brief section following this one.

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3.1.1. Preparation of DNA

1. Grow yeast cells in appropriate media (10 mL per sample to be analyzed) to A600 ¼ 0.5–1.5. Before taking samples out of shaking incubator, prepare 1.5 mL microfuge tubes: label tubes; add acid-washed glass beads (425–600 m) to 0.3–0.5 mL level of tube; add 100 ml 5 mg/mL proteinase K, 5% SDS. When this is done, spin down yeast cells in desktop centrifuge 2000 rpm (800 g) 2 min at room temperature. Pour off supernatant and resuspend pellets in TE solution. Transfer resuspended pellets to labeled microfuge tube containing glass beads and proteinase K/SDS. Vortex 1 min at top speed (see Note 3); place on ice 15 s; vortex 1 min. 2. Hold the microfuge tube upside down and make a small hole in the bottom with a hot dissecting needle (heated over a flame) (see Note 4). Collect the lysed cell suspension by placing the tube onto a clean labeled 1.5 mL microfuge tube and spinning 8 K rpm in a microfuge 5–10 s. Discard the tube with glass beads (the suspension should now be in the lower microfuge tube) and spin the lower microfuge tube 8–10 min at full speed in microfuge. Since there is proteinase K and SDS in the sample it does not matter whether these spins are done at 4C or room temperature. Transfer 320 mL of the supernatant to a new labeled 1.5 mL microfuge tube and incubate 37C (up to 50C is fine) at least 2 h. 3. Extract sample with phenol and chloroform. Add 80 mL 4 M ammonium acetate to sample and 1 mL 100% ethanol; precipitate. Spin sample 10 min at top speed in microfuge at 4C, discard supernatant (it can be poured off), and take up the sample in 30 mL TE solution plus 1 mL 10 mg/ml RNase A (see Note 5).

3.1.2. Gel Electrophoresis and Blotting (37, 38) 3.1.2.1. Electrophorese DNA on a TopoisomerResolving Gel

Pour a large (21 cm or more from wells to end of gel) agarose gel containing 40 mg/ml chloroquine diphosphate (see Note 6). The agarose concentration to be used depends on the size of the closed circular DNA being analyzed: for a 1.4–2 kb plasmid; use 1.5% agarose; for 2–3 kb, use 1.2%; for 3–4.5 kb, use 1%; for 4–6 kb, use 0.8%; and for 6–8 kb, use 0.7% (e.g., for the 2 mm plasmid). Use TAE buffer in the gel and running buffer and add chloroquine (133 ml of 60 mg/ml chloroquine diphosphate solution for a 200 ml gel) after letting the melted agarose cool to 55–65C. Use the same chloroquine concentration in the gel and running buffer; it is not necessary to recirculate the gel buffer solution during electrophoresis. Add 1 ml of loading dye to 15 ml of each sample (it is not necessary to add chloroquine to the samples), load samples and run at 2.5 V/cm (measure cm from electrode to electrode) for 18–20 h (see Note 7). The gel can be run at room temperature and need not be shielded from light. Do not add ethidium bromide to the gel or running buffer! (See Note 8)


3.1.2.2. Blot Gel to Nylon Membrane

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1. Cut out part of gel to be blotted (discard empty lanes; it is also desirable to slice away the extreme edges of the gel, as the menisci make it not flat and more difficult to achieve uniform contact with the nylon membrane) and measure its size. 2. Soak gel in 0.5 N HCl 2 10 min in a glass or plastic tray. No shaking is necessary. 3. Soak in 0.5 N NaOH 2 10 min. 4. Meanwhile, with a razor blade and gloves, cut out a piece of nylon membrane such as Genescreen (Dupont) or Duralon UV (Stratagene) the same size as the gel. Also cut three pieces of 3 mm paper to the same size and three pieces slightly larger. Soak the membrane in 6 SSC. 5. Soak gel in 20 SSC plus a few milliliter of 0.9 M Tris/ 1.5 M HCl to neutralize (neutralization does not have to be complete for nylon; somewhere in the pH 6–8 range is fine). Soaking can be from 5–10 min to a half hour. 6. Lay gel, bottom up, on a flat plastic gel tray. Lay the nylon membrane over the gel (be sure you have marked your gel in some way; by loading lanes asymmetrically, or cutting a corner of the gel and the membrane, so you know the orientation when it comes time to look at the autoradiogram) and roll out gently but firmly with a plastic pipet. It is not necessary to apply much pressure here, and too much pressure can crack the gel; the rolling is done only to remove bubbles. Soak the three 3 mm paper sheets that are larger than the gel in 6 SSC and lay over gel plus membrane and roll out bubbles gently but firmly. Turn gel plus membrane plus 3 mm paper onto a stack of paper towels; for larger gels these need to be unfolded (see Note 9). You are blotting! Soak 3 mm paper pieces the same size as gel in 20 SSC and lay over the top of the gel and roll out bubbles. Cover with a film wrap such as Saran Wrap, then a paper towel, then a flat plastic gel tray. Put a light weight (an empty one liter bottle is good) on top. 7. After 15–20 min, remove weight, gel tray, paper towel, and Saran wrap. Pick up gel plus 3 mm paper on both sides and remove wet paper towels from underneath. Replace blot, wet the top with 20 SSC and replace topping. You can use a heavier weight, such as two half-full one liter bottles. 8. When blotting is complete (2 h to overnight), briefly immerse membrane in 6 SSC (to clean of agarose fragments, stray fruit flies, etc.), keeping track of which side the nucleic acid is on, then lay the membrane face up on a piece of 3 mm paper. Irradiate in a UV crosslinker (e.g., a Stratalinker from Stratagene or the equivalent). (You are irradiating the side the material is on to crosslink it to the membrane).

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9. Put the blot in a sealable plastic pouch. The easiest way to do this is to slide it in on a piece of 3 mm paper without touching the membrane to the pouch, then touch the moist membrane against the bag and allow surface tension to keep the membrane in the bag while you remove the 3 mm paper. You are ready to hybridize. 3.1.2.3. Hybridize Blot with a Probe Specific to the Closed Circular DNA You Wish to Analyze

1. Prepare a double-stranded, radioactively labeled probe using DNA which is specific to the closed circular DNA you wish to analyze, using any of the many commercially available kits for random-primed labeling. 2. Add hybridization mix to the bag containing your blot (about 4 ml for one pint bag, 10 ml for one quart, 15 ml for two quart; enough to ensure no bubbles). Push bubbles to top and heat-seal the bag. Place in bath or hybridization oven at desired temperature, usually 65C, for 5 min to 2 h. 3. Denature labeled double-stranded probe by heating in a screw cap microfuge tube at 95C for 3 min. Put probe on ice. Add probe to a few ml – 2–5 ml, depending on blot size– hybridization buffer in a plastic tube at room temperature. 4. While the probe is denaturing at 95C, take the blot out of the bath or hybridization oven, slice a corner of the pouch (if you can slice through only one side of the pouch this makes adding the probe easier), and insert a pipet tip to hold the opening open after squeezing or pipeting out hybridization buffer. Use a 5 or 10 ml pipet to add hybridization buffer containing labeled probe when ready and reseal. Replace in bath or hybridization oven and allow to hybridize from 8 to 24 h. 5. Prepare Church Gilbert wash buffers #1 (500 ml) and #2 (1 l) and 0.2 SSC/0.1% SDS (500 ml) and place at desired temperature (usually 65C) to prepare for washing blot.

3.1.2.4. Wash and Expose Blot

1. Wash blot: slice bag open over a plastic tray, carefully remove membrane from bag, dispose of radioactive bag and buffer, and wash blot 2 5 min at room temperature using hot (65C) Church Gilbert wash #1. Wash again at room temperature 2 5 min with hot (65C) Church Gilbert wash #2. Then add fresh wash #2 and place blot in 65C oven (or bath) for 10–25 min (shaking is not necessary). Pour off #2 wash and wash at 65C for 10–30 min with 0.2 SSC/0.1% SDS. 2. Remove blot from tray, lay on 3 mm paper. Wrap in Saran wrap and expose to film or to phosphorimager screen.

3.1.3. Analysis

3.1.3.1. Obtain the intensities of the individual topoisomer bands (see Fig. 7.1) either by using a phosphorimager or by using a densitometer to scan an autoradiogram (see Note 10).


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Fig. 7.1 Finding the centers of topoisomer distributions. Yeast harboring the TALS episome (derived from the TRP1ARS1 plasmid) was grown in glucose and galactose medium, as indicated, and TALS topoisomers were resolved on a 1.5% agarose gel containing 40 mg/ml chloroquine diphosphate (19; TALS is 1.8 kb). The linear scans which show the relative intensities of the topoisomers were obtained by scanning the autoradiogram on a flatbed scanner and using the public domain program NIH Image (available at http://rsb.info.nih.gov/nih-image/) to trace individual lanes. The bands marked by asterisks are the most intense. (The traces are for illustrative purpose; intensities of individual bands were obtained using a phosphorimager.) Plotting (1/Lki).ln(Ii/Imax) versus Lki (for simplicity, only the galactose lane is shown) yields a straight line having slope –0.267 and intercept 0.106 (r2 ¼ 0.992); thus b/2 m ¼ 0.20, and the center is 0.20 downward (more positively supercoiled) from the most abundant topoisomer, as shown by the dot. A similar plot for the glucose lane yields b/2 m ¼ 0.447, so the net difference in linking number is 0.6.

3.1.3.2. Use a spreadsheet program such as Excel to calculate the Gaussian center of the topoisomer distribution in each lane as follows: 1. Determine the band having the greatest intensity; assign it a relative linking number (Lk) of zero. Assign linking numbers to the bands having more negative linking number (more slowly migrating in a 40 mg/ml chloroquine gel) –1, –2, –3, etc, and the bands corresponding to more positive linking number þ1, þ2, etc.

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2. Plot (1/Lki) ln (Ii/Imax) versus Lki, where Lki corresponds to the assigned relative linking number of the band i having relative intensity Ii, and Imax is the intensity of the maximally intense band. Note that Imax corresponds to Lki ¼ 0 and so cannot be used in the plot (Fig. 7.1). 3. Find the slope (m) and intercept (b) of the resulting straight line (see Notes 11 and 12). Calculate b/2 m; this will be a number between –0.5 and 0.5 if your assignment of the most intense band was correct (that is, if this is the band closest to the center of the distribution). Regardless of whether you correctly assigned the band or not, the exact center of the distribution can be found a distance b/2 m from the band you assigned as the most intense, with the direction being toward more negative linking number if b/2 m is positive, and toward more positive linking number if b/2 m is negative. 4. You can now calculate differences in topology from one sample to another by examining the relative shifts in linking number from one lane to another, taking care that you have assigned the shift measured by b/2 m relative to the correct topoisomer band in each lane (see Note 13). 3.2. Comparison of Topology Among Species Having Relatively Large Differences in Linking Number

In some cases it may be desirable to compare topology between molecules having larger differences in linking number, for example, to determine the average number of nucleosomes present on a minichromosome by comparing the number of supercoils present in plasmid chromatin relative to relaxed, naked DNA (39). In such cases it may be advantageous to use 2D electrophoresis [Fig. 7.2; (40)]. Normally, molecules which are highly positively or

Fig. 7.2. Using 2D gel electrophoresis to determine linking number of plasmid chromatin relative to naked DNA. Relaxed, naked DNA was run together with plasmid chromatin SV40 DNA and a ‘‘connector’’ DNA (B), and the connector DNA was run alone (A). The first dimension was run downward at 1.3 mg/ml chloroquine diphosphate, and the second dimension was run left to right at 0.3 mg/ml chloroquine diphosphate, using 0.7% agarose. Linking numbers, relative to the center of the relaxed, naked DNA sample (assigned a value of zero) are indicated. From Drabik et al. (37), with permission.


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negatively supercoiled under the conditions of electrophoresis are compressed as rapidly migrating species, while those having a nearly relaxed configuration under electrophoresis conditions may be compressed together with the slowly migrating nicked circular molecules. By electrophoresing the same molecules at different chloroquine concentrations, these compressed species can be resolved as individual topoisomers on a single gel. Thus, although this technique does not increase sensitivity, it does allow resolution of more topoisomer species on a single gel which can aid in assessing large linking number differences. This protocol differs from that already given by the following modifications. 3.2.1. Preparation of Relaxed Naked DNA and of ‘‘Connector’’ DNA

These can be prepared from purified plasmid DNA (the same species being analyzed), if available, or from the material purified as in parts 1–3 above if not (for example, for 2 micron plasmid or other plasmids having only yeast sequences). For a truly accurate count of nucleosome number, the same buffer conditions and temperature should be used to relax naked DNA and isolated plasmid chromatin; the preparation of the latter is beyond the scope of this chapter. At the least, the naked DNA should be relaxed at the same temperature as the yeast cells from which plasmid chromatin DNA was prepared were grown (usually 30C). DNA and minichromosomes can be relaxed using commercially available topoisomerase I under manufacturer’s suggested conditions (see Note 14). For small plasmids ( 0.97. It may happen that a topoisomer distribution

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does not yield a good fit to a straight line because one or two points deviate from what would otherwise be a good fit. This can occur if there is a spot on the autoradiogram or if linearized plasmid, which generally runs a bit faster than nicked circular molecules, is accidentally included in the analysis. It can also be due to using bands which are very faint and which therefore include too much background noise. These defects can be corrected by simply dropping the offending point from the analysis. If more than one or two (at most) points do not fit the straight line, or if it is not obvious which points do not fit, this may be an indication of other problems. For example, the sample may not have been at equilibrium or background may be too high. Repeating the experiment several times will reveal if a sample not fitting a Gaussian distribution is inherent (for example, in a kinetic analysis during changing conditions). If this is so the center can be estimated from a less than optimal straight line fit, or by determining the point in the distribution corresponding to the ‘‘center of gravity’’ – the point at which there is equal intensity corresponding to topoisomers of higher and lower linking number. 12. The slope times the length of the closed circular DNA molecule in bp corresponds to the parameter NK. This parameter reflects the breadth of the topoisomer distribution, corrected for plasmid size, and is usually about 400–700 (in bp RT) for plasmid chromatin. Interestingly, this is smaller than the number found for relaxed naked DNA, which is about 1150 for plasmids larger than about 1.5 kb (23, 24). This reflects a broader distribution for plasmid chromatin than for naked DNA, which is likely to be due to heterogeneity in nucleosome number or structure and may reflect a dynamic structure for plasmid chromatin in vivo (36). 13. To obtain quantifiable results with standard deviations, it is necessary to repeat topoisomer analysis in independent experiments. It should go without saying that results from analysis of a single isolation and analysis of DNA topology should never be presented. 14. If relaxation of naked DNA goes to completion, an excellent fit to a Gaussian distribution should be obtained (see Note 11), with NK about 1150 bp RT (see Note 12). If a trailing edge of topoisomers is observed either with naked DNA or plasmid chromatin relaxation by topoisomerase was probably incomplete. 15. At these chloroquine concentrations, naked DNA molecules will migrate as positively supercoiled species, and DNA from plasmid chromatin migrates as negatively supercoiled species.


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For a given plasmid, it may be necessary to experiment with different chloroquine concentrations in one or both dimensions.

Acknowledgment I thank Dr. Len Lutter for providing Fig. 7.2.

References 1. Keller, W., and Wendel, I. (1974) Stepwise relaxation of supercoiled SV40 DNA. Cold Spring Harbor Symp. Quant. Biol., 39, 199–208. 2. Germond, J.E., Hirt, B., Oudet, P., GrossBellard, M., and Chambon, P. (1975) Folding of the DNA double helix in chromatinlike structures from simian virus 40. Proc. Natl. Acad. Sci. U.S.A., 72, 1843–1847. 3. Simpson, R.T., Thoma, F., and Brubaker, J.M. (1985) Chromatin reconstituted from tandemly repeated cloned DNA fragments and core histones: a model system for study of higher order structure. Cell, 42, 799–808. 4. Norton, V.G., Imai, B.S., Yau, P., and Bradbury, E.M. (1989) Histone acetylation reduces nucleosome core particle linking number change. Cell, 57, 449–457. 5. Keller, W., Muller, U., Eicken, I., Wendel, I., and Zentgraf, H. (1978) Biochemical and ultrastructural analysis of SV40 chromatin. Cold Spring Harbor Symp. Quant. Biol., 42, 227–243. 6. Pederson, D.S., Venkatesan, M., Thoma, F., and Simpson, R.T. (1986) Isolation of an episomal yeast gene and replication origin as chromatin. Proc. Natl. Acad. Sci. U.S.A., 83, 7206–7210. 7. Morse, R.H., Pederson, D.S., Dean, A., and Simpson, R.T. (1987) Yeast nucleosomes allow thermal untwisting of DNA. Nucl. Acids. Res., 15, 10311–10330. 8. White, J.H., Cozzarelli, N.R., and Bauer, W.R. (1988) Helical repeat and linking number of surface wrapped DNA. Science, 241, 323–327. 9. Hayes, J.J., Tullius, T.D., and Wolffe, A.P. (1990) The structure of DNA in a nucleosome. Proc. Natl. Acad. U.S.A., 87, 7405–7409.

10. Freeman, L.T., and Garrard, W.T. (1992) DNA supercoiling in chromatin structure and gene expression. Crit. Rev. Euk. Exp., 2, 165–209. 11. Lenfant, F., Mann, R.K., Thomsen, B., Ling, X., and Grunstein, M. (1996) All four core histone N-termini contain sequences required for the repression of basal transcription in yeast. EMBO J., 15, 3974–3985. 12. Smith, M.M., Yang, P., Santisteban, M.S., Boone, P.W., Goldstein, A.T., and Megee, P.C. (1996) A novel histone H4 mutant defective in nuclear division and mitotic chromosome transmission. Mol. Cell. Biol., 16, 1017–1026. 13. Wechsler, M.A., Kladde, M.P., Alfieri, J.A., and Peterson, C.L. (1997) Effects of Sinversions of histone H4 on yeast chromatin structure and function. EMBO J., 16, 2086–2095. 14. Kim, U.J., Han, M., Kayne, P., and Grunstein, M. (1988) Effects of histone H4 depletion on the cell cycle and transcription of Saccharomyces cerevisiae. EMBO J., 7, 2211–2219. 15. Norton, V.G., Marvin, K.W., Yau, P., and Bradbury, E.M. (1990) Nucleosome linking number change controlled by acetylation of histones H3 and H4. J. Biol. Chem., 265, 19848–19852. 16. Morse, R.H., and Cantor, C.R. (1986) Effect of trypsinization and histone H5 addition on DNA twist and topology in reconstituted minichromosomes. Nucl. Acids Res., 14, 3293–3310. 17. Thomsen, B., Bendixen, C., and Westegaard, O. (1991) Histone hyperacetylation is accompanied by changes in DNA topology in vivo. Eur. J. Biochem., 201, 107–111.

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18. Lutter, L.C., Judis, L., and Paretti, R.F. (1992) The effects of histone acetylation on chromatin topology in vivo. Mol. Cell. Biol., 12, 5004–5014. 19. Stafford, G.A., and Morse, R.H. (1997) Chromatin remodeling by transcriptional activation domains in a yeast episome. J. Biol. Chem., 272, 11526–11534. 20. Wong, J., Shi, Y.-B., and Wolffe, A.P. (1997) Determinants of chromatin disruption and transcriptional regulation instigated by the thyroid hormone receptor: hormoneregulated chromatin disruption is not sufficient for transcriptional activation. EMBO J., 16, 3158–3171. 21. Boeger, H., Griesenbeck, J., Strattan, J.S., and Kornberg, R.D. (2003) Nucleosomes unfold completely at a transcriptionally active promoter. Mol. Cell, 11, 1587–1598. 22. Korber, P., Luckenbach, T., Blaschke, D., and Horz, W. (2004) Evidence for histone eviction in trans upon induction of the yeast PHO5 promoter. Mol. Cell. Biol., 24, 10965–10974. 23. Depew, R.E., and Wang, J.C. (1975) Conformational fluctuations of DNA helix. Proc. Natl. Acad. Sci. U.S.A., 72, 4275–4280. 24. Pulleyblank, D.E., Shure, M., Tang, D., Vinograd, J., and Vosberg, H.-S. (1975) Action of nicking-closing enzyme on supercoiled and nonsupercoiled closed circular DNA: formation of a Boltzmann distribution of topological isomers. Proc. Natl. Acad. Sci. U.S.A., 72, 4280–4284. 25. Morse, R.H., and Cantor, C.R. (1985) Nucleosome core particles suppress the thermal untwisting of core DNA and adjacent linker DNA. Proc. Natl. Acad. Sci. U.S.A., 82, 4653–4657. 26. Ambrose, C., McLaughlin, R., and Bina, M. (1987) The flexibility and topology of simian virus 40 DNA in minichromosomes. Nucl. Acids Res., 15, 3703–3721. 27. Lutter, L.C. (1989) Thermal unwinding of simian virus 40 transcription complex DNA. Proc. Natl. Acad. Sci. U.S.A., 86, 8712–8716. 28. Saavedra, R.A., and Huberman, J.A. (1986) Both DNA topoisomerases I and II relax 2 m plasmid DNA in living yeast cells. Cell, 45, 65–70.

29. White, J.H., Gallo, R., and Bauer, W.R. (1989) Dependence of the linking deficiency of supercoiled minichromosomes upon nucleosome distortion. Nucl. Acids Res., 17, 5827–5835. 30. White, J.H., Gallo, R., and Bauer, W.R. (1989) Effect of nucleosome distortion on the linking deficiency in relaxed minichromosomes. J. Mol. Biol., 207, 193–199. 31. Bauer, W.R., Hayes, J.J., White, J.H., and Wolffe, A.P. (1994) Nucleosome structural changes due to acetylation. J. Mol. Biol., 236, 685–690. 32. Pederson, D.S., and Morse, R.H. (1990) Effect of transcription of yeast chromatin on DNA topology in vivo. EMBO J., 9, 1873–1881. 33. Bauer, W., and Vinograd, J. (1970) Interaction of closed circular DNA with intercalative dyes. II. The free energy of superhelix formation in SV40 DNA. J. Mol. Biol., 47, 419–435. 34. Hsieh, T.-S., and Wang, J.C. (1975) Thermodynamic properties of superhelical DNAs. Biochemistry, 14, 527–535. 35. Bates, A.D., and Maxwell, A. (1993) DNA Topology. IRL Press at Oxford University Press, Oxford, UK. 36. Morse, R.H. (1991) Topoisomer heterogeneity of plasmid chromatin in living cells. J. Mol. Biol., 222, 133–137. 37. Sambrook, J., Fritsch, E.F., and Maniatis, T. (1989) Molecular cloning: a laboratory manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. 38. Church, G., and Gilbert, W. (1984) Genomic sequencing. Proc. Natl. Acad. Sci. USA, 81, 1991–1995. 39. Drabik, C.E., Nicita, C.A., and Lutter, L.C. (1997) Measurement of the linking number change in transcribing chromatin. J. Mol. Biol., 267, 794–806. 40. Lee, C.-H., Mizusawa, H., and Kakefuda, T. (1981) Unwinding of double-stranded DNA helix by dehydration. Proc. Natl. Acad. Sci. U.S.A., 78, 2838–2842. 41. Givens, R.M., Saavedra, R.A., and Huberman, J.A. (1996) Topological complexity of SV40 minichromosomes. J. Mol. Biol., 257, 53–65.

Chapter 8 Preparation and Analysis of Uniquely Positioned Mononucleosomes Daria A. Gaykalova, Olga I. Kulaeva, Vladimir A. Bondarenko, and Vasily M. Studitsky Abstract Short DNA fragments containing single, uniquely positioned nucleosome cores have been extensively employed as simple model experimental systems for analysis of many intranuclear processes, including binding of proteins to nucleosomes, transcription, DNA repair and ATP-dependent chromatin remodeling. In many cases such simple model templates faithfully recapitulate numerous important aspects of these processes. Here we describe several recently developed procedures for obtaining and analysis of mononucleosomes that are uniquely positioned on 150–600 bp DNA fragments. Key words: Positioned nucleosome cores, chromatin, reconstitution.

1. Introduction A variety of processes in eukaryotic nuclei (such as DNA replication, repair, recombination and transcription) occur on DNA organized in chromatin. The minimal structural unit of chromatin is the nucleosome core: 147-bp DNA organized in 1 3/4 superhelical coils on the surface of the histone octamer (1). Short (150–350 bp) DNA fragments containing single, uniquely positioned nucleosome cores have been extensively employed recently for analysis of many intranuclear processes, including binding of regulatory proteins to nucleosomes (2, 3), transcription of nucleosomal templates (4–7), DNA repair in chromatin (8), ATP-dependent chromatin remodeling (9–13), and analysis of nucleosome structure (14–16). In many cases such simple model Gaykalova and Kulaeva equally contributed to this work. Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_8 Springerprotocols.com

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templates faithfully recapitulate important aspects of these processes (3, 5, 6, 8, 12, 13, 17). At the same time, this experimental system has the following advantages as compared with polynucleosomal templates: (a) Structure of single, uniquely positioned nucleosomes and changes in the nucleosome structure during various processes of DNA metabolism can be analyzed with a high resolution. (b) Electrophoretic mobility of mononucleosomes formed on a short DNA fragment (190–350 bp) in a native gel strongly depends on nucleosome position relative to DNA ends (18, 19) and on its histone composition (17, 20). Therefore positioning and histone composition of mononucleosomes that are often changed during various processes can be easily monitored by analysis in a native gel. (c) Since the initial sample usually contains only one positioned nucleosome, its fate during various processes can be determined with certainty. (d) The mobility of mononucleosomes in a native gel often is considerably changed upon binding of various protein complexes to the templates. Often conformationally different complexes having the same protein/DNA stochiometry have different mobilities in the gel (17). Thus analysis of protein binding to nucleosomes and proteininduced changes in the conformation of the complexes are possible using mononucleosomal templates. Several strategies could be used to obtain single, uniquely positioned nucleosome cores on a short DNA fragment. Typically, nucleosomes are assembled on a DNA fragment containing a nucleosome-positioning DNA sequence. Natural positioning sequences are not very strong (21). Therefore for DNA fragments that are 150–350-bp long, the result of nucleosome reconstitution usually is a mixture of single, differently positioned nucleosomes. Uniquely positioned nucleosomes can be obtained by reducing the length of DNA fragment to 150 bp (5, 14). If nucleosome-free DNA region has to be present on the template, the nucleosomes formed on 150-bp DNA fragment can be ligated to the nucleosome-free DNA fragment (5). Using this approach, uniquely positioned nucleosomes can be formed even on a random DNA sequence (22). Alternatively, single, differently positioned nucleosomes can be formed on a longer DNA fragment (150–350 bp), and then the templates containing nucleosomes positioned at the desired region of the fragment can be gel purified (23, 24). The method is based on the observation that positional isomers of nucleosome cores have different electrophoretic mobilities (19, 23, 25). A disadvantage of this approach is a low yield of the desired complexes (23). Most of the technical problems associated with use of weak nucleosome-positioning DNA sequences can be avoided by employing stronger nucleosome-positioning DNA sequences. Particularly strong nucleosome-positioning sequences have been obtained using SELEX approach, starting with a pool of synthetic random-sequence DNA molecules (26, 27). Many of these

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sequences have affinities for histone binding that are more than six-fold higher than those of the strongest natural positioning sequences, and more than 100-fold higher than the affinity of an ‘‘average’’ DNA sequence (27). The following properties make the high-affinity sequences very useful in vitro: (a) The main advantage of using these sequences is ability to form single, uniquely positioned nucleosomes on the vast majority of DNA molecules present during nucleosome reconstitution (on DNA fragments of 150–600 bp in size). In most cases obtained nucleosome preparations are homogeneous and therefore can be studied without further purification. (b) Knowledge of the consensus DNA sequences determining high DNA-histone affinity and nucleosome positioning (28, 29) allows intelligent design of the templates with desired nucleosome positioning. Since the consensus sequences are localized within the central 70-bp, H3/H4 tetramer-binding region of nucleosomal DNA, the sequences of nucleosomal DNA binding H2A/H2B histone dimers can be modified if needed. (c) Uniquely positioned H3/H4 tetramers can be formed (22). (d) Finally, polynucleosomes containing an array of precisely positioned nucleosomes can be assembled (30). Several methods for reconstitution of nucleosome cores on DNA have been described (31). In our laboratory we commonly use two methods for nucleosome assembly on the high-affinity DNA sequences. One involves incubation of purified DNA and core histones at high salt followed by reduction in the salt concentration by dialysis. At intermediate salt concentration H3/H4 bind to DNA as a tetramer and at a lower salt concentration binding of two H2A/H2B dimers results in formation of nucleosome cores (32). In the second approach, the histone octamers are transferred from ‘‘donor’’ chromatin onto DNA (33); however the resulting nucleosome preparation contains some amount of donor chromatin that could affect some reactions. For example, in our laboratory nucleosomes obtained by the transfer method have to be purified from donor chromatin after formation of transcription elongation complexes and their immobilization on a resin (34). Importantly, in both protocols some excess of competitor DNA or donor chromatin is present. The presence of the competitor allows formation of nucleosomes only on the high-affinity DNA sequences of the DNA fragment. Two methods are the most appropriate for analysis of nucleosome positioning on the high-affinity DNA sequences: low resolution mapping with restriction enzymes and high-resolution mapping using DNase I or hydroxyl radical footprinting (35). Both methods are based on the resistance of nucleosomal DNA to external probes. In the first method, isolated cores are incubated with a restriction enzyme present in the amount sufficient for complete digestion of free DNA and then analyzed in a nucleoprotein gel (36, 37). This method allows rapid mapping of multiple positioned nucleosomes but the resolution is relatively

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low (10 bp). It is most useful for quantitative information on the fraction of cores assembled in each position on a DNA fragment, but not for high-resolution analysis of nucleosome positioning. The second method is more involved and requires highly homogeneous preparation of uniquely positioned nucleosomes, but it allows determination of nucleosome positioning with singlenucleotide resolution. Below we describe several procedures for obtaining and analysis of mononucleosomes that are uniquely positioned on 150–600 bp DNA fragments containing sequences having high affinity to core histones H3/H4 and directing precise nucleosome positioning (28, 29).

2. Materials 2.1. Miscellaneous Items

1. Dialysis membranes (Spectra/Por; molecular weight cut-off of 8000 and 12,000–14,000). 2. G-25 Quick spin columns (Boehringer Mannheim). 3. Siliconized Eppendorf tubes (PGC Scientific). 4. 3 MM chromatography paper (Whatman) 5. QIAquick Gel Extraction kit (Qiagen). 6. Chicken blood (Truslow Farms). 7. CM C-25 Sephadex. 8. Centiprep-30.

2.2. Enzymes

1. T4 polynucleotide kinase (New England Biolabs, NEB). 2. Restriction enzymes (NEB). 3. Micrococcal nuclease (MNase, Worthington, 10,000 U/ml). 4. Klenow fragment of Escherichia coli DNA polymerase I (NEB). 5. Taq DNA polymerase (NEB) 6. DNase I (Sigma)

2.3. Buffers and Solutions

1. PBS buffer: phosphate-buffered saline (Gibco BRL). 2. Buffer A: 15 mM Tris–HCl (pH 7.5), 15 mM NaCl, 60 mM KCl, 0.15 mM spermine, 0.5 mM spermidine, 15 mM mercaptoethanol, 0.34 M sucrose, and 0.1 mM PMSF. 3. Buffer B: 10 mM Tris–HCl (pH 7.5), 350 mM NaCl, 0.5 mM EDTA, and 0.1 mM PMSF. 4. NLB (nuclei lysis buffer): 0.25 mM EDTA and 0.1 mM PMSF.


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5. TAE buffer: 0.04 M Tris–acetate, pH 8.0, and 1 mM EDTA. 6. HE buffer: 10 mM Na–HEPES, pH 8.0, and 1 mM EDTA. 7. CRB 1–6 (core reconstitution buffers): all six buffers contain HE, 5 mM mM -mercaptoethanol, 0.1% NP-40, and NaCl at the following concentrations: buffer 1 –2 M, 2 – 1.2 M, 3 – 1 M, 4 – 0.8 M, 5 – 0.6 M, and 6 – 0.01 M. 8. SP6 buffer: 45 mM Na–HEPES, pH 8.0, 6 mM MgCl2, 2 mM spermidine, 2 mg/ml BSA, and 10 mM -mercaptoethanol. 9. 1 TB (transcription buffer): 20 mM Tris–HCl (pH 8.0), 5 mM MgCl2, 2 mM -mercaptoethanol, and indicated concentration of KCl, mM. 10. 5 DSS (DNase I stop solution): 250 mM EDTA and 1% SDS. 11. 40 Fe (II) stock solution for hydroxyl radical reaction: 80 mM ammonium iron (II) sulfate hexahydrate, (NH4)Fe(SO4)2 6H2O. 12. 100 EDTA stock solution: 0.4 M Na–EDTA. 13. 50 H2O2 stock solution: 30% hydrogen peroxide. 14. 5 Ascorbate stock solution: 0.1 M sodium salt of L-ascorbic acid. 15. 1 RLB (RNA loading buffer): 95% formamide, 10 mM EDTA, 0.1% SDS, and 0.01% of each bromophenol blue and xylene cyanol dyes. 16. 10 HRSS (hydroxyl radical stop solution): 0.1 M thiourea, 0.3 M Na–acetate, 30 mM EDTA, and 2 mg/ml glycogen. 17. 4 Chromatin loading buffer: 100 mM Tris–HCl (pH 7.5), 40 mM EDTA, 40% sucrose, and 1 mg/ml of Salmon Testes DNA. 2.4. Proteins

1. H2A/H2B and H3/H4 histone pairs isolated from adult chicken blood by chromatography on hydroxyapatite (32, 38).

3. Methods 3.1. Preparation of Short DNA Templates Containing a Strong NucleosomePositioning Sequence

1. Design the short DNA template (see Note 1). 2. 50 -end-label one of the PCR primers with [32P] ATP (6000 Ci/mmol) using T4 polynucleotide kinase (see Note 2). 3. PCR-amplify DNA fragments in 500 ml volume (5 100 ml reactions) using Taq DNA polymerase (New England Biolabs).

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4. Resolve the obtained DNA fragments in a 1.5 % (w/v) agarose gel containing 0.5 mg/ml ethidium bromide and TAE buffer at 4–6 V/cm for 1.5–3 h, depending on the resolution required for clear band separation. 5. Using a long wavelength UV lamp (to reduce nicking of DNA), identify and excise the required band(s). 6. Purify the fragment using QIAquick Gel Extraction kit (Qiagen). 7. Extract the samples once with one volume of 1:1 (v/v) phenol:chloroform. 8. Precipitate DNA with three volumes of ethanol, wash with 70% (v/v) ethanol, dry, and dissolve in 100 ml of HE buffer. 9. Determine DNA concentration by measuring the A260 (using A260 ¼ 20 for 1 mg/ml DNA) and store at 20C. 3.2. Preparation of Donor Chromatin from Chicken Erythrocytes

3.2.1. Red Cell Isolation

The protocol described below is a modified version of the method published earlier (39) (see Note 3). 1. Collect red cells from 200 ml chicken blood by centrifugation at 1800 g for 10 min at 4C. 2. Carefully remove white cells forming from the top of the pellet. 3. Resuspend cells in 48 ml PBS buffer. 4. Collect red cells by centrifugation at 3000 g for 5 min at 4C. 5. Repeat steps 3 and 4 two more times.

3.2.2. Nuclei Isolation

1. Conduct all manipulations at 4C with pre-cooled buffers, unless indicated otherwise. 2. Resuspend cells in PBS buffer (half of the volume of the pellet). 3. Add 10 volumes of buffer A supplemented with 0.5% NP-40 and mix by inversion. 4. Collect nuclei by centrifugation at 12,000 g for 10 min at 4C. 5. Resuspend nuclei in 200 ml of buffer A (no NP-40). 6. Collect nuclei by centrifugation at 12,000 g for 10 min at 4C. 7. Repeat steps 3 and 4 several times until red color disappears. 8. Resuspend purified nuclei in a small volume (50–100 ml) of buffer A. 9. Resuspend 1 ml of nuclei in 0.9 ml of HE buffer, add 0.1 ml of 10% SDS, and measure A260. Concentration of nuclei should be 200–400 A260/ml.

3.2.3. Chromatin Preparation

1. Adjust nuclei concentration to 100 A260/ml.


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2. For analytical digestion with micrococcal nuclease (MNase) warm 1 ml of nuclei to 37C. 3. Add MgCl2 and CaCl2 to 1 mM final concentration. 4. Add 1 ml of MNase to 10 U/ml final concentration and incubate at 37C. 5. Remove 0.1 ml aliquots after 1, 2, 4, 8, 15, 30, and 60 min and stop the digestion by adding EDTA to 10 mM and SDS to 1% final concentration. 6. Extract the samples once with one volume of 1:1 (v/v) phenol:chloroform. 7. Precipitate DNA with three volumes of ethanol, wash with 70% (v/v) ethanol, dry, and dissolve in HE buffer. 8. Analyze DNA in 1% agarose gel and identify the digestion point where the sizes of DNA fragments are 3–20 kb (Fig. 8.1, 15 min digestion point was selected). 9. For preparative digestion with MNase warm the nuclei in a 500 ml conical glass flask to 37C. 10. Add MgCl2 and CaCl2 to 1 mM final concentration, MNase to 10 U/ml final concentration, and incubate at 37C for the required time. 11. Stop the digestion by adding EDTA to 10 mM final concentration. 12. Collect nuclei by centrifugation at 12,000 g for 10 min at 4C. 13. Estimate the volume of the pellet and resuspend the nuclei in equal volume of NLB buffer (nuclei should become semitransparent). 14. Collect nuclear debris by centrifugation at 12,000 g for 10 min at 4C. MNase, min: 1 2 4 8 15 30 60

M

Fig. 8.1. Time course of digestion of chicken erythrocyte nuclei with micrococcal nuclease (MNase). Nuclei were digested for various time intervals (1, 2, 4, 8, 15, 30, or 60 min), DNA purified, and analyzed in 1% agarose gel (ethidium bromide stain). A 15 min digestion point was selected for preparative digestion of the nuclei (indicated by arrow). M – 100-bp DNA ladder (New England Biolabs).

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15. Remove the supernatant containing soluble chromatin (it should be slightly opaque & gray). Discard the pellet. 16. Resuspend an aliquot of chromatin in 0.9 ml of HE buffer, add 0.1 ml of 10% SDS, and measure A260. Concentration of chromatin should be 50–80 A260/ml. 1. Adjust concentration of chromatin to 50–100 A260/ml. 2. Pre-soak CM C-25 Sephadex (Pharmacia, 36 mg of resin per mg of chromatin) in buffer B for 1 h. 3. Slowly add 2 M NaCl to chromatin to 0.35 M final concentration (see Note 4). 4. Add 1/3 of pre-soked CM C-25 Sephadex to chromatin and slowly stir for 2 h at 4C. 5. Collect the resin by centrifugation at 12,000 g for 10 min at 4C. Remove supernatant containing soluble chromatin. 6. Repeat steps 4 and 5 two more times. 7. Analyze protein composition of –H1/5 chromatin by SDS–PAGE (Fig. 8.2). Dissolve 10 mg of histones in 10–60 ml of Laemmli loading buffer. Electrophorese histones in a 18% (acrylamide:bis ¼ (30:0.15)) Laemmli gel (17 17 0.15 cm) for 5–6 h at 32 mA. Stain the gel with Coomassie Blue. 8. Concentrate –H1 chromatin to 100 A260/ml (5 mg/ml) on Centriprep-30 by centrifugation at 1300 g for required time at 4C.

-H1/5 chromatin

9. Dialyze –H1/5 chromatin overnight against buffer B and store at –70C (see Note 5). +H1/5 chromatin

3.2.4. Removal of Linker Histones H1/H5 from Chromatin

H1/5

H3 Core H2A/H2B histones H4 M

Fig. 8.2. Analysis of protein composition of histone H1/5-containing and H1/5-depleted chicken erythrocyte chromatin. Histones were resolved by 18% Laemmli SDS–PAGE. Positions of histones on the gel are indicated. M – 10–250 kD protein markers (BioRad).


3.3. Reconstitution of Mononucleosome Cores on High-Affinity DNA Sequences

3.3.1. Reconstitution of Mononucleosome Cores Using Ppurified Core Histones

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1. Cool 500 ml each of CRB1 to CRB6 buffers to 4C. 2. Dissolve 0.5–3 mg of DNA in the CRB1 buffer. 3. Add Salmon Testes DNA to achieve two-fold weight excess over the amount of the DNA fragment and purified core histones to a molar ratio of 1.5–2.0 of core histone octamer to DNA (use 1.5 excess of H2A/H2B dimer, see Note 6). Adjust volume to 40–100 ml. 4. Dialyze against CRB1 at 4C overnight. 5. Next morning, dialyze successively against CRB2, CRB3, CRB4, and CRB5, each for 1 h. Then dialyze against CRB6 for 3 h or overnight. 6. Transfer the reconstitute to a siliconized Eppendorf tube (see Note 7) and store at 4C (do not freeze). 7. Check the quality of reconstitution by native 4.5% PAGE (23). Electrophorese the cores in a 4.5% (acrylamide:bis ¼ 40:1) gel containing 20 mM HEPES (pH 8.0), 1 mM EDTA, and 5% (v/v) glycerol (17 17 0.15 cm) for 2–3 h at 6 V/cm. 8. Dry the gel on Whatman 3MM paper and expose with a PhosphorImager screen at room temperature for 2–3 h or overnight if necessary. Assembled nucleosomes are expected to migrate as a single band in the gel (Fig. 8.3).

3.3.2. Reconstitution of the Mononucleosome Cores Using Donor –H1 Chromatin

1. Cool 500 ml each of CRB4 to CRB6 buffers to 4C. 2. Mix 1–3 mg of the DNA fragment with long –H1/H5 donor chromatin at a ratio of 1:5 (w:w) in 0.04–0.1 ml of CRB3 buffer (see Note 8). 3. Dialyze successively against CRB4 and CRB5, each for 2 h at 4C. Then dialyze the sample against CRB6 for 3 h or overnight.

Nucleosome

157-bp DNA M

Fig. 8.3. Analysis of end-labeled 157-bp DNA and nucleosomes by native 4.5% PAGE. Nucleosomes are analyzed after reconstitution on the 601 high-affinity DNA sequence (27) without further purification. Note that nucleosome preparations are homogeneous and do not contain considerable amount of contaminating histone-free DNA. Positions of DNA and nucleosomes in the gel are indicated. M – end-labeled pBR322-Msp I digest (New England Biolabs).

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4. Transfer the reconstitute to a siliconized Eppendorf tube (see Note 7) and store at 4C (do not freeze). 5. Check the quality of reconstitution by analysis by native 4.5% PAGE (23). Assembled nucleosomes are expected to migrate as a single band. 3.4. Mapping the Positions of Nucleosome Cores

3.4.1. Mapping with Restriction Enzymes

1. Digest 20 ng (DNA) of labeled cores in 20 ml of SP6 buffer with 5–10 U of corresponding restriction enzyme at 37C for 20 min (see Note 9). If needed, add labeled DNA to the sample as an intrinsic control for enzyme activity. 2. Stop the reaction by adding 1/3 volume of 4 chromatin loading buffer. Analyze the digests by native 4.5% PAGE. 3. Dry the gel on Whatman 3MM paper and expose with a PhosphorImager screen at room temperature for 2–3 h or overnight if necessary. An example of restriction enzyme mapping of nucleosome core positions is shown in Fig. 8.4.

AflIII StyI

AluI

EagI 220 bp

601

M

-

Af S Al E

Nucleosome

DNA

Fig. 8.4. Analysis of nucleosome positioning on 220 bp DNA fragment using the restriction enzyme sensitivity assay. The DNA fragment contains the 601 high-affinity DNA sequence (27). If nucleosomes are positioned symmetrically, they have the same mobilities in a native gel, but are differentially sensitive to restriction enzymes and therefore can be resolved in a native gel after incubation with the enzymes. Top: restriction map of the 220 bp 601 template. The template was end-labeled (indicated by the asterisk). The cleavage sites for AflIII, StyI, AluI, and EagI and the expected position of nucleosome core (oval ) are indicated. Bottom: analysis of nucleosome cores in the native 4.5% PAGE. Mobilities of nucleosomes and histone-free DNA are indicated. Cores were digested with AflIII, StyI, AluI, or EagI as indicated. M, end-labeled MspI digest of pBR322. Note that some amount of histone-free DNA was added to nucleosomal templates to serve as an internal control for the efficiency of digestion. All enzymes completely digested histone-free DNA.


3.4.2. Mapping with DNase I endonuclease

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1. Use 20–100 ng (DNA) of DNA-end-labeled nucleosome or histone-free DNA (10,000–50,000 cpm) per reaction in 20 ml of TB40 buffer (see Note 10). 2. Add 0.1–0.2 units of DNase I per 100 ng of total DNA in the reaction (including Shredded Salmon Sperm DNA that could be present in the reconstitute). Incubate for 30 s at 37C (see Note 11). 3. Stop reaction by adding ¼ volumes of 5 DSS. 4. Adjust volume to 0.1 ml and extract the samples once with one volume of 1:1 (v/v) phenol:chloroform. 5. Precipitate DNA with three volumes of ethanol, wash with 70% (v/v) ethanol, dry, and dissolve in 5 ml of RLB buffer. Denature samples at 95C. 6. Electrophorese DNA in an 8% (acrylamide:bis ¼ 19:1) gel containing 8 M urea and 0.5 TBE buffer (17 34 0.03 cm) for 40 min at 2000 V. 7. Dry gel on Whatman 3MM paper and expose to a PhosphorImager screen at room temperature overnight. An example of DNase I footprinting of a positioned nucleosome is shown in Fig. 8.5A.

3.4.3. Mapping with Hydroxyl Radicals

1. The efficiency of DNA cleavage by hydroxyl radicals does not depend on DNA concentration, but is strongly inhibited by some chemicals: MgCl2 (> 20 mM), glycerol (>0.1%), and mercaptoethanol (>1 mM) (35). Therefore before hydroxyl radical footprinting the samples have to be dialyzed against a buffer that does not contain these chemicals. 2. Use 20–100 ng (DNA) of DNA-end-labeled nucleosomes or histone-free DNA (10,000–50,000 cpm) per reaction in 50 ml of HE buffer (the amount of nucleosomes should be twice the amount of histone-free DNA). 3. Prepare the following solutions just before the experiment: (a) 2 mM ammonium iron (II) sulfate hexahydrate – 4 mM EDTA, (b) 0.6% hydrogen peroxide, and (c) 20 mM sodium ascorbate. 4. Put siliconized Eppendorf tubes containing nucleosomes or histone-free DNA on the side and place 3.3-, 6.6-, or 9.9 ml drops each of the solutions on the wall of the tube. Quickly mix all components and incubate 2 min at room temperature (see Note 12). 5. Stop reactions by adding 6.6, 7.7 or 8.8 ml of 10 HR stop solution. 6. Adjust volume to 100 ml and extract the samples once with one volume of 1:1 (v/v) phenol:chloroform.

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A

DNase I DNA Nucl M

256 bp

B

OH radical DNA Nucl M

217 bp

Fig. 8.5. Fine mapping of translational positions of 605 nucleosome cores (27) and rotational orientation of DNA on the surface of the octamer using footprinting. A. DNase I footprinting. A 256 bp histone-free end-labeled DNA fragment or nucleosome cores were incubated with DNase I, DNA purified, and analyzed by denaturing PAGE. The predominant position of nucleosome core is indicated by oval. M, end-labeled MspI digest of pBR322. B. Hydroxyl radical footprinting. A 217 bp histone-free end-labeled DNA fragment or nucleosome cores were incubated in the presence of hydroxyl radicals, DNA purified, and analyzed by denaturing PAGE. Note that the expected 10 bp periodicity of DNA cutting is not immediately obvious after treatment with DNase I because of the sequence specificity of the enzyme. At the same time, hydroxyl radicals produce clear 10 bp periodic pattern allowing determination of precise rotational orientation of DNA on the surface of the octamer.

7. Precipitate DNA with three volumes of ethanol, wash with 70% (v/v) ethanol, dry, and dissolve in 5 ml of RLB buffer. Denature samples for 1–2 min at 95C. 8. Electrophorese DNA in an 8% (acrylamide:bis ¼ 19:1) gel containing 8 M urea and 0.5 TBE buffer (17 34 0.03 cm) for 40 min at 2000 V. Use sequencing markers if high accuracy is needed. 9. Dry gel on Whatman 3MM paper and expose to a PhosphorImager screen at room temperature overnight. An example of hydroxyl radical footprinting of a positioned nucleosome is shown in Fig. 8.5B.


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4. Notes 1. A strong nucleosome-positioning sequence can be placed at any location within the DNA fragment. A single, precisely positioned nucleosome can be formed on DNA fragments up to at least 600 bp long. 2. For footprinting studies only one DNA strand must be endlabeled. The 50 -end of the fragment can be end labeled by the protein kinase and the 30 -end – by the Klenow fragment (23). To determine rotational orientation of DNA on the surface of the histone octamer, two nucleosome preparations containing different labeled DNA strands need to be prepared. 3. Donor chromatin isolated from chicken erythrocytes contains mostly unmodified core histones (39). The main advantage of using this source of histones is the presence of low amount of proteases in chicken erythrocytes. 4. Chromatin becomes insoluble at 150 mM NaCl and then becomes soluble again when the concentration of NaCl reaches 350 mM. 5. Donor –H1 chromatin could be further purified by chromatography on Sephacryl S-400HR (Pharmacia) (33). 6. The presence of the excess of H2A/H2B histones guarantees formation of complete nucleosomes. We found that at stochiometric ratio of H2A/H2B to H3/H4 histones some amount of incomplete nucleosomes (primarily missing one H2A/H2B dimer) is formed. 7. Use of siliconized tubes is essential to prevent nucleosome disruption during storage at low concentration (1–10 mg/ ml). Presumably, siliconization prevents irreversible adsorption of histones to the walls of the tube. 8. The ratio of DNA to donor chromatin may need to be adjusted because different chromatin preparations produce slightly different results. 9. Recognition site for the restriction enzyme must be present within the region covered by nucleosome in the desired position. We found that restriction enzyme sites are protected even when they are located 5 bp in the nucleosome core (31). 10. For both DNase I and hydroxyl radical footprinting techniques it is essential to compare the results of footprinting of nucleosomes and histone-free DNA (Fig. 8.5). Therefore end-labeled nucleosomes and histone-free DNA samples should be prepared in parallel. 11. Addition of different amounts of DNase I results in variable extents of DNA cleavage. The fraction of DNA molecules

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containing the cleavages should not exceed 30% to guarantee that only one cleavage per molecule is introduced. 12. Addition of different amounts of the reagents results in variable extents of DNA cleavage. The fraction of DNA molecules containing the cleavages should not exceed 30% to guarantee that only one cleavage per molecule is introduced.

Acknowledgments We thank John Widom for plasmids containing the nucleosomepositioning sequences. This work was supported by NIH grant GM58650 to V.M.S.

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9. Wittmeyer, J., Saha, A., and Cairns, B. (2004) DNA translocation and nucleosome remodeling assays by the RSC chromatin remodeling complex. Meth. Enzymol. 377, 322–343. 10. Lorch, Y., and Kornberg, R. D. (2004) Isolation and assay of the RSC chromatinremodeling complex from Saccharomyces cerevisiae.Meth. Enzymol. 377, 316–322. 11. Eberharter, A., Langst, G., and Becker, P. B. (2004) A nucleosome sliding assay for chromatin remodeling factors. Meth. Enzymol. 377, 344–353. 12. Kassabov, S. R., and Bartholomew, B. (2004) Site-directed histone-DNA contact mapping for analysis of nucleosome dynamics.Meth. Enzymol. 375, 193–210. 13. Mizuguchi, G., Shen, X., Landry, J., Wu, W. H., Sen, S., and Wu, C. (2004) ATP-driven exchange of histone H2AZ variant catalyzed by SWR1 chromatin remodeling complex. Science 303, 343–348. . 14. Dyer, P. N., Edayathumangalam, R. S., White, C. L., Bao, Y., Chakravarthy, S., Muthurajan, U. M., and Luger, K. (2004) Reconstitution of nucleosome core particles from recombinant histones and DNA. Meth. Enzymol. 375, 23–44. 15. Luger, K., Rechsteiner, T. J., and Richmond, T. J. (1999) Preparation of nucleosome core particle from recombinant histones. Meth. Enzymol. 304, 3–19. 16. Hanson, B. L., Alexander, C., Harp, J. M., and Bunick, G. J. (2004) Preparation and


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Chapter 9 Monitoring DNA Breaks in Optically Highlighted Chromatin in Living Cells by Laser Scanning Confocal Microscopy Michael J. Kruhlak, Arkady Celeste, and Andre´ Nussenzweig Abstract The recognition and repair of DNA lesions occurs within a chromatin environment. Genetically tagging fluorescent proteins to DNA damage response proteins has provided spatial and temporal details concerning the establishment of biochemical subnuclear regions geared toward metabolizing genomic lesions. A specific marker for chromatin regions containing DNA breaks is required to study the initial dynamic structural changes in chromatin when DNA breaks occur. Here we present the experimental protocols used to investigate the dynamics of chromatin structure immediately after the simultaneous photoactivation of PAGFP-tagged core histone H2B and introduction of DNA breaks using UVA laser microirradiation on a laser scanning confocal microscope. Key words: DNA breaks, histones, chromatin, PAGFP, photoactivation, confocal microscopy, and UVA laser microirradiation.

1. Introduction The appearance of DNA breaks presents severe implications for the integrity of the genome. A double-strand break is thought to be defining initial event that leads to a chromosomal translocation, which has the potential to transform a normal cell into the unstable tumorigenic state. Studying the cell biology of DNA breaks has focused largely on the proteins involved in sensing and metabolizing the genomic lesions. Physically, DNA breaks occur in a chromatin environment, therefore, it is of interest to determine how this environment is modulated upon the introduction of DNA breaks. It is known that the biochemical environment is altered as specific proteins are recruited (1) and post-translational modifications Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_9 Springerprotocols.com

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occur on a multitude of proteins, as in the phosphorylation of the core histone variant H2AX (2, 3), now a hallmark indicator of the presence of DNA breaks. This modification of a chromatin structural component also emphasizes the need to understand how the chromatin environment is modified upon the introduction of DNA breaks. Biochemical studies have provided valuable evidence for how large scale chromatin structure is influenced by DNA lesions (4, 5). However, the spatio-temporal context of these changes can only be provided by cell biology approaches, in particular, live-cell imaging. Tagging DNA damage response proteins with genetically encoded fluorescent proteins has allowed researchers to monitor the redistribution and temporal recruitment of response proteins to sites of DNA breaks (6, 7). However, there is a lag period from the initial introduction of the experimentally induced DNA break until the steady-state accumulation of the protein at the DNA damage site. Furthermore, the multitude of response proteins display different recruitment kinetics and localize to distinct subcompartments in the DNA lesion environment (8) and, therefore, not all proteins are representative of the chromatin structure surrounding the lesion. Expression of GFP-tagged proteins has also allowed the application of photobleaching techniques to measure the kinetics of proteins in the spatial and biochemical context of the DNA breaks (6, 9). Photobleaching of fluorescent proteins in living cells involves directed exposure of a subset of fluorescent proteins to a short high intensity pulse of excitation light that photochemically alters the fluorophore resulting in the irreversible loss of emission light from the fluorescent protein, but does not physically disrupt the structural or biochemical activity of the tagged protein (10, 11). Photobleaching a subset of GFP-tagged proteins allows one to monitor the steady-state dynamics of the protein of interest by observing the redistribution of photobleached and non-photobleached proteins within a specific region of the cell. The inverse and complementary approach is to optically highlight a subset of fluorescent proteins and monitor the dynamics of the highlighted proteins (10). The advent of the photoactivatable variant of GFP, PAGFP (12) is a good example, where the non-activated PAGFP does not emit green fluorescence when exposed to 488 nm light, whereas upon a short high intensity pulse of 400 nm light the protein is covalently photoconverted to the photoactivated state that now absorbs 488 nm light to successfully emit green fluorescence. Only those PAGFP molecules exposed to the pulse of 400 nm light are photoactivated and emit green fluorescence when excited by 488 nm light. Thus PAGFP can be used as a highlighter protein in living cells (13).

Monitoring DNA Breaks in Chromatin

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Multiple methods have been employed to experimentally introduce DNA breaks within living mammalian cells to study the cell biological response to such genomic insults, including, but not limited to, ionizing radiation and laser microirradiation (2, 14–16). Laser scanning confocal microscopes equipped with continuous wave UVA lasers have been used to introduce DNA breaks in living cells (16, 17). However, most LSCMs equipped with continuous wave UVA lasers provide wavelengths that fall outside the efficient absorbance spectrum of the DNA molecule (18). Therefore, it is necessary to pre-sensitize cells with biochemical agents that either bind to the DNA molecule, such as the cell permeant Hoechst 33342 minor groove DNAbinding dye that causes minimal disruption to the structure of the DNA molecule (19), or those that are incorporated into the DNA molecule during DNA replication, such as the halogenated analog of deoxyuridine, BrdU (20–22). The presence of the DNA sensitizing agent facilitates the introduction of DNA breaks by UVA laser microirradiation using laser lines commonly found on confocal microscopes (351 and 364 nm) or laser microdissection microscopes (337 or 355 nm). The use of a laser scanning confocal microscope to introduce DNA breaks provides the distinct advantage of being able to subsequently image the specimen with high spatial and temporal resolution immediately after the introduction of DNA breaks (17). However, laser microdissection microscopes, which employ a pulsed UVA laser, are not optimally set up for high-resolution fluorescence imaging. DNA is organized into chromatin structure with the nucleosome being the fundamental repeating unit, consisting of two molecules each of the core histones H2A, H2B, H3, and H4. The core histones are conserved stable structural components of the nucleosome that exhibit slow steady-state exchange (23, 24), under normal growth conditions. Here we describe a method used to optically highlight and subsequently monitor chromatin in specific subnuclear regions containing DNA breaks in living cells with high spatial and temporal resolution. Development of the method required the ability to introduce DNA breaks in desired subnuclear regions; a specific stable marker for chromatin; that the chromatin marker highlight particular regions containing DNA breaks; and capability to visualize the subnuclear regions in real time in living cells. Thus, we combined UVA (364 nm) laser microirradiation and simultaneously photoactivation of H2B-PAGFP expressing cells pre-sensitized with the DNA-binding dye Hoechst 33342 with laser scanning confocal microscopy to achieve our goals. Here we will describe in detail how the experiments are set up and performed.

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2. Materials 2.1. Cell Culture

1. Wild-type mouse embryo fibroblasts established from day 13.5 C57BL/6 embryos. 2. Dulbecco’s-modified Eagle’s medium (DMEM; Invitrogen) supplemented with 10% fetal calf serum, 100 U/ml penicillin (Gibco/BRL), 100 mg/ml streptomycin (Gibco/BRL), and 2 mM L-glutamine (Gibco/BRL). 3. Gridded chamber coverglass (cat# P35G-1.5-7-C-grid; MatTek). 4. Fugene 6 liposome transfection reagent (Roche). 5. Plasmid DNA used for transfection, H2B-PAGFP. 6. Microscopy medium: 25 mM HEPES in phenol red free DMEM (Invitrogen) supplemented with 10% fetal calf serum, 100 U/ml penicillin (Gibco/BRL), 100 mg/ml streptomycin (Gibco/BRL), and 2 mM L-glutamine (Gibco/BRL). 7. Hoechst 33342 (Sigma) 10 mg/ml working concentration in DMSO.

2.2. Immunolabeling

1. 1 PBS (Gibco/BRL). 2. 16% paraformaldehyde solution, EM grade (Electron Microscopy Sciences): the day of the experiment freshly prepare 2% paraformaldehyde by diluting in 1 PBS. 3. 0.5% Triton X-100 in 1 PBS. 4. 1% bovine serum albumin (Sigma) in 1 PBS; 0.05% goat serum (Sigma) in 1 PBS. 5. Primary antibody mouse monoclonal anti-phospho-histone H2A.X (Ser139, clone JBW301, Upstate Biotech), 1 mg/ml stock solution. 6. Secondary antibody goat anti-mouse IgG highly crossadsorbed AlexaFluor 546 (Molecular Probes/Invitrogen), stock 2 mg/ml solution.

2.3. Laser Scanning Confocal Microscopy

1. Laser scanning confocal microscope equipped with UVA (364 nm) and two visible (488 and 543 nm) laser lines, and an acousto-optical tunable filter (AOTF) (see Note 1). 2. 40 C-apochromat (N.A. 1.2) water immersion lens (see Note 2). 3. Temperature-controlled stage heater (Carl Zeiss MicroImaging) (see Note 3). 4. Objective lens heater (Bioptechs) (see Note 4). 5. Microscope software with photobleaching and time series acquisition modules (see Note 5). 6. Laser power meter capable of measuring UV and visible wavelengths (the Ophir Nova by Ophir Optronics).


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3. Methods The method described below is dependent on two main contributing experimental factors. First, the transfection and pre-sensitization of the cells must be performed appropriately as to minimize alterations to the growth efficiency and biochemical stability of the cells. Overexpression of the PAGFP-tagged histone H2B will cause artifacts in the distribution and mobility of the protein, whereas inappropriate use of the Hoechst DNA-binding dye will disrupt DNA metabolic events, such as transcription, and detrimentally alter the growth state of the cells. Judicial use of both reagents will minimize both artifacts. Second, the appropriate amount of 364 nm UVA laser intensity is necessary for introducing DNA breaks. A calibration experiment is required to establish the most appropriate laser intensity needed to achieve a reasonable number of DNA breaks using UVA laser microirradiation. Artifacts will arise if the UVA laser intensity used to introduce DNA breaks is too high. Once these two experimental factors are established, running the experiments is straightforward. 3.1. Cell Culture, Transfection, and Pre-sensitization

1. Grow cells to 70% confluency in gridded chamber coverglass dishes in 2 ml of DMEM growth medium. 2. Transfect cells with H2B-PAGFP plasmid DNA using Fugene 6, as per the protocol suggested by the manufacturer, and incubate for 24 h at 37C, 5% CO2. 3. Warm microscopy medium to 37C. Add 0.75 ml of 10 mg/ml Hoechst 33342 solution for each milliliter of medium used, for a final concentration of 7.5 mg/ml of Hoechst 33342 dye. 4. Replace growth medium in chamber dishes with Hoechst dye containing microscopy medium and incubate cells for 30 min at 37C and 5% CO2 (see Note 6). Rinse the cells once with microscopy medium not containing Hoechst dye and add 2 ml per chamber dish of microscopy medium without Hoechst dye. The cells are now ready to be placed on the microscope.

3.2. Calibration of Laser Microirradiation Intensity for Introducing DNA Breaks

The purpose of this section is to record the microscope parameters and appropriate laser intensity needed to introduce DNA breaks in pre-sensitized cells. The aim is to determine the appropriate level of 364 nm laser intensity needed to introduce DNA breaks without overly damaging the cells. Exposure to laser intensity is governed by two factors: the actual output intensity of the laser available at the objective lens and the dwell time of the laser as it scans across the region of interest. There are a number of settings in the microscope software that influence these parameters; they

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will be mentioned in detail below. Once the parameters are established they should be maintained consistently throughout future experiments employing this method. Transfected cells are not absolutely necessary for the calibration step, but using them will help determine the detector settings required when running future photoactivation experiments. 1. Attach the objective heater to the 40 C-apochromat water immersion objective lens and set to 37C. 2. Attach the heated stage insert and set to 37C. 3. Add a small drop (20 ml) of distilled water on the objective lens and place the chamber coverglass with pre-sensitized cells in the heated stage and find best focus using brightfield illumination (see Note 7). Align and focus the condenser lens to establish Köhler illumination for subsequent acquisition of differential interference contrast (DIC) images. 4. Again using brightfield illumination, find a group of cells that are roughly in the same plane of focus. Record the position of the cells in relation to the grid imprinted on the coverslip. 5. Collect an initial image using a configuration suitable for collecting green fluorescence from PAGFP and corresponding DIC image. In establishing the optical configuration, a main dichroic mirror suitable for UV and 488 nm laser light must be used, at a minimum, since the 364 nm laser line will subsequently be used for laser microirradiation. Also, at this point the H2B-PAGFP will not emit green fluorescence, but it is important that the configuration be set up to allow acquisition of such fluorescence after photoactivation. The configuration that we consistently use includes a HFT UV/ 488 main dichroic mirror, a NFT 490 secondary beam splitter, and band-pass 505–550 nm emission filter. The pinhole diameter is opened slightly for live-cell imaging to provide an optical slice thickness of 1.5 mm. The DIC image is not confocal. 6. The laser power output of the Argon ion multi-line (458, 477, 488, and 514 nm) visible laser should be set to 25% maximum output with the 488 nm laser line used to image the photoactivated PAGFP. For imaging, the 488 nm laser intensity should be modulated to 3% transmission of the 25% output. The scan parameters for a 512 512 pixel frame size include a pixel dwell time of 3.2 ms (scan speed of 7 in the Zeiss LSM510 software) and optical zoom factor providing 0.22 mm X–Y pixel dimensions (Zoom factor 2 in the Zeiss LSM510 software). 7. Use the DIC image to place a ROI in the nucleus of each cell in the group. Remember to use ROIs of consistent dimensions (see Note 8).


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8. Next set up the so-called ‘‘bleaching’’ parameters that will be used to introduce DNA breaks and photoactivate the H2BPAGFP using the 364 nm UVA laser. We set the power output of the 80 mW Argon ion multi-line (351 nm and 364 nm) UV laser to 50% of maximum with the 364 nm laser line being used for UVA laser microirradiation. The AOTF-modulated transmission of the 364 nm laser line should be set to 20% of the 50% maximum output. The same scan speed (pixel dwell time of 3.2 ms) is used (see Note 9), with 10 scan iterations. The number of iterations includes the number of times the laser will scan through a given ROI (see Note 10). 9. Choose the option to scan once before exposure to the UVA laser, which will provide an image of the cells prior to introducing DNA breaks and photoactivating the H2B-PAGFP. A single post-UVA laser irradiation image is sufficient during the calibration step. 10. For the calibration procedure, move to different groups of cells, adjusting the positions of the ROIs and changing the AOTF-modulated transmission of the 364 nm laser line, with all other settings kept the same. Start by using the scale 2, 5, 10, 15, 20, 30, 40, 50, 75, and 100% transmission, with a different setting used for each new group of cells (see Note 11). Five minutes after the last group of cells has been exposed to UVA laser microirradiation, fix the cells in freshly prepared 2% paraformaldehyde in 1 PBS at room temperature. Remember to record the position of each group of cells on the gridded coverslip. 11. With the chamber coverglass dish removed and the remaining distilled water cleaned from the objective lens, use the laser power meter to measure the 364 nm laser intensity at the objective for each AOTF-modulated transmission value and record the values (Fig. 9.1) (see Note 12). 3.3. Immunolabeling of Laser Microirradiated Cells

This step is necessary to support the UVA laser intensity calibration of the microscope. The phosphorylated histone variant H2AX is a sensitive marker for the presence of DNA breaks and is used to determine whether the amount of UVA laser intensity is appropriate for introducing DNA breaks. Immunolabeling of the fixed cells can be done in the chamber coverglass dish. 1. After 5 min of fixation in 2% paraformaldehyde, rinse the cells with three successive changes of 1 PBS. 2. Permeabilize the cells by incubating with 0.5% Triton X-100 in 1 PBS for 5 min at room temperature. 3. Rinse the cells three times with 1 PBS.

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Fig. 9.1. UVA (364 nm) laser intensity measurements. The laser power, in mW, measured at the 40 C-apochromat objective lens in relation to the various AOTF-modulated percent transmission amounts of the laser light.

4. Incubate with blocking buffer for 60 min at room temperature. 5. Remove blocking buffer. 6. Add primary anti-phosphorylated H2AX antibody at a final concentration of 1 mg/ml in 1 PBS and incubate in a humidified chamber for 60 min at room temperature. 7. Rinse three times with 1 PBS. 8. Add secondary goat anti-mouse AlexaFluor 546 (see Note 13) antibody at a final concentration of 10 mg/ml and incubate in a humidified chamber in the dark for 60 min at room temperature. 9. Rinse three times with 1 PBS. 10. Leave cells in 1 PBS. 3.4. Imaging the Immunolabeled Cells

Measuring the 364 nm UVA laser intensity during the calibration step provides a quantitative measure of the amount of laser microirradiation applied to the cells. The degree of phosphorylated H2AX present will be compared to the amount of laser intensity used. Since the appearance of phosphorylated H2AX is a sensitive indicator of the presence of DNA breaks, it will be used to help empirically determine the most suitable amount of laser microirradiation. The aim is to determine the minimal amount of UVA laser microirradiation that provides a consistent phosphorylated H2AX signal. 1. Relocate the cells on the gridded chamber coverglass using the 40 C-apochromat objective lens.


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2. Choose an optical configuration suitable for exciting and collecting fluorescence for two channels, the green fluorescence from the H2B-PAGFP and the red fluorescence from the AlexaFluor 546, marking the presence of phosphorylated H2AX. A multi-track configuration (the two laser lines are actively switched on and off, but are not on simultaneously) with 488 and 543 nm laser line excitation. The power output of the 488 nm laser should be set to 30% maximum output and modulated to 10% transmission of the 30% output, whereas the HeNe 543 nm laser line should be set to maximum output and modulated at 50% transmission. The scan parameters for a 512 512 pixel frame size include a pixel dwell time of 2.56 ms (scan speed 8 in the Zeiss LSM510 software) and optical zoom factor providing 0.22 mm X–Y pixel dimensions (Zoom factor 2 in the Zeiss LSM510 software). For example, the multi-track configuration used with our microscope consistently includes a HFT UV/488/543/633 main dichroic mirror, a NFT 545 secondary beam splitter, and 505–530 nm band-pass emission filter in front of PMT detector accepting the H2B-PAGFP green fluorescence and a 560–615 nm bandpass emission filter in front of a second PMT detector accepting the AlexaFluor 546 red fluorescence. The pinhole diameter is set to one Airy unit which provides an optical slice thickness of 1.0 mm. A DIC image is also collected. 3. Move to the distinct groups of cells exposed to different amounts of UVA laser microirradiation while noting the presence of phosphorylated H2AX (Fig. 9.2). If the nucleus displays phosphorylated H2AX throughout the entire nucleus, beyond the ROI exposed to UVA laser microirradiation, that is a clear indication that the amount of UVA laser microirradiation used was too high and needs to be reduced. 3.5. Imaging Photoactivated Chromatin Containing DNA Breaks

Now that the appropriate microscope parameters suitable for UVA laser microirradiation have been determined, the actual time-lapse experiments can be performed (Fig. 9.3). 1. Place the transfected H2B-PAGFP expressing cells in the gridded chamber coverglass dish on the heated 40 C-apochromat water immersion lens. 2. The heated stage insert should be set to 37C. 3. Set up the 364 nm laser microirradiation and photoactivation microscope parameters determined to be the most suitable from the calibration step. For our microscope configuration we use 50% UVA laser output, 20% AOTFmodulated transmission, pixel time of 3.2 ms (scan speed 7), optical zoom set to 0.22 mm pixel size, and 10 iterations of the ROI.

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Fig. 9.2. Calibration of the degree of phosphorylated H2AX present in relation to the amount of UVA laser intensity used for microirradiation. Several groups of wild-type MEFs were exposed to different amounts of 364 nm laser intensity. A single rectangular ROI was placed on each cell in a group and the cells subsequently microirradiated in that ROI. Confocal images of the resulting phosphorylated H2AX (g-H2AX, left column) are shown. The corresponding DIC images are also shown (right column). Scale bar, 20 mm.


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Fig. 9.3. Simultaneous photoactivation of H2B-PAGFP and introduction of DNA breaks using UVA (364 nm) laser microirradiation. (A) a schematic indicating that DNA breaks are introduced upon exposure to 364 nm laser microirradiation in specific ROIs when cells are pre-sensitized with Hoechst DNA-binding dye, whereas H2B-PAGFP is photoactivated with the same 364 nm laser intensity in the presence or absence of the Hoechst dye. (B) Time-lapse series of a single pre-sensitized wild-type mouse embryo fibroblast expressing H2B-PAGFP. In this example, three ROIs were placed in the cell nucleus, one rectangle and two circles (shown in the Pre-UV image; with the oval outline denoting the boundary of the nucleus). After fixation and immunolabeling, the same cell was relocated and confocal image collected of the resultant DNA breaks, marked by the presence of phosphorylated H2AX (g-H2AX, far right bottom panel). A corresponding DIC image is also shown. Scale bar, 20 mm.

4. Find a group of cells using brightfield illumination, as described above (see Note 14). 5. Position the ROIs. 6. Acquire a time series of images including the pre-UVA laser exposure image and multiple images post-UVA laser exposure. The frequency of image acquisition is to be determined by the temporal information desired. Be careful that the sample is not imaged too frequently that overexposure to the 488 nm laser during imaging of the photoactivated H2B-PAGFP inadvertently disrupts the cells.

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4. Notes 1. We used a Zeiss LSM510 META laser scanning confocal microscope equipped with a 80 mW Argon ion multi-line (351 and 364 nm) continuous wave UVA laser (Coherent Enterprise II 653, Coherent Laser Group, Santa Clara, CA), a 45 mW Argon ion multi-line (458, 477, 488, and 514 nm) visible laser (Lasos 60), and a 1.0 mW 543 nm HeNe laser. On this model of microscope, the UV and visible laser lines enter the scanhead from separate optical fibers and, therefore, to ensure that both UV and visible lasers are aligned the UV/V is collimator setting needs to be optimized for the objective lens used. 2. We used a Zeiss 40 C-apochromat (N.A. 1.2) lens for several of reasons: First, the use of a water immersion lens reduces refractive index mismatch between the specimen and the objective lens, thereby reducing the degree of spherical aberration in the resultant image. Second, the C-apochromat lens is corrected for multiple wavelengths helping to ensure that the 364 and 488 nm laser wavelengths are focused at the same plane in the specimen (see also Note 1). Last, the transmission efficiency of our C-apochromat lens is greater at 364 nm than for our 63 oil immersion Plan-apochromat (N.A. 1.4) lens. Overall, the optical advantages gained by using the 40 C-apochromat lens outweighed the disadvantage of using an objective lens of lower numerical aperture (N.A. 1.2 versus N.A. 1.4). 3. We used a Zeiss heated stage insert that controls only the temperature of the stage and not the humidity or CO2 levels. For short-time course experiments such as described in this article, HEPES-buffered microscopy medium is sufficient to regulate the CO2 levels. 4. We used an objective heater from Bioptechs Inc., the large size to fit over the larger diameter collar of our 40 Capochromat lens. It is necessary to use an objective heater in conjunction with the heated stage if an environmental chamber is not used. The objective lens acts as a heat sink and the use of an objective heater helps stabilize the temperature and prevent temperature-related focal drift. 5. The software controlling the microscope and image acquisition parameters must have an accompanying module that allows the adaptive placement of ROIs. The software must also have a ‘‘bleaching’’ option in the module so that scanning of the 364 nm UVA laser can be limited to a short high intensity exposure in the confined area of the ROI. The Zeiss LSM510 software (version 3.2) used in our experiments has a bleaching option where the ROI can be established (both size and


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position in the image or specimen). It also allows the user to choose the desired wavelength and AOTF-modulated transmission (laser intensity) used to scan within the ROI. Although the ‘‘bleaching’’ option is used, we actually use the parameters to photoactivate the PAGFP rather than photobleach the PAGFP. An AOTF that modulates the laser intensity during the scanning process is necessary to restrict the high intensity UVA laser pulse to the desired ROI. The bleaching option in the Zeiss software is coupled to the time series acquisition, and thereby the bleach parameters can be run immediately before the time series is acquired. 6. Various cell types take up the cell permeant Hoechst 33342 dye at different rates and exhibit different degrees of sensitivity in the presence of the dye (25). The final concentration of Hoechst dye and the length of the incubation period in the presence of the dye may need to be adjusted for certain cell types. Overall, the aim is to add sufficient dye to sensitize the cells, but not too much to compromise the biochemical stability of the cells. Generally, after the addition of the dye the cells should continue to grow in culture through multiple cell cycles. If the majority of the cells exhibit poor survival or replication defects, then the concentration of the Hoechst dye used is too great and should be reduced. Once the appropriate dye concentration and incubation time have been established for a given cell type, those conditions must be used consistently for future experiments. 7. It is imperative that exposure to UV excitation by wide-field fluorescence illumination, such as through a DAPI filter, be strictly avoided. Such exposure will lead to the inadvertent introduction of DNA breaks and will compromise the experimental results. However, since the non-photoactivated H2BPAGFP does not emit green fluorescence it is difficult to initially determine the transfection efficiency of the cells and whether it is worth continuing with further steps in the experiment. An extra chamber coverglass dish of transfected cells, not to be used for laser microirradiation, can be exposed to UV light using a DAPI filter since this exposure will also photoactivate the H2B-PAGFP and will provide some indication of the transfection efficiency of the cells. Again, this chamber dish should not be used in the laser microirradiation experiments. 8. Given the mechanics of scanning the laser across the sample, horizontal rectangular ROIs are preferable over vertical ROIs. The length (number of pixels in the X-axis) of the ROI may vary depending on the dimensions of the nucleus within individual cells, however, the thickness (number of pixels in the Y-axis) should be kept consistent.

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9. Version 3.2 of the Zeiss LSM510 software forces one to use the same scan speed in the bleaching settings as that used in the image acquisition scan settings. Newer versions of the Zeiss software provide more flexibility in choosing different scan speeds for bleaching (photoactivating) and imaging. 10. The laser power output, the AOTF-modulated transmission of the laser allowed to reach the objective lens, the speed at which the laser is scanned across the sample (pixel dwell time), the distance that the laser covers over the particular time (optical zoom), and the number of scan iterations all influence the amount of laser intensity directed at the region of the nucleus. It is important to establish a good balance between all these parameters to ensure that the appropriate amount of laser intensity is used. The parameters mentioned above were found to be suitable for our Zeiss LSM510 META confocal microscope by running the calibration described above, but they will most definitely need to be adjusted for other similar microscopes. 11. Inherently there is an unavoidable time factor involved in performing the calibration procedure and this is why it is important to start at the lower laser intensity and progress to greater laser intensity. 12. Measuring the laser intensity at the objective will provide accurate values of the laser intensity suitable for running the UVA laser microirradiation experiments. As the UVA laser ages the output decreases and so recording periodic measurements of the laser intensity at the objective will allow adjustments to be made to the parameters to ensure the appropriate amount of laser intensity is used in future experiments. 13. The UVA microirradiated cells have H2B-PAGFP, and the red fluorescence emitting AlexaFluor 546 fluorophore is used for detecting the presence of phosphorylated H2AX. Our Zeiss LSM510 META microscope has a HeNe 543 laser line, and the excitation spectrum of the AlexaFluor 546 fluorophore is suited to this laser line. 14. Since the non-photoactivated PAGFP signal is very low, it is difficult to determine which cells are expressing H2B-PAGFP. To help overcome this, temporarily adjust the brightness and contrast of the displayed image. The presence of a faint green signal indicates that the cell may be expressing the protein. Return the contrast settings to normal and run the experiment. If the cell is indeed expressing the H2B-PAGFP, one will find out in the first image immediately after UVA laser microirradiation. If the activated PAGFP signal does not appear, most likely that cell is not expressing the H2B-PAGFP. Simply move on to a different cell and try again.


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Acknowledgments We thank George Patterson (NICHD/NIH) for the generous gift of the PAGFP construct and helpful technical assistance. We are grateful to Tom Misteli (NCI/NIH) for providing the H2BGFP construct. References 1. Downs, J. A., Nussenzweig, M. C., and Nussenzweig, A. (2007). Chromatin dynamics and the preservation of genetic information. Nature 447, 951–8. 2. Rogakou, E. P., Boon, C., Redon, C., and Bonner, W. M. (1999). DNA doublestranded breaks induce histone H2AX phosphorylation on serine 139. J Cell Biol 146, 905–16. 3. Rogakou, E. P., Pilch, D. R., Orr, A. H., Ivanova, V. S., and Bonner, W. M. (1998) J Biol Chem 273, 5858–68. 4. Smerdon, M. J., Kastan, M. B., and Lieberman, M. W. (1979). Distribution of repairincorporated nucleotides and nucleosome rearrangement in the chromatin of normal and xeroderma pigmentosum human fibroblasts. Biochemistry 18, 3732–9. 5. Takahashi, K., and Kaneko, I. (1985). Changes in nuclease sensitivity of mammalian cells after irradiation with 60Co gammarays. Int J Radiat Biol Relat Stud Phys Chem Med 48, 389–95. 6. Houtsmuller, A. B., and Vermeulen, W. (2001). Macromolecular dynamics in living cell nuclei revealed by fluorescence redistribution after photobleaching. Histochem Cell Biol 115, 13–21. 7. Lukas, C., Falck, J., Bartkova, J., Bartek, J., and Lukas, J. (2003). Distinct spatiotemporal dynamics of mammalian checkpoint regulators induced by DNA damage. Nat Cell Biol 5, 255–60. 8. Bekker-Jensen, S., Lukas, C., Kitagawa, R., Melander, F., Kastan, M. B., Bartek, J., and Lukas, J. (2006). Spatial organization of the mammalian genome surveillance machinery in response to DNA strand breaks. J Cell Biol 173, 195–206. 9. Essers, J., Houtsmuller, A. B., van Veelen, L., Paulusma, C., Nigg, A. L., Pastink, A., Vermeulen, W., Hoeijmakers, J. H., and Kanaar, R. (2002). Nuclear dynamics of RAD52

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group homologous recombination proteins in response to DNA damage. Embo J 21, 2030–7. Lippincott-Schwartz, J., Altan-Bonnet, N., and Patterson, G. H. (2003). Photobleaching and photoactivation: following protein dynamics in living cells. Nat Cell Biol Suppl, S7–14. White, J., and Stelzer, E. (1999). Photobleaching GFP reveals protein dynamics inside live cells. Trends Cell Biol 9, 61–5. Patterson, G. H., and Lippincott-Schwartz, J. (2002). A photoactivatable GFP for selective photolabeling of proteins and cells. Science 297, 1873–7. Patterson, G. H., and Lippincott-Schwartz, J. (2004). Selective photolabeling of proteins using photoactivatable GFP. Methods 32, 445–50. Cremer, C., Cremer, T., Fukuda, M., and Nakanishi, K. (1980). Detection of laser – UV microirradiation-induced DNA photolesions by immunofluorescent staining. Hum Genet 54, 107–10. Meldrum, R. A., Botchway, S. W., Wharton, C. W., and Hirst, G. J. (2003). Nanoscale spatial induction of ultraviolet photoproducts in cellular DNA by three-photon near-infrared absorption. EMBO Rep 4, 1144–9. Walter, J., Cremer, T., Miyagawa, K., and Tashiro, S. (2003). A new system for laserUVA-microirradiation of living cells. J Microsc 209, 71–5. Kruhlak, M. J., Celeste, A., Dellaire, G., Fernandez-Capetillo, O., Muller, W. G., McNally, J. G., Bazett-Jones, D. P., and Nussenzweig, A. (2006). Changes in chromatin structure and mobility in living cells at sites of DNA double-strand breaks. J Cell Biol 172, 823–34. Sutherland, J. C., and Griffin, K. P. (1981). Absorption spectrum of DNA for

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wavelengths greater than 300 nm. Radiat Res 86, 399–409. Geierstanger, B. H., and Wemmer, D. E. (1995). Complexes of the minor groove of DNA. Annu Rev Biophys Biomol Struct 24, 463–93. Limoli, C. L., and Ward, J. F. (1993). A new method for introducing double-strand breaks into cellular DNA. Radiat Res 134, 160–9. Djordjevic, B., and Djordjevic, O. (1965). Chromosomal aberrations in synchronized mammalian cells treated with 5-bromodeoxyuridine and irradiated by ultra-violet light. Nature 206, 1165–6. Regan, J. D., Setlow, R. B., and Ley, R. D. (1971). Normal and defective repair of damaged DNA in human cells: a sensitive assay utilizing the photolysis of bromodeoxyuridine. Proc Natl Acad Sci U S A 68, 708–12.

23. Kimura, H., and Cook, P. R. (2001). Kinetics of core histones in living human cells: little exchange of H3 and H4 and some rapid exchange of H2B. J Cell Biol 153, 1341–53. 24. Siino, J. S., Nazarov, I. B., Svetlova, M. P., Solovjeva, L. V., Adamson, R. H., Zalenskaya, I. A., Yau, P. M., Bradbury, E. M., and Tomilin, N. V. (2002) Photobleaching of GFP-labeled H2AX in chromatin: H2AX has low diffusional mobility in the nucleus. Biochem Biophys Res Commun 297, 1318–23. 25. Fried, J., Doblin, J., Takamoto, S., Perez, A., Hansen, H., and Clarkson, B. (1982) Effects of Hoechst 33342 on survival and growth of two tumor cell lines and on hematopoietically normal bone marrow cells. Cytometry 3, 42–7.

Chapter 10 Methods to Study Transcription-Coupled Repair in Chromatin He´le`ne Gaillard, Ralf Erik Wellinger, and Andre´s Aguilera Abstract Transcription-coupled repair (TCR) is a sub-pathway of nucleotide excision repair that allows for the enhanced repair of the transcribed strand of active genes. A classical method to study DNA repair in vivo consists in the molecular analysis of UV-induced DNA damages at specific loci. Cells are irradiated with a defined dose of UV light leading to the formation of DNA lesions and incubated in the dark to allow repair. About 90% of the photoproducts consist of cyclobutane pyrimidine dimers, which can be cleaved by the DNA nicking activity of the T4 endonuclease V (T4endoV) repair enzyme. Strand-specific repair in a suitable restriction fragment is determined by alkaline gel electrophoresis followed by Southern blot transfer and indirect end-labeling using a single-stranded probe. Recent approaches have assessed the role of transcription factors in TCR by analyzing RNA polymerase II occupancy on a damaged template by chromatin immunoprecipitation (ChIP). Cells are treated with formaldehyde in vivo to cross-link proteins to DNA and enrichment of a protein of interest is done by subsequent immunoprecipitation. Upon reversal of the protein–DNA cross-links, the amount of coprecipitated DNA fragments can be detected by quantitative PCR. To perform ChIP on UV-damaged templates, we included an in vitro photoreactivation step prior to PCR analysis to ensure that all precipitated DNA fragments serve as substrates for the PCR reaction. Here, we provide a detailed protocol for both the DNA repair analysis and the ChIP approaches to study TCR in chromatin. Key words: DNA Damage, UV, cyclobutane pyrimidine dimer, nucleotide excision repair, transcription-coupled repair, chromatin immunoprecipitation.

1. Introduction DNA repair is an essential defense system that allows cells to minimize the deleterious effects of endogenous and environmental damage. Such damage may otherwise lead to persistent DNA modifications, mutations, replication errors, genomic instability, cancer, aging, or might even be lethal. Nucleotide excision repair (NER) is a major, evolutionarily conserved DNA repair pathway Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_10 Springerprotocols.com

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that fixes bulky DNA lesions, including UV-induced photoproducts, and adducts derived from cisplatin and 4-nitroquinoline oxide treatment (1, 2). A few human disorders are characterized by defective nucleotide excision repair (3). These disorders include xeroderma pigmentosum (XP), Cockayne syndrome (CS), and Trichothiodystrophy (TTD). Hypersensitivity to UV light exposure is a characteristic of all three diseases but sun exposure is accompanied by an approximate 1000-fold increase of developing skin cancer in the case of XP. Interestingly, human genes leading to these disorders are involved in the nucleotide excision repair but also include basal transcription complexes. Thus, nucleotide excision repair and transcription might cooperate in order to archive an optimized repair of the genome. Within NER two damage-sensing pathways are recognized. The major one is termed global genome repair (GGR) and is capable of repairing the entire genome, while repair of the transcribed strand of active genes is enhanced by the so called transcription-coupled repair (TCR). GGR and TCR share the core proteins of excision repair and only differ in factors involved in damage recognition. In yeast, GGR requires Rad7, a protein carrying leucine-rich repeats, and Rad16, a member of the SWI2/ SNF2 subfamily of ATPases (4). Initial work of Hanawalt and colleagues has led to a more detailed understanding of TCR in Escherichia coli discovering that TCR is mediated by a single transcription-repair coupling factor (5). In eukaryotes the precise mechanism of TCR remains poorly understood. A fundamental difference between eukaryotic and prokaryotic cells is the presence of the nuclear envelope. We have recently shown that TCR has to cope with mRNA export in eukaryotes, possibly explaining why it cannot take advantage of a single transcription-repair coupling factor as found in prokaryotic cells (6). As ongoing transcription is required for TCR, damage recognition is likely triggered by the elongating RNA polymerase (RNAP) itself, whose progression gets obstructed at the site of damage. RNAP arrests at injuries in the template strand triggering, possibly via additional specific factors, the recruitment of the DNA repair machinery (5, 7–9). The mammalian Cockayne syndrome B protein (CSB) and its yeast orthologue Rad26 share conserved functions (10, 11) and represent putative eukaryotic transcriptionrepair coupling factor (TCRF) candidates. In yeast, the Rpb9 subunit of RNAPII has also been shown to contribute to TCR (12–14). Recently, we have shown that proteins of the THO, Sub2-Yra1 and Thp1-Sac3 complexes, which are involved in mRNP biogenesis and export (15), are also required for efficient TCR (6). On the other hand, RNAPII is subjected to ubiquitylation and proteasome-mediated degradation in response to UV-generated DNA damage (16–19). It has been proposed that degradation of damage-stalled RNAPII complexes might be an alternative to TCR

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(20). Indeed, recent studies have shown that arrested RNAPII elongation complexes are the preferred substrate for ubiquitylation, which is dependent on the C-terminal repeat domain (CTD) of RNAPII and on the Def1 protein in yeast (21, 22). TCR can be studied in vitro using cell extracts or purified components. Since TCR mutants are usually not UV-sensitive unless GGR is disrupted, in vivo methods to study TCR are limited to the molecular analysis of DNA damages at specific loci. DNA repair can be analyzed at low or high resolution (23). Cells are irradiated with a defined dose of UV light leading to the formation of cyclobutane pyrimidine dimers (CPDs) and 6–4 photoproducts (6–4 PP) and incubated in the dark to allow repair. It is important to avoid light activation of the photolyase repair enzyme during repair in order to ensure that only NER takes place. The majority (about 90%) of the photoproducts consist of CPDs, which can be cleaved by the lesion specific DNA nicking activity of the T4 endonuclease V (T4endoV) repair enzyme (24). Isolated DNA is first cleaved with suitable restriction enzymes, then treated with T4endoV and separated by electrophoresis in alkaline conditions to analyze the migration of single-stranded DNA. Strand-specific repair can be determined by Southern blot transfer and indirect end-labeling using a single-stranded, fragment-specific probe. An example is shown in which repair of the housekeeping RPB2 gene was analyzed in the mRNA biogenesis mutant sub2–206 (Fig. 10.1A). Repair is detected by the recovery of the intact DNA fragment migrating on top of a smear-like signal. Within this smear-like signal specific bands are apparent that mark the presence of DNA-damage hot-spots within the fragment. Since UV-damage is sequence specific, CPD hot-spots in the Crick strand will lead to damage cold-spots in the Watson strand and vice versa. Recovery of the intact DNA fragment at various repair time points can be measured by calculating the relative signal intensity of the intact DNA fragment versus the total signal. Repair curves are usually displayed as the percentage of repair (Fig. 10.1B). Note that in wild-type cells repair of the transcribed strand occurs much faster than repair of the non-transcribed strand. In sub2 mutants repair of the transcribed strand is slower indicating that Sub2 is required for effective TCR (6). More recent approaches have assessed the role of transcription factors in TCR by analyzing RNAPII occupancy on a damaged template by chromatin immunoprecipitation (ChIP) (6, 25, 26). The ChIP-assay was originally developed to determine the natural site of action of the chromatin-binding proteins of the POLYCOMB complex (27) and adapted for virtually any chromatinbound factor (28–30). As depicted in Fig. 10.2, proteins are cross-linked to DNA in vivo by formaldehyde treatment of cells, thus preventing redistribution of chromatin components during chromatin preparation, and enrichment of a protein of interest is

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Fig. 10.1. Example for an experiment using the method described to analyze the CPD repair rates on the transcribed (TS) and the non-transcribed (NTS) strands of the constitutively expressed RPB2 gene. (A.) Yeast cells in which the Sub2 mRNP biogenesis factor is mutated were irradiated with 230 J/m2 UV light or mock treated (no UV). Repair was analyzed by indirect end-labeling in a 4.4 kb (NsiI/PvuI) RPB2 fragment. The remaining intact restriction fragment after treatment of damaged DNA with T4endoV (+UV, +T4endoV) corresponds to the fraction of undamaged DNA. Non-irradiated DNA (–UV) and DNA not treated with T4endoV (–T4endoV) were used as controls. (B.) Graphical representation of the values obtained by quantification of the gels shown in A. Repair curves of wild-type yeast cells are shown for comparison (dash lines).

done by subsequent immunoprecipitation. Upon reversal of the protein–DNA cross-links, the amount of coprecipitated DNA fragments can be detected by quantitative PCR. In order to analyze RNAPII occupancy on a UV-damaged template by ChIP, we


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Fig. 10.2. Strategy for the mapping of chromosomal interaction sites of non-histone chromatin-associated factors after UV irradiation. Yeast cells are irradiated with UV light to generate lesions in the DNA (A.), incubated for recovery, crosslinked with formaldehyde in vivo at different time points (B.), and subsequently disrupted mechanically. Chromatin fragments released from the cells are shared by sonication (C.) to an average size suitable for the desired resolution of the mapping experiment. Antibodies recognizing a chromosomal non-histone protein are added and protein–DNA complexes recovered by absorption to protein A sepharose beads (D.). Samples from the INPUT and PRECIPITATE are heat treated to reverse the cross-links and DNA is isolated (E.). DNA damages (CPDs) are repaired by in vitro photoreactivation with E. coli photolyase (F.) and DNA is purified. The DNA is used as template in quantitative PCR reactions with primer pairs derived from different chromosomal regions (G.). Bound and unbound sequences are equally represented in the genome and will be amplified accordingly from the INPUT sample. Only sequences specifically bound by the protein of interest in vivo coimmunoprecipitate with this factor and will give rise to PCR products from the PRECIPITATE.

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included a photoreactivation step prior to PCR analysis (see Fig. 10.2) because it has been shown that UV photoproducts such as CPDs block the elongation of Taq-DNA polymerases (31). Thus, in vitro elimination of CPDs by photoreactivation ensures that all precipitated DNA fragments serve as substrates for the PCR reaction. An example is shown for wild-type and cells defective in the THO mRNP biogenesis factor (Fig. 10.3).

Fig. 10.3. Example for an experiment using the method described to analyze the association of Rpb1 (the largest subunit of RNAPII holoenzyme) with the YLR454w ORF under the control of the GAL1 promoter in UV-irradiated yeast cells. Wild-type or hpr1D yeast cells were irradiated with 230 J/m2 UV light or mock treated (no UV). After formaldehyde cross-linking at the indicated time points, cell lysates were prepared and subjected to immunoprecipitation with an antibody against the Rpb1 C-terminal domain (CTD). DNA from INPUT and PRECIPITATE were isolated, subjected to photoreactivation by photolyase to repair most of the damages, and analyzed by quantitative real-time PCR analysis using primer pairs specific for the 50 - and 30 -end of the YLR454w gene and for the 9716–9863 intergenic region of chromosome V, which was used as nontranscribed control. Standard curves for all three pairs of primers were performed for each PCR analysis, all PCR reactions being performed in triplicate. The enrichment for each PCR amplification of interest was calculated as the ratio between the region-specific signal and the intergenic signal of the PRECIPITATE normalized with respect to the corresponding ratios of the INPUT. The complete set of results and their interpretation can be found in our recent publication (6).


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2. Materials 2.1. UV Irradiation and Repair

1. Yeast cultures grown in appropriate medium. 2. Round pyrex trays (SCHOTT, 19 cm diameter). 3. BS03 irradiation chamber equipped with UV-C lamps (Philips TUV 15 W, irradiation peak at 254 nm) and UV-Mat dosimeter (Dr. Gröbel UV-Elektronik GmbH; see Note 1). 4. Medium containing appropriate amino acids at a 3 concentration. 5. Dark rooms equipped with safety lights (Philips TLD RED). 6. Refrigerated table centrifuge (Beckman Coulter, Rotor C0650).

2.2. Purification of DNA for Repair Analysis

1. Freshly made spheroplasting buffer: 1 M sorbitol, 10 mM Tris–HCl pH 7.6, 100 mM EDTA pH 8.0, 0.1% v/v betamercaptoethanol. 2. Zymolyase stock solution: 1000 U/ml (USB). 3. Solution I: 800 mM GuHCl, 30 mM EDTA pH 8.0, 30 mM Tris–HCl pH 7.6, 5% Tween-20, 0.5% Triton X-100. 4. RNase A stock solution: 10 mg/ml (Roche; see Note 5). 5. Proteinase K stock solution: 20 mg/ml (Roche). 6. NaCl stock solution: 2.5 M. 7. D/I: dichloroform/isoamylalcohol (24:1). 8. Solution II: 1% w/v CTAB (cetyltrimethylammonium bromide), 50 mM Tris–HCl pH 7.6, 10 mM EDTA pH 8.0. 9. Na solution: 500 mM NaCl, 10 mM Tris–HCl pH 7.6, 1 mM EDTA pH 8.0. 10. Isopropanol (at room temperature). 11. 70% ethanol (at room temperature).

2.3. Repair Analysis by Indirect EndLabeling

1. Appropriate DNA fragments (see Note 11). 2. Nucleotide stock solution: 10 mM dATP, 10 mM dTTP, 10 mM dGTP (Roche). 3.

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PdCTP (3000 mCi/mmol; Amersham Biosciences).

4. Taq polymerase and 10 reaction buffer (Biotools). 5. Thermocycler (Perkin–Elmer). 6. Restriction enzymes and appropriate 10 buffers (Roche). 7. Strataclean resin (Stratagene). 8. 50T5E buffer: 50 mM Tris–HCl pH 7.5, 5 mM EDTA pH 8.0.

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9. 50% v/v slurry of Sephadex G-50 (SIGMA) in 50T5E buffer (keep at 4C). 10. T4-endonuclease V (Epicentre Technologies). 11. Agarose. 12. Alkaline electrophoresis buffer: 50 mM NaOH, 1 mM EDTA pH 8.0. 13. Freshly made 5 alkaline-loading buffer: 12.5% Ficoll 400, 5 mM EDTA pH 8.0, 0.125% bromocresol green, 250 mM NaOH. 14. Denaturating solution: 1.5 M NaCl, 0.5 M NaOH. 15. Neutralizing solution: 0.5 M Tris–HCl pH 7.6, 1.5 M NaCl. 16. 20 SSC: 3 M NaCl, 0.3 M sodium citrate pH 7.0. 17. 3MM Whatman paper. 18. Filter paper. 19. Hybond N blotting membrane (Amersham Biosciences). 20. Hybridization solution: 0.25 M Na2HPO4, 7% SDS, 1 mM EDTA. 21. 20 SSPE: 3 M NaCl, 0.2 M NaH2PO4, 20 mM EDTA pH 7.4. 22. Wash solution I: 0.1 SSPE, 5 mM EDTA, 0.5% SDS. 23. Wash solution II: 0.5 SSC, 0.1% SDS. 24. Horizontal gel apparatus (OWL, gel size: 20 25 cm). 25. Peristaltic pump, glass plates. 2.4. ChIP Analysis

1. Formaldehyde 37% (Merck). 2. Glycine stock solution: 2.5 M in H2O. 3. 10 PBS: 1.37 M NaCl, 27 mM KCl, 100 mM Na2HPO4, 20 mM KH2PO4 pH 7.4. 4. Lysis buffer: 50 mM HEPES/KOH pH 7.5, 140 mM NaCl, 1 mM EDTA pH 8.0, 1% Triton X-100, 0.1% sodium deoxicholate, 1 mM benzamidine, 50 mg/ml TPCK, 0.25 mM TLCK, 1 mg/ml leupeptine, 2 mg/ml antiparine, 1 mg/ml papstatin A, 1 mg/ml aprotinin, 1 mM PMSF (see Note 16). 5. Glass beads (450–600 microns, SIGMA; see Note 17). 6. Bioruptor (Diagenode). 7. TE/SDS buffer: 10 mM Tris–HCl pH 7.6, 1 mM EDTA pH 8.0, 1% SDS. 8. Affinity purified, specific antibody (e.g., 8WG16 monoclonal antibody [COVANCE], see Fig. 10.3). 9. Protein A sepharose CL-48 (Amersham Biosciences).


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10. Lysis/NaCl Buffer: 50 mM HEPES KOH pH 7.5, 140 mM NaCl, 1 mM EDTA pH 8.0, 1% Triton X-100, 0.1% sodium deoxicholate, 500 mM NaCl. 11. Buffer II: 10 mM Tris–HCL pH 8.0, 1 mM EDTA pH 8.0, 250 mM LiCl, 0.5% NP40, 0.5% sodium deoxicholate. 12. TE buffer: 10 mM Tris–HCl pH 7.6, 1 mM EDTA pH 8.0. 13. Proteinase K stock solution: 20 mg/ml (Roche). 14. GFX purification columns (GE Healthcare). 15. Photolyase (TREVIGEN). 16. BS03 irradiation chamber equipped with UV-A lamps (Philips TLD 15 W/05, emission peak at 360 nm; Dr. Gröbel UVElektronik GmbH). 17. Power SYBR green PCR master mix (Applied Biosystems) 18. Appropriate oligonucleotides (see Note 18). 19. 7500 Fast Real-Time PCR System (Applied Biosystems).

3. Methods 3.1. UV Irradiation and Repair Kinetics of Yeast Cells

1. Grow 400 ml yeast cultures in the appropriate medium and temperature to an absorbance at 600 nm of about 0.8. 2. Harvest cells by centrifugation in a JA-10 rotor (Beckman Coulter) at 6000g and 25C for 5 min. 3. Resuspend cells in minimal medium without amino acids to an absorbance at 600 nm of 1.2 (see Note 2). 4. Do all steps from irradiation to lysis of spheroplasts in dark rooms equipped with safety light to avoid photoreactivation (see Note 3). 5. Pour 200 ml cell suspension into a pyrex tray to produce a 4-mm layer. Take care to keep the remaining cell suspension away from UV light for the non-irradiated control (see Note 4). 6. Irradiate at room temperature with 200 J/m2 of UV light (predominantly 254 nm) generated by germicidal lamps. 7. Immediately after irradiation, pour the cell suspension into an Erlenmeyer flask containing 100 ml of concentrated medium (with 3 amino acids). 8. For the non-irradiated control, pour 40 ml cell suspension into an Erlenmeyer flask containing 20 ml of concentrated medium.

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9. Incubate both the irradiated and non-irradiated cells on a shaker at 30C (with safety light). 10. For repair analysis, remove 50 ml aliquots at 0, 30, 60, 90, and 120 min, chill on ice, and harvest cells by centrifugation at 3000 rpm, 4C for 5 min. The non-irradiated sample is taken at 0 min. 11. For ChIP analysis, remove 40 ml aliquots at 0, 15, 30, 60, and 90 min and process as described below. 3.2. Purification of DNA for Repair Analysis

1. Wash the cells with 50 ml cold H2O. 2. Harvest cells by centrifugation at 3000 rpm, 4C for 5 min. 3. Resuspend cells in 3 ml cold H2O and split each sample into two 2 ml eppendorf tubes. 4. Harvest cells by centrifugation at 3000 rpm, 4C for 5 min. 5. Freeze one of the tubes in liquid nitrogen and store it at –80C. 6. Resuspend the cell pellet in 1 ml spheroplasting buffer by vortexing (see Note 6). 7. Add 150 ml zymolyase stock solution and incubate at 30C for 45 min with nutation. 8. Spin down for 2 min at 6000 rpm in a microfuge and discard the supernatant. Be careful to remove all the supernatant. 9. Wash the pellet once with 500 ml H2O (without resuspending the cells). 10. Resuspend the cell pellet in 200 ml H2O. Add 250 ml solution I, vortex for 10 s, add 25 ml RNase A stock solution, then 25 ml proteinase K stock solution and incubate in a water bath at 50C for 30 min (see Note 7). Invert the tube occasionally. 11. If the DNA happens to precipitate during the treatment, add 100 ml 2.5 M NaCl and incubate at 50C until the DNA is solved to completion. 12. Add 0.5 volume (env. 300 ml) D/I (prewarmed to RT), vortex for 10 s, and spin down for 10 min at full speed in a microfuge. Note that a white layer is formed at the interphase. Transfer the aqueous (upper) phase into a 2 ml tube. 13. Add 2 volumes (env. 1.2 ml) solution II to the aqueous phase, invert gently approximately 20 times until the genomic DNA precipitates (Note that plasmid DNA will precipitate with the genomic DNA). 14. Spin down at full speed in a microfuge at room temperature for 15 min and carefully discard the supernatant. 15. Solve the pellet in 500 ml Na solution (see Note 8).


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16. Add 0.6 volume (300 ml) isopropanol, invert several times until the DNA precipitates and spin down at full speed in a microfuge for 15 min at room temperature (see Note 9). 17. Wash the pellet with 100 ml 70% ethanol (see Note 9). Make sure to remove all of the ethanol. 18. Solve the DNA pellet in 100 ml TE (see Note 10). 19. Check DNA concentration by agarose gel electrophoresis and ethidium bromide staining (loading 5 ml on the gel is fine, the DNA preparation usually yields enough material for five repair analyses). 3.3. Repair Analysis by Indirect EndLabeling

1. Digest DNA to completion with the appropriate restriction enzyme in a total volume of 45 ml. 2. Add 0.5 ml Strataclean resin, vortex, and spin down at full speed in a microfuge for 1 min. 3. Purify over self-made Sephadex G-50 column (in 50T5E buffer; see Note 12). 4. Split the flow-through containing the DNA into two new tubes (20 ml each). 5. Add 3 ml T4-endonulease V (1 U/ml) to one tube (+T4endoV) and 5 ml H2O to the second one (-T4-endoV) and incubate all the tubes at 37C for 1 h. 6. Spin down the tubes and add 2 ml T4-endonulease V (1 U/ml) to the (+T4-endoV) tubes and pursue incubation at 37C for at least 1 h. 7. Add 5.5 ml loading buffer to stop the reaction and denature the DNA (see Note 13). 8. Prepare one horizontal agarose gel (20 25 cm, OWL; 300 ml 1.3% agarose in H2O). Soak the gel in 1 L 1 alkaline electrophoresis buffer for at least 2 h on a shaker at room temperature. After 1 h, exchange the buffer for 1 L fresh alkaline electrophoresis buffer. Precool the gel and fresh electrophoresis buffer for at least 1 h at 4C. 9. Load 25 ml DNA samples per slot and allow the DNA to enter the gel at 300 mA (env. 60 V) for 10–20 min, then continue electrophoresis at 250 mA (env. 50 V) using peristaltic pump in both directions for 15–17 h at 4C, until the dye (bromocresol green) migrates about 14–16 cm (see Note 14). 10. Depending on the size of the restriction fragment of interest, a short treatment (at most 10 min) of the gel in HCl solution might increase the transfer efficiency of large DNA fragments. However, be aware that HCl treatment will decrease the signal coming from the short fragments. Importantly, all

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the gels should be treated exactly the same in order to get comparable results. 11. Rinse the gel shortly with H2O and soak it in denaturating solution for 30 min or more on a shaker. This step might be reduced or even skipped if the gel was not previously treated with HCl solution. 12. Rinse the gel shortly with H2O and soak it in neutralizing solution for 30 min or more on a shaker. Replace buffer for fresh one after the first 15 min of incubation. 13. For Southern transfer, lay two sheets of 3MM Whatman paper presoaked in 20 SSC over a plastic tray bridging a tray with 20 SSC. The paper remains in contact with 20 SSC on both sides. Carefully place the gel on the paper. Cover it with a Hybond N membrane. Cover the membrane with three 3MM Whatman papers presoaked in 20 SSC, 10 cm of filter paper, a glass plate, and a weight of about 250 g. Transfer DNA to the membrane overnight (or 5 h replacing the wet filter papers for fresh ones after 2 h). 11. After transfer, cross-link the DNA to the membrane by UV irradiation (700 J/m2 on each side in a cross-linker). Rinse the membrane with 2 SSC. 12. Place the membrane in rotary cylinders with the DNA bound side facing inside and incubate them in 20 ml hybridization solution at 65C for 2–16 h in an hybridization oven. 13. Prepare strand-specific probes by primer extension. Primer extension reaction contains 2–5 ng appropriate and purified DNA fragment (see Note 11), 10 pmol appropriate primer, 1 Taq polymerase reaction buffer, 1 ml nucleotide stock solution, 1 U Taq polymerase, and 5 ml 32PdCTP in a total volume of 50 ml. Ten to twenty rounds of primer extension (typically 45 s denaturating step at 94C, 5 min annealing step at 56C, and 30 s elongating step at 72C) are used to increase the amount of radioactive DNA. Purify the probe from unbound radioactive nucleotides on a G-50 Sephadex column. 14. Replace the hybridization solution used for the prehybridization with 10 ml of hybridization solution and add the denatured probe. Hybridize at 65C overnight. 15. Pour off the hybridization solution, wash the membrane three times with wash solution I and once with wash solution II for 20 min each at 65C. 16. Air dry the membrane on a sheet of filter paper and expose them to PhosphorImager screens (Fuji). 17. Quantify repair with a PhosphorImager (Fuji FLA5100) and the ImageGauge application software (Fuji). The


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background signal of the gel is determined in a background box normally set next to gel lanes or above gel slots and the average pixel value subtracted to all signal intensities. The CPD content is calculated using the Poisson distribution (Mellon 1987), ln (RFa/RFb), where RFa and RFb represent the intensities of the intact restriction fragment of the T4endoV and mock-treated DNA, respectively (see Note 15). Repair curves are generally calculated as the fraction of CPDs removed versus repair time, the initial damage being set to 0% repair. 3.4. ChIP Analysis

1. Cross-link the cells by adding formaldehyde to a final concentration of 1% (2.16 ml for 40 ml cell suspension) and incubate 15 min on a shaker at room temperature (see Note 19). 2. Add 2.2 ml glycine solution (125 mM final concentration) and incubate further on a shaker at room temperature for 5 min. 3. Pellet the cells in 50 ml conical tubes at 1500g, 4C in a swing rotor for 5 min. 4. Wash the cells twice with 50 ml cold 1 PBS. The cell pellets might be frozen in liquid nitrogen and kept at 80C at this point. 5. Wash the cells once with 500 ml lysis buffer. 6. Resuspend the cells in 400 ml lysis buffer and transfer to eppendorf tubes containing 1 volume of 0.5 mm glass beads (normal tubes might be used, but screw-tops prevent leaking). 7. Lyse the cells by vigorous vortexing for 45 min at 4C. 8. Use needles to make a hole at the bottom of each tube and place them in 2 ml eppendorf tubes of which the cap was previously cut off. Spin down the tubes 5 s in a microfuge to recover the extracts. 9. Transfer the extracts to 1.5 ml eppendorf tubes and place on ice. Sonicate each sample in a bioruptor for 15 min, position high, 30 s ‘ON’ followed by 30 s ‘OFF’ at 4C (see Note 20). 10. Centrifuge at full speed in a table microfuge for 5 min at 4C to clear the extract of debris and unlysed cells. Transfer the supernatant to a new tube. 11. Centrifuge at full speed in a table microfuge for 15 min at 4C to clear any remaining debris and transfer the supernatant to a new tube (see Note 21). 12. Take a 20 ml aliquot (INPUT) and process immediately as described in point 23.

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13. Add the antibodies to the extracts (e.g., 5–10 ml commercial anti-Rpb1 antibody) and incubate overnight at 4C with nutation. 14. Wash the protein A-coated beads (about 15 ml dry volume per reaction) in 1 ml cold 1 PBS for 5 min at 4C with nutation. Spin down in a table microfuge at 4C for 5 s and discard supernatant. Wash twice with lysis buffer using the same procedure to equilibrate the beads. 15. Recover the antibody–protein–DNA complexes with equilibrated protein A beads (see Note 22). Incubate at 4C for 2 h with nutation. 16. Spin down in a table microfuge at 4C for 5 s, discard supernatant, resuspend the beads in 1 ml lysis buffer, and incubate 5 min in agitation at 4C. 16. Spin down in a table microfuge at 4C for 5 s, discard supernatant, resuspend the beads in 1 ml lysis/NaCl buffer, and incubate 5 min in agitation at 4C. 17. Spin down in a table microfuge at 4C for 5 s, discard supernatant, resuspend the beads in 1 ml buffer II, and incubate 5 min in agitation at 4C. 18. Spin down in a table microfuge at 4C for 5 s, discard supernatant, resuspend the beads in 1 ml TE, and incubate 5 min in agitation at 4C. 19. Spin down in a table microfuge for 5 s, discard supernatant, and take all the liquid off using a clean Hamilton syringe. 20. Elute the precipitated material from the beads by adding 60 ml TE 1% SDS to the beads. Incubate 10 min at 65C (see Note 23). 21. Spin down at full speed in a table microfuge for 2 min and transfer the supernatant (env. 60 ml) to a new tube. 22. Take a 20 ml aliquot (PRECIPITATE) and process immediately as described in point 23. Keep the remaining at 4C as backup. 23. Add the 20 ml aliquots to PCR tubes containing 100 ml TE 1% SDS and incubate overnight at 65C to reverse the cross-links (see Note 24). 24. Transfer to a 1.5 ml tube, add 120 ml TE and 5 ml poteinase K stock solution, and incubate at 37C for 2 h. 25. Purify over GFX columns according to manufacturer protocol and elute in 200 ml TE. 26. Photoreactivate all samples with 200 ng photolyase under photoreactivating light for 30 min. We photoreactivate the samples as 50 ml drops in inverted eppendort caps (see Note 25).


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27. Purify over GFX columns and elute in 160 ml TE. 28. Quantify the enriched DNA fragments in the PRECIPITATE by real-time quantitative PCR. We perform real-time quantitative PCR using SYBR green dye and the absolute quantification protocol in the 7500 Real-Time PCR system (Applied Biosystems). Standard curves for all pairs of primers are performed for each analysis. All PCR reactions are performed in triplicate. The enrichment for each PCR of interest is calculated as the ratio between the region-specific signal and the intergenic signal of the PRECIPITATE, normalized with respect to the corresponding ratios of the INPUT. An example is shown in Fig. 10.3.

4. Notes 1. Alternatively, an homemade irradiation chamber containing several germicidal lamps arranged in parallel and an illumination stage at a distance of about 40 cm from the lamps might be used. UVX radiometer (UVP) with a UVX-25 photocell will be required to measure the flux and calculate the irradiation time. 2. The cell density in the liquid culture as well as the UV absorbance of the medium need to be considered for DNA-damage induction. Some people irradiate in medium without sugar (or even in PBS), we prefer to use sugar-containing medium to minimalize the additional stresses suffered by the yeast cells. However, the presence of sugar in the media, which does absorb some of the UV light, makes it necessary to irradiate the cells at relatively high dosage to generate a reasonable amount of damage (about 0.3–2 CPD per kilobase). 3. The use of safety lights (Philips TLD RED) and brown flasks is highly recommended because photoreactivation is a fast process and could be promoted by traces of day light. 4. Rectangular plastic trays might also been used, as long as they do not absorb UV light and produce a 4-mm layer of cell suspension. 5. The RNase A should be carefully boiled to eliminate residual DNAse activity. 6. It is important that the cells are well resuspended. Usually, we deduce the volume of the cell pellet from the water used in the spheroplasting buffer to ensure that the final concentrations are correct.

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7. The tubes might be slowly cooled down to room temperature by leaving them overnight in the turned-off water bath. 8. It is important to solve the DNA pellet to completion. If it does not go into solution, add 50–100 ml 2.5 M NaCl and incubate at 50C for 15 min. 9. Both the ethanol and the isopropanol should be stored at room temperature to avoid coprecipitation of CTAB. 10 The best is to keep the DNA at 4C at least 1 day before going on with the repair analysis to ensure that it is completely solved. 11. To generate strand-specific probes for indirect end-labeling, DNA fragments of 200–300 bp that hybridize close to the end of the restriction fragment of interest should be used. The DNA fragment might be purified from plasmid DNA or amplified by PCR and purified twice over GFX columns to get rid of all the oligonucleotides used as primer (traces of residual primers would impede the strand specificity of the probe generated by primer extension). Since CPD distribution is strand specific, each strand will produce a characteristic CPD pattern (see Fig. 10.1). 12. To prepare Sephadex G-50 columns, cut off the bottom and the cap of an 1.5 ml eppendorf tube, insert a piece (about 4 mm long) of cigarette filter inside the tube, and place it in a 2 ml eppendorf tube. Add 1 ml of Sephadex G-50 slurry (in the appropriate buffer) in the tube and centrifuge at 2000 rpm for 2 min in a microfuge. The Sephadex G-50 containing tube is then placed in a new 2 ml tube, which serves to collect the purified DNA after centrifugation at 2000 rpm for 2 min in a microfuge. 13. The experiment should be planned such that DNA is loaded directly after having added the denaturating loading buffer. 14. Since the large amount of dye generates background signal, it is recommended to use less dye in the samples and to load a larger amount of loading buffer in an empty lane (if available) to follow the migration more easily. 15. Using this technique, region-specific damage can also be quantified (relative to total lesions). In this case, regionspecific damage is calculated as the signal of that region in the T4endoV-treated DNA divided by the signal of the whole lane. The corresponding signal of the mock-treated DNA is subtracted as background. 16. Lysis buffer lacking protease inhibitors might be stored at 4C. Once the protease inhibitors are added, the buffer should be kept on ice (or at 4C) and used within 2 days. Protease stock solutions (100 ) are prepared by solving benzamidine in


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H2O, PMSF in isopropanol, and TLCK in 50 mM sodium acetate pH 5.0. Protease stock solutions (1000 ) are prepared by solving aprotinin in HEPES/KOH pH 8.0, leupeptin and antipain in H2O, and Papstatin A in ethanol. All stock solutions are stored at 20C, except for aprotinin, which is stored at 4C. All proteases are from SIGMA. 17. Before use, the beads are acid washed by soaking overnight in nitric acid, extensive rinsing with distilled water and baking at 200C until dry (might take up to 1 week depending on the amount). Some people add an additional final wash with 70% ethanol and bake the beads at 80C. 18. We use the Primer Express application software (Applied Biosystems) to design the oligonucleotides. Beside the primer pairs required to amplify the regions of interest, a primer pair amplifying a region devoided of the protein against which the immunoprecipitation is set (e.g., intergenic region) is required. 19. Others incubate for 30 min at 30C and claim that for certain IPs, overnight incubation at 4C is helpful. 20. The extracts might be briefly centrifuged and the pellets resuspended in lysis buffer prior to sonication. The sonification conditions should be set previously for each sonicator to generate DNA fragments between 100 and 500 pb, with a peak at 300 bp. 21. At this point 5 ml of the extracts might be run on an agarose gel to check fragment lengths. After this step, salt or detergent concentrations may be increased or decreased for more or less stringent IPs, respectively. 22. We use about 50 ml swollen (wet) beads per reaction. For pipetting, cut off the end of a pipette tip to make the opening wider. During washes, do not spin the beads at full speed because they can get crushed. 23. To improve the efficiency of recovery, incubation might be done for up to 30 min and repeated a second time, pooling the eluates afterwards. 24. It is recommended to use a thermocycler to prevent evaporation. 25. Alternatively, the samples might be photoreactivated in an homemade irradiation chamber containing several UV-A lamps arranged in parallel and an illumination stage at a distance of about 40 cm from the lamps. Since photolyase does not require a particular reaction buffer, 200 ng photolyase (freshly diluted to 100 ng/ml in TE) are directly added to each sample, which is then split in four inverted eppendorf caps. To manipulate and transport the inverted eppendorf caps containing the samples, it is useful to place them in

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plastic petri dishes, which can also be easily labeled. After photoreactivation, the four drops of each samples are pooled.

Acknowledgments Research in AA’s laboratory is funded by grants from the Spanish Ministery of Science and Innovation and from the Junta de Andaluca.

References 1. Hanawalt, P. C. (2001). Controlling the efficiency of excision repair. Mutat. Res. 485, 3–13. 2. Prakash, S., and Prakash, L. (2000). Nucleotide excision repair in yeast. Mutat. Res. 451, 13–24. 3. de Boer, J., and Hoeijmakers, J. H. (2000). Nucleotide excision repair and human syndromes. Carcinogenesis 21, 453–60. 4. Verhage, R., Zeeman, A. M., de Groot, N., Gleig, F., Bang, D. D., van de Putte, P., and Brouwer, J. (1994). The RAD7 and RAD16 genes, which are essential for pyrimidine dimer removal from the silent mating type loci, are also required for repair of the nontranscribed strand of an active gene in Saccharomyces cerevisiae. Mol. Cell. Biol. 14, 6135–42. 5. Mellon, I. (2005). Transcription-coupled repair: a complex affair. Mutat. Res. 577, 155–61. 6. Gaillard, H., Wellinger, R. E., and Aguilera, A. (2007). A new connection of mRNP biogenesis and export with transcription-coupled repair. Nucleic Acids Res. 35, 3893–906. 7. Laine, J. P., and Egly, J. M. (2006). When transcription and repair meet: a complex system. Trends Genet. 22, 430–6. 8. Tornaletti, S. (2005). Transcription arrest at DNA damage sites. Mutat. Res. 577, 131–45. 9. Svejstrup, J. Q. (2002). Mechanisms of transcription-coupled DNA repair. Nat. Rev. Mol. Cell Biol. 3, 21–9. 10. Venema, J., Mullenders, L. H., Natarajan, A. T., van Zeeland, A. A., and Mayne, L. V. (1990). The genetic defect in Cockayne syndrome is associated with a defect in repair of UV-induced

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DNA damage in transcriptionally. Proc. Natl. Acad. Sci. U S A 87, 4707–11. van Gool, A. J., Verhage, R., Swagemakers, S. M., van de Putte, P., Brouwer, J., Troelstra, C., Bootsma, D., and Hoeijmakers, J. H. (1994). RAD26, the functional S. cerevisiae homolog of the Cockayne syndrome B gene ERCC6. EMBO J. 13, 5361–9. Li, S., Chen, X., Ruggiero, C., Ding, B., and Smerdon, M. J. (2006). Modulation of Rad26- and Rpb9-mediated DNA repair by different promoter elements. J. Biol. Chem. 281, 36643–51. Li, S., Ding, B., Chen, R., Ruggiero, C., and Chen, X. (2006). Evidence that the transcription elongation function of Rpb9 is involved in transcription-coupled DNA repair in Saccharomyces cerevisiae. Mol. Cell. Biol. 26, 9430–41. Li, S., and Smerdon, M. J. (2002). Rpb4 and Rpb9 mediate subpathways of transcriptioncoupled DNA repair in Saccharomyces cerevisiae. EMBO J. 21, 5921–9. Aguilera, A. (2005). Cotranscriptional mRNP assembly: from the DNA to the nuclear pore. Curr. Opin. Cell Biol. 17, 242–50. Luo, Z., Zheng, J., Lu, Y., and Bregman, D. B. (2001). Ultraviolet radiation alters the phosphorylation of RNA polymerase II large subunit and accelerates its proteasome-dependent degradation. Mutat. Res. 486, 259–74. Beaudenon, S. L., Huacani, M. R., Wang, G., McDonnell, D. P., and Huibregtse, J. M. (1999). Rsp5 ubiquitin-protein ligase mediates DNA damage-induced degradation of the large subunit of RNA polymerase II in Saccharomyces cerevisiae. Mol. Cell. Biol. 19, 6972–9.


18. Ratner, J. N., Balasubramanian, B., Corden, J., Warren, S. L., and Bregman, D. B. (1998). Ultraviolet radiation-induced ubiquitination and proteasomal degradation of the large subunit of RNA polymerase II. Implications for transcription-coupled DNA repair. J. Biol. Chem. 273, 5184–9. 19. Bregman, D. B., Halaban, R., van Gool, A. J., Henning, K. A., Friedberg, E. C., and Warren, S. L. (1996). UV-induced ubiquitination of RNA polymerase II: a novel modification deficient in Cockayne syndrome cells. Proc. Natl. Acad. Sci. U S A 93, 11586–90. 20. Svejstrup, J. Q. (2003). Rescue of arrested RNA polymerase II complexes. J. Cell Sci. 116, 447–51. 21. Reid, J., and Svejstrup, J. Q. (2004). DNA damage-induced Def1-RNA polymerase II interaction and Def1 requirement for polymerase ubiquitylation in vitro. J. Biol. Chem. 279, 29875–8. 22. Somesh, B. P., Reid, J., Liu, W. F., Sogaard, T. M., Erdjument-Bromage, H., Tempst, P., and Svejstrup, J. Q. (2005). Multiple mechanisms confining RNA polymerase II ubiquitylation to polymerases undergoing transcriptional arrest.. Cell 121, 913–23. 23. Li, S., Waters, R., and Smerdon, M. J. (2000). Low- and high-resolution mapping of DNA damage at specific sites. Methods 22, 170–9. 24. Lloyd, R. S. (1999). The initiation of DNA base excision repair of dipyrimidine

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Chapter 11 Cytometric Analysis of DNA Damage: Phosphorylation of Histone H2AX as a Marker of DNA Double-Strand Breaks (DSBs) Toshiki Tanaka, Dorota Halicka, Frank Traganos, and Zbigniew Darzynkiewicz Abstract Phosphorylation of histone H2AX on Ser 139 is a sensitive reporter of DNA damage, particularly if the damage involves induction of DNA double-strand breaks (DSBs). Phosphorylated H2AX has been named H2AX and its presence in the nucleus can be detected immunocytochemically. Multiparameter analysis of H2AX immunofluorescence by flow or laser-scanning cytometry allows one to measure extent of DNA damage in individual cells and to correlate it with their position in the cell cycle and induction of apoptosis. This chapter presents the protocols and outlines applications of multiparameter cytometry in analysis of H2AX phosphorylation as a reporter of the presence of DSBs. Key words: H2AX, H2AX phosphorylation, DNA double-strand breaks, Multiparameter flow cytometry, Laser-scanning cytometry, Immunocytochemistry, Apoptosis.

1. Introduction DNA damage that involves formation of DSBs induces phosphorylation of Ser-139 at the carboxy terminus of histone H2AX (1, 2) one of several variants of the nucleosome core histone H2A (3–5). The phosphorylation takes place on H2AX molecules in megabase chromatin domains flanking the DSBs and is mediated by the PI-3-like protein kinases, ATM- (1, 2, 6, 7) ATR- (8), and/or DNA-dependent protein kinase (DNAPK) (9, 10). The Ser-139 phosphorylated H2AX has been named H2AX (11). Development of an Ab specific to H2AX made it possible to detect H2AX phosphorylation and thus to assay immunocytochemically DNA damage and repair in Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_11 Springerprotocols.com

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situ, in chromatin of individual cells (12). Compared with the alternative method of DNA damage assessment which is based on analysis of electrophoretic mobility of DNA released from individual cells (comet assay) the immunocytochemical approach is less cumbersome and offers much greater sensitivity (13). Shortly after induction of DSBs, e.g., by ionizing radiation or other genotoxic agents, the presence of H2AX in chromatin can be detected with this Ab in the form of discrete nuclear foci (2, 11). Because each focus represents a single DSB2 their frequency is considered to report the incidence of DSBs. Several checkpoint and DNA repair proteins such as Rad50, Rad51, and Brca1 co-localize with H2AX (14). It was recently proposed that phosphorylated H2AX may function as an anchor holding broken DNA ends in close proximity in chromatin, facilitating their repair (15). H2AX also mediates translocation of the p53-binding protein 1 (53BP1) to the radiation-induced foci (7). Western blotting and detection of H2AX immunofluorescene (IF) provide two major approaches to analyze H2AX phosphorylation. Measurement of H2AX IF by multiparameter flow or laser-scanning cytometry (16–20) is particularly advantageous. The major benefit of the cytometric approach stems from the fact that H2AX phosphorylation in situ, in chromatin of individual cells, can be measured with high sensitivity and accuracy and the expression of H2AX can be directly correlated, within the same cells, with their DNA content, induction of apoptosis or any other cell attribute of interest. Large cell numbers, thus, may be rapidly analyzed and the data provide information on the extent of H2AX phosphorylation with respect to their cell cycle phase, commitment to die in response to DNA damage, their surface immunophenotype, etc. Cytometry also allows one to analyze intercellular variability in H2AX phosphorylation within cell populations and to identify rare cell subpopulations, otherwise undetectable by Western blotting. The chapter which follows is focused on applications of multiparameter cytometry in analysis of H2AX phosphorylation as a reporter of the presence of DSBs.

2. Materials 1. Primary antibody: anti-phospho-histone H2A.X (Ser-139) mAb, clone JBW301 (Upstate) 2. Secondary antibodies: Alexa Fluor 488 F(ab0 )2 fragment of goat anti-mouse IgG (H+L) (Invitrogen) 3. Buffer: 1 phosphate-buffered saline (PBS) (Sigma)

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4. Solutions (stored at 4C) a. 1% methanol-free formaldehyde (Polysciences, Inc.) dissolved in PBS b. 1% (w/v) solution of bovine serum albumin (BSA; Sigma) dissolved in PBS b. 1 mg/ml 4,6-diamidino-2-phenylindole (DAPI, Sigma) dissolved in PBS d. 0.1% Triton X-100 (Sigma) dissolved in PBS (see Note 2) e. 5 mg/ml PI in the presence of 100 mg/ml of RNase A (Sigma) dissolved in PBS f. 80% ethanol in dH2O

3. Methods 3.1. Staining of Cells to be Measured by Flow Cytometry

3.1.1. Fixation of Cells

1. Suspend 106 cells in 0.5 ml of PBS. 2. Transfer this cell suspension to 5 ml of 1% formaldehyde in tube for 15 min at 1–4C (on ice), then centrifuge (200 g, 5 min). 3. Suspend the pellet in 80% ethanol for at least 2 h; the cells can be stored in 80% ethanol for up to several days at –20C.

3.1.2. Staining

1. Centrifuge (200 g, 5 min) cells suspended in 80% ethanol then rinse (200 g, 5 min them twice) with PBS. 2. Add 2–3 ml 1% BSA solution into the tubes and centrifuge (200 g, 5 min). 3. Discard the supernatant and transfer the cells suspended in 100 ml of 1% BSA to new tubes. 4. Add 1 ml (1:100) H2AX primary antibody into the suspension, cover the tubes with parafilm to prevent drying and incubate for 2 h at room temperature (see Note 1). 5. Add 2–3 ml 1% BSA into the tubes and centrifuge (200 g, 5 min). 6. Discard the supernatant, suspend cells to 100 ml 1% BSA solution and transfer to new tubes. 7. Add 1 ml (1:100) Alexa Fluor 488 anti-mouse antibody, cover the tubes to prevent drying and incubate for 30 min at room temperature in the dark.

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8. Add 2–3 ml 1% BSA into the tubes and centrifuge them for 5 min. 9. Add 0.5–1.0 ml of 10 mg/ml propidium iodide (PI) – RNase solution into the tubes and transfer to tubes used for FACScan. Incubate cells for 30 min at room temperature in the dark. 10. Measure the samples by FACScan. 3.2. Staining of Cells to be Measured by Laser Scanning Cytometry (LSC) 3.2.1. Fixation (For Cells Growing on Chamber Slides)

The cells may be deposited on slides either by growing them in ‘‘Chamber’’-slide vessels (the cells that grow attached) or by attaching them to slides by cytocentrifugation (the cells that grow in suspension or cells detached by trypsinization). 1. Discard the medium and wash each chamber by PBS once. 2. Add ice-cold 1% formaldehyde into each chamber and fix for 15 min on ice. 3. Remove formaldehyde, wash each chamber by PBS once. (see Note 3) 4. Remove the chamber sides and immerse a slide into cold (–20C) 80% ethanol. The slides can be stored in 80% ethanol for up to several days at –20C.

3.2.2. Fixation for Cells Deposited on Slides by Cytocentrifugation

3.2.3. Staining

1. Place the cells-bearing slides in Coplin jars containing 1% formaldehyde at 0–4C for 15 min 2. Rinse the slides with PBS and transfer them to Coplin jars containing 80% ethanol at –20C and store them under these conditions 1. Wash the slide twice with PBS, for 5 min each time. 2. Incubate with 0.1% Triton X-100 for 15 min at room temperature. 3. Incubate with 1% BSA for 30 min at room temperature. 4. Incubate with 50–100 ml 1% BSA containing 1:200 H2AX antibody in each chamber area (100–200 ml to each slide) for overnight at 4C. Rigorously prevent drying by covering the site containing cells and antibody solution with a coverslip made of parafilm or polyethylene foil, and keeping the slides in the moist chamber. 5. Wash slide by PBS for 5 min once. 6. Incubate with 50–100 ml 1% BSA containing 1:100 Alexa Flour 488 antibody in each chamber area (100–200 ml to each slide) for 40 min at room temperature preventing drying. 7. Wash a slide by PBS for 5 min once. 8. Incubate with 100 ml 1 mg/ml DAPI in each chamber area for 10 min at room temperature.


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9. Wash a slide by PBS and put one drop of mounting (antifade) medium on each chamber or cytospin area, then cover it with the cover glass. 10. Measure cell fluorescence by LSC. 3.3. Instrument Settings

1. Flow cytometer The fluorescence of red (PI) and green (Alexa Fluor 488) of individual cells induced by excitation with a 488 nm argon ion laser is measured using a FACScan cytometer (Becton Dickinson) with the standard optics and CELLQuest software (Becton Dickinson). At least 10,000 cells should be measured per sample. All experiments should be repeated at least three times. 2. LSC Cellular blue (DAPI) and green (Alexa Fluor 488) fluorescence emission is measured simultaneously in the same cells using iCys LSC (CompuCyte) utilizing standard filter settings; blue and green fluorescence is excited with violet diode and 488-nm argon ion lasers, respectively. The intensities of integrated fluorescence and maximal pixel are measured and recorded for each cell. At least 3000 cells were measured per sample. Each experiment should be run at least in triplicate.

3.4. Results and Data Interpretation

The protocols should be able to measure the induction of H2AX phosphorylation in HL-60 cells treated with topotecan (Tpt) measured by FACScan as well as induction of H2AX phosphorylation in A549 cells treated with Tpt measured by LSC. As mentioned earlier, H2AX phosphorylation (H2AX expression) measured by cytometry is often a reporter of DNA damage, particularly when the damage involves formation of DSBs. This is the case, for example, when the damage of DNA was induced by DNA topoisomerase I inhibitor Tpt (as shown in Figs. 11.1 and 11.2), DSBs are caused by ionizing radiation (1, 2) or generated

Fig. 11.1. This figure demonstrates selective response of S-phase to Tpt-induced DNA damage. The data was obtained using FACScan flow cytometer (BD Biologicals, San Jose, CA). FL-1 shows the level of H2AX immunofluorescence (green fluorescence) and FL-3 shows DNA content (red fluorescence of DNA-bound PI).

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Fig. 11.2. This figure also demonstrates selective response of S-phase to Tpt-induced DNA damage. The data was obtained using iCys laser scanning cytometer (CompuCyte, Westwood, MA). X axis shows the level of H2AX immunofluorescence and Y axis shows DNA content.

during apoptotic DNA fragmentation (13). It should be stressed, however, that H2AX may be phosphorylated on Ser-139 also in the absence of induction of DSBs. This can be seen during replication stress induced by inhibition of DNA synthesis (21) or during chromatin condensation such as during mitosis in some cell types as well as upon induction of premature chromosome condensation (PCC) (22). A caution, therefore, has to be exercised in interpreting expression of H2AX as a marker of DSBs. (see Notes 4 and 5)

4. Notes 1. The concentration of H2AX Ab can be between 1:100 and 1:400. Titrate the Ab initially and use it subsequently at the concentration that gives maximal signal-to-noise ratio, i.e., the maximal IF difference between the untreated cells and the cells subjected to treatment that generated DNA damage 2. The presence of nonionic detergent such as Triton X-100 is helpful since it increases permeability of the plasma membrane to primary and secondary Abs 3. Be careful while washing the cells growing on chamber slides because the cells in mitotic phase or undergoing apoptosis may be detached from the slide during vigorous washing. 4. The gating analysis can be carried out to obtain mean values of H2AX IF for cells in G1, S and G2M phases of the cell cycle by selecting the cells with DNA index (DI) 1.0–0.2 (G1); 1.2 > (S) < 1.8 and 2.0–0.2 (G2), respectively.


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5. The data can also be expressed as a change in mean H2AX IF (; delta) either of all cells or after gating analysis cells in G1, S or G2M phase of the cycle, due to the treatment, by subtracting the mean IF value of the untreated cells from the mean IF value of the treated ones in respective phases of the cell cycle.

References 1. Rogakou EP, Pilch DR, Orr AH, Ivanova VS, Bonner WM. (1998) DNA double-stranded breaks induce histone H2AX phosphorylation on serine 139. J Biol Chem. 273, 5858–5868. 2. Sedelnikova OA, Rogakou EP, Panuytin IG, Bonner W. (2002) Quantitative detection of 125IUdr-induced DNA double-strand breaks with -H2AX antibody. Radiation Res. 158, 486–492. 3. West MH, Bonner WM. (1980) Histone 2A, a heteromorphous family of eight protein species. Biochemistry. 19, 3238–3245. 4. Thatcher TH, Gorovsky MA. (1994) Phylogenetic analysis of the core histones H2A, H2B, H3 and H4. Nucleic Acids Res. 22, 174–179. 5. Pehrson JR, Fuji RN. (1998) Evolutionary conservation of histone macro-H2A subtypes and domains. Nuclei Acids Res. 26, 2837–2842. 6. Burma S, Chen BP, Murphy M, Kurimasa A, Chen DJ. (2001) ATM phosphorylates histone H2AX in response to DNA double-strand breaks. J Biol Chem. 276, 42462–42467. 7. Anderson L, Henderson C, Adachi Y. (2001) Phosphorylation and rapid relocalization of 53BP1 to nuclear foci upon DNA damage. Mol Cell Biol. 21, 1719–1729. 8. Furuta T, Takemura H, Liao-Z-Y, Aune GJ, Redon C, Sedelnikova OA, Pilch DR, Rogakou EP, Celeste A, Chen HT, Nussenzweig A, Aladjem MI, Bonner WM, Pommier Y. (2003) Phosphorylation of histone H2AX and activation of Mre11, Rad50, and Nbs1 in response to replication-dependent DNA double-strand breaks induces by mammalian topoisomerase I cleavage complexes. J Biol Chem. 278, 20303–20312. 9. Park EJ, Chan DW, Park JH, Oettinger MA, Kwon J. (2003) DNA-PK is activated by nucleosomes and phosphorylated H2AX within the nucleosomes in an acetylationdependent manner. Nuclei Acids Res. 31, 6819–6827.

10. Wang H, Wang M, Wang H, Bocker W, Iliakis G. (2005) Complex H2AX phosphorylation patterns by multiple kinases including ATM and DNA-PK in human cells exposed to ionizing radiation and treated with kinase inhibitors. J Cell Physiol. 202, 492–502. 11. Rogakou EP, Boon C, Redon, C, Bonner WM. (1999) Megabase chromatin domains involved in DNA double-strand breaks in vivo. J Cell Biol. 146, 905–916. 12. Banath JP, Olive PL. (2003) Expression of phosphorylated histone H2AX as a surrogate of cell killing by drugs that create DNA double-strand breaks. Cancer Res. 63, 4347–4350. 13. Huang X, Okafuji M, Traganos F, Luther E, Holden E, Darzynkiewicz Z. (2004) Assessment of histone H2AX phosphorylation induced by DNA topoisomerase I and II inhibitors Topotecan and Mitoxantrone and by the DNA cross-linking agent Cisplatin. Cytometry. 58A, 99–110. 14. Paull TT, Rogakou EP, Yamazaki V, Kirchgesser CU, Gellert M, Bonner WM. (2000) A critical role for histone H2AX in recruitment of repair factors to nuclear foci after DNA damage. Curr Biol. 10, 886–895. 15. Bassing CH, Alt FW. (2004) H2AX may function as an anchor to hold broken chromosomal DNA ends in close proximity. Cell Cycle. 3, 149–153. 16. Tanaka T, Kurose A, Halicka HD, Traganos F, Darzynkiewicz Z. (2006) 2-Deoxy-D-glucose reduces the level of constitutive activation of ATM and phosphorylation of histone H2AX. Cell Cycle. 5, 878–882. 17. Tanaka T, Kurose A, Halicka HD, Huang X, Traganos F, Darzynkiewicz Z. (2006) Nitrogen oxide-releasing aspirin induces histone H2AX phosphorylation, ATM activation and apoptosis preferentially in S-phase cells. Cell Cycle. 5, 1669–1674.

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18. Tanaka T, Halicka HD, Huang X, Traganos F, Darzynkiewicz Z. (2006) Constitutive histone H2AX phosphorylation and ATM activation, the reporters of DNA damage by endogenous oxidants. Cell Cycle. 5, 1940–1945. 19. Tanaka T, Huang X, Jorgensen E, Gietl E, Traganos F, Darzynkiewicz Z, Albino AP. (2007) ATM activation accompanies histone H2AX phosphorylation in A549 cells upon exposure to tobacco smoke. BMC Cell Biol. 8, 26. 20. Tanaka T, Huang X, Halicka HD, Zhao H, Traganos F, Albino AP, Dai W, Darzynkiewicz Z. (2007) Cytometry of ATM activation and histone H2AX phosphorylation to estimate extent of DNA damage induced

by exogenous agents. Cytometry 71A, 648–661. 21. Kurose A, Tanaka T, Huang X, Traganos F, Dai W, Darzynkiewicz Z (2006). Effects of hydroxyurea and aphidicolin on phosphorylation of ATM on Ser 1981 and histone H2AX on Ser 139 in relation to cell cycle phase and induction of apoptosis. Cytometry A 69A, 212–221. 22. Huang X, Kurose A, Tanaka T, Traganos F, Dai W, Darzynkiewicz Z (2006) Sequential phosphorylation of Ser-10 on histone H3 and Ser-139 on histone H2AX and ATM activation during premature chromosome condensation: Relationship to cell-cycle and apoptosis. Cytometry A 69A, 222–229.

Chapter 12 Detection of DNA Damage Induced by Topoisomerase II Inhibitors, Gamma Radiation and Crosslinking Agents Using the Comet Assay Lori A. Hazlehurst Abstract The comet assay is a simple gel electrophoresis method for visualizing and quantifying DNA damage. The comet assay is sensitive and reproducible and can be used to detect single-strand DNA breaks, doublestrand DNA breaks, protein-associated DNA strand breaks and DNA crosslinks. The comet assay uses fluorescent DNA-binding dyes to detect both damaged DNA that resides in the tail region and undamaged DNA that is retained in the head region following gel electrophoresis. This assay is a single cellbased assay and thus is highly adaptable for measuring DNA damage in clinical samples. Furthermore, unlike other assays the detection of DNA damage is not dependent on the random incorporation of radiolabeled nucleotides. Again this can be problematic with clinical samples as proliferation rates are often slow and culturing of primary patient specimens for 48 h required to randomly label DNA is often not possible. In this chapter we will outline the comet assay for the detection of DNA damage induced by topoisomerase II inhibitors, cross-linking agents and gamma radiation. Key words: Comet assay, filter elution assay, topoisomerase II poison, DNA crosslinks, DNA repair.

1. Introduction Topoisomerase II poisons, DNA cross-linking agents and radiation remain the mainstay of chemotherapy. These chemotherapeutic agents will typically induce apoptosis or programmed cell death. However, the initial cytotoxic lesion occurs at the level of DNA. Activation of pathways that increase the rate of DNA repair, for DNA cross-linking agents and radiation, can decrease the amount of detectable DNA damage. Likewise, decreases in the levels and/or activity of topoisomerase II can diminish the Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_12 Springerprotocols.com

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amount of protein-associated DNA damage induced by topoisomerase II poisons. These differences in amount of initial DNA damage for specific types of agents can be detected by the comet assay. By adjusting buffers one can easily detect single or double DNA strand breaks, and the addition of proteinase K allows for the detection of protein-associated DNA strand breaks. Previous methods to detect DNA damage such as alkaline and neutral elution assays give similar results as the comet assay. The alkaline and neutral elution assays are based on time of elution of damaged DNA through a gravity drip filter, whereas the comet assay is based on migration of damaged DNA in agarose using gel electrophoreses principles (1, 2). Elution assays generate large volumes of radioactive waste and specialized equipment is needed which includes a fraction collector and peristaltic pump. Additional experimental advantages of the comet assay compared to the alkaline or neutral elution assays are (1) DNA damage can be measured in cycling and non-cycling cells. The alkaline and neutral elution assay requires 3H and 14C thymidine incorporation into DNA for 24–48 h prior to performing the assay. This is to ensure random labeling of DNA. Thus elution assays are not suitable for quiescent or slowly dividing cells and (2) the comet assay can be performed on a minimal sample size (less then 5,000 cells). Thus, in summary the comet assay is adaptable to measuring DNA damage in patient specimens were sample size is often limited and ex-vivo growth can be difficult.

2. Materials 2.1. Buffers for Detection of DoubleStrands Breaks

1. Lysis buffer for the detection of double-strand DNA breaks: 0.5% SDS and 30 mM disodium ethylendiamine tetraacetic acid (EDTA) pH 8.0. This buffer maybe stored at room temperature. If detecting protein-associated strand breaks produced by treatment with a topoisomerase II inhibitor 25 mg proteinase K per 100 ml of lysis buffer should be added the day of the experiment. 2. Electrophoresis buffer for double-strand DNA breaks: TBE (90 mM Tris, 90 mM boric acid, 2 mM EDTA pH 8.0. 3. DNA-staining solution Syber green (Invitrogen, Carlsbad, CA) 1:10,000 dilution in TBE.

2.2. Buffers for Detection of SingleStrand Breaks

1. Lysis buffer for the detection of single-strand breaks: 2.5 M Na2Cl, 100 mM EDTA, 1% sodium sarcosinate, 1% Triton X-100, 10% DMSO and 10 mM Tris pH 10.00. 2. Alkaline unwinding buffer and electrophoresis buffer for singlestrand breaks: 0.03 M NaOH and 1 mM EDTA pH 12.2. The

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pH must be above 12.0 to ensure adequate denaturating of the DNA. Furthermore, once slides have been exposed to alkaline buffer they should not be placed in a neutral buffer until after electrophoresis has occurred. 2.3. Electrophoresis

1. 1% low gelling temperature agarose (Nusieve GTG) made in PBS. Once the agarose solution is dissolved place in 45C water bath. The agarose should be made fresh daily. 2. Fully frosted microscope slides aid in retaining the agarose attached to the slide. Best to solidify the agar cell suspension on the slides at 4C for 3 min before placing in lysis buffer (see Note 1). 3. A horizontal electrophoresis chamber is needed. A large flat bed is recommended to accommodate at least 10–12 individual slides. A power supply that can generate a constant voltage of 20 V is required.

2.4. Image Capturing and Analysis

1. Epifluorescence microscope with good-quality objectives suitable for fluorescent imaging. The microscope needs to be equipped with a black and white camera capable of collecting images in ‘‘real time.’’ 2. Florescent filters to accommodate an excitation peak of 495 nm with a peak emission of 520 nm characteristic of Syber green. 3. Several commercial imaging packages are available for image analysis.

3. Methods 3.1. Detection of DNA Damage Induced by Topoisomerase II Poisons

Topoisomerase II is an ATP-dependent enzyme, which is capable of cleaving double-stranded DNA and strand passage thereby relieving torsional strain. During the enzymatic function, topoisomerase II will form a transient covalent bond with the 50 end of the break site. A topoisomerase II poison will stabilize the normally transient covalent bond resulting in protein–DNA complex referred to as a cleavable complex (3). This cleavable complex can be measured by a variety of methods including SDS–KCl precipitation of DNA–protein complexes, alkaline and neutral elution as well as by the comet assay (4–7). If DNA damage is mediated by topoisomerase II, DNA double-strand breaks can be detected when proteinase K is added to the lysis buffer. The addition of proteinase K will result in the digestion of the cleavable complex and thereby expose the double-strand DNA break. DNA damage

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can typically be observed following a 1-h treatment with a topoisomerase II poison (see Note 3). Depending on the agent DNA damage can be persistent making repair difficult to measure. However some topoisomerase II poisons such as amsacrine, repair of damaged DNA can be detected within 2 h following removal of the drug from the media (4, 8). 1. Treat cells for 1 h with a topoisomerase II poison. Place 5,000 cells in a 2.0 ml Eppendorf tube and add 1 ml cold PBS. Spin cells down at 4C for 5 min at 1200 g. Aspirate and add 500 ml cold PBS and 1.5 ml of 1% agar (agar is kept at 45) and mix well (see Note 4). Quickly and gently overlay cell suspension on frosted microscope slide and allow to solidify for 3 min at 4C. Gently place slides in cold neutral lysis buffer containing 25 mg/ ml proteinase K at 4C for 1 h. Transfer slides to 37C and incubate for an additional 12–16 h. After 12–16 h equilibrate agarose slides with TBE for 1 h changing out solutions every 15 min. Gently place slides into electrophoresis box containing TBE and slides are subjected to electrophore at 25 V for 20 min. 2. After electrophoresis, DNA is stained with Syber green for 25 min in TBE. 3. To reduce background staining wash slides twice with TBE for 3 min. Slides can be stored in a humidified box in a refrigerator. This same procedure can be used to detect DNA double-strand breaks by gamma radiation (proteinase K is not required in the lysis buffer). 3.2. Detection of DNA Crosslinks

Depending on the agent used crosslinks can be detected within 2 h of drug treatment, with maximal levels of crosslinking occurring between 6 and 12 h (9–11). Following drug treatment, cells are irradiated with a known amount of gamma radiation (500–1000 rads) to introduce random single-strand breaks. Effectively the more crosslinks introduced, the fewer single-strand breaks will be detected in irradiated samples. Percent crosslinks can be calculated using either comet moment or tail moment as follows: % crosslinks ¼ [1–(comet momentdrug treated–comet moment control)/(comet momentRads treated only –comet moment control)]*100 1. Treat cells for 2 h with a cross-linking agent such as melphalan. Repair of DNA crosslinks can be easily measured by removing drug after 2 h and measuring crosslinks across time. To introduce single-strand breaks immediately following drug treatment samples are irradiated at 900 rads. Irradiation should be performed on ice and samples should be kept cold as repair of radiation-induced DNA damage occurs quickly at room


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temperature. Additionally, samples should be placed in lysis buffer as quickly as possible. Place 5,000 cells in a 2.0 ml eppendorf and add 1 ml cold PBS. Spin cells down at 4C for 5 min at 1200 g. Aspirate and add 500 ml of cold PBS and 1.5 ml of 1% agar (agar is kept at 45) and mix well. Quickly and gently overlay cell suspension on frosted microscope slide and allow agarose to solidify for 3 min at 4C. Gently place slides in cold lysis buffer (for single strand breaks) at 4C for 1 h. Transfer slides to alkaline unwinding assay for 1 h. Gently place slides into electrophoresis box containing the alkaline buffer and electrophoresis for 20 min at 25 V (see Note 2). 2. After electrophoresis wash slides several times in TBE to equilibrate the slides and then stain DNA with Syber green for 25 min in TBE. It is essential that the agar is no longer alkaline as the florescent properties of Syber green are negatively affected in alkaline conditions. 3. To reduce background staining wash slides twice with TBE for 3 min. Slides can be stored in a humidified box in refrigerator. 3.3. Detection of Radiation Induced DNA-Strand Breaks

Radiation-induced strand breaks can be measured as single or double-strand breaks. Repair of radiation-induced DNA occurs quickly and thus time points of repair should be performed in minutes and not hours. Detection of single-strand breaks can occur as low as 200 rads, while double-strand breaks detection is in the range of 2,000 rads. It is important that samples are irradiated on ice, and samples should be placed in lysis buffer as quickly as possible to reduce concerns of repair. For detection of doublestrand breaks neutral lysis buffer (proteinase K is not required) is used and TBE is used as the electrophoresis buffer as described above (see Note 3). For detection of single-stand breaks an alkaline unwinding buffer is used to denature double-stranded DNA and for electrophoresis as described above under detection of crosslinks.

3.3.1. Quantification of Comets

Several different methods are available for quantifying comets. These include measuring the tail length, tail moment and the comet moment. The tail length is the most problematic method for quantification, as it does not account for the amount of DNA that has migrated. The end result is the length of the tail will eventually plateau despite increasing DNA damage (Fig. 12.1). Both the comet moment and the tail moment take into account the amount and distance the DNA has migrated and thus is a more useful measurement for quantifying DNA damage. Olive originally defined tail moment as the percentage of DNA in the tail multiplied by the distance between the means of the head and tail distributions (12). The comet moment was described later by Kent et al. (13). The comet moment has the advantage of only requiring defining the center of the comet head and is described below. When the comet moment is plotted against comet tail

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Fig. 12.1. A typical comet assay to measure DNA damage. The relation between comet moment, intensity and distance traveled is shown.

there is a close correlation. However, for low levels of DNA damage where a distinct tail region is difficult to ascertain the comet moment may be advantageous compared to the tail moment. 3.3.2. Statistical Considerations

1. A minimum of 50 images should be analyzed per sample (See Notes 5–9). Three independent experiments should be performed and these data can be used to determine the mean and standard deviation. 2. If performing a dose–response ANOVA is a good statistical method for testing for significant differences between treatment groups.

4. Notes 1. A major initial complaint of this assay is the detachment of agar from the slides. We have found that solidifying in the cold prior to adding to lysis buffer helps reduce this problem. Moreover, if care is taken when handling the slides and changing the buffers the detachment of the agar from the slide is a rare occurrence. 2. Once a lysis buffer has been added, slides should be kept in the dark to prevent any DNA damage caused by fluorescent lighting (14).


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3. DNA damage caused by topoisomerase II inhibitors can be detected as either single or double-strand breaks depending on the buffers used. Typically lower doses are needed to detect single-strand breaks. The advantage of testing double-strand breaks is that this type of damage is more specific for toposiomerase II inhibitors. For example, if the drug tested is a toposiomerase I inhibitor this will cause single-strand breaks but not double-strand breaks. Thus measuring double-strand breaks that occur only when proteinase K is added to the lysis buffer limits the possibilities of proteins that are associated with DNA to those proteins that covalently bind doublestranded DNA. 4. The thickness and percentage of agarose will effect migration of DNA. For these reasons it is important that the agar is completely in solution and that the amount of agar added to each slide is equivalent. 5. Avoid capturing images located around the edges of the slide as those tend to have different lengths compared to the center. 6. To avoid viewer bias it is best to blind the person capturing the images to the conditions. A minimum of 50 images per condition should be analyzed. 7. Avoid capturing images where comets are overlapping. If the slides appear overcrowded with comets it is best to start over as overlapping comets introduce error in calculating comet and tail moments. 8. When quantifying initial DNA damage avoid capturing images that are apoptotic. Apoptotic comets are easily recognized as having fat and long tails with a clear separation of the head indicative of DNA damage due to the activation of caspase dependent endonucleases. 9. The total DNA content (DNA present in tail and head combined) measured in control untreated and treated cells should be similar. If they are not this is typically due to saturation of the comet head and gain settings and exposure times should be adjusted accordingly.

References 1. Kohn, K.W. 1991. Principles and practice of DNA filter elution. Pharmacol Ther 49:55–77. 2. Ostling, O., and Johanson, K.J. 1984. Microelectrophoretic study of radiationinduced DNA damages in individual mammalian cells. Biochem Biophys Res Commun 123:291–298.

3. Tewey, K.M., Rowe, T.C., Yang, L., Halligan, B.D., and Liu, L.F. 1984. Adriamycininduced DNA damage mediated by mammalian DNA topoisomerase II. Science 226:466–468. 4. Hazlehurst, L.A., Krapcho, A.P., and Hacker, M.P. 1995. Comparison of azaanthracenedione-induced DNA damage and

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5.

6.

7.

8.

9.

cytotoxicity in experimental tumor cells. Biochem Pharmacol 50:1087–1094. Hazlehurst, L.A., Valkov, N., Wisner, L., Storey, J.A., Boulware, D., Sullivan, D.M., and Dalton, W.S. 2001. Reduction in druginduced DNA double-strand breaks associated with beta1 integrin-mediated adhesion correlates with drug resistance in U937 cells. Blood 98:1897–1903. Olive, P.L., Banath, J.P., and Durand, R.E. 1990. Detection of etoposide resistance by measuring DNA damage in individual Chinese hamster cells. J Natl Cancer Inst 82:779–783. Pommier, Y., Schwartz, R.E., Zwelling, L.A., Kerrigan, D., Mattern, M.R., Charcosset, J.Y., Jacquemin-Sablon, A., and Kohn, K.W. 1986. Reduced formation of proteinassociated DNA strand breaks in Chinese hamster cells resistant to topoisomerase II inhibitors. Cancer Res 46:611–616. Fox, M.E., and Smith, P.J. 1990. Longterm inhibition of DNA synthesis and the persistence of trapped topoisomerase II complexes in determining the toxicity of the antitumor DNA intercalators mAMSA and mitoxantrone. Cancer Res 50:5813–5818. Emmons, M., Boulware, D., Sullivan, D.M., and Hazlehurst, L.A. 2006. Topoisomerase

10.

11.

12.

13.

14.

II beta levels are a determinant of melphalaninduced DNA crosslinks and sensitivity to cell death. Biochem Pharmacol 72:11–18. Hazlehurst, L.A., Enkemann, S.A., Beam, C.A., Argilagos, R.F., Painter, J., Shain, K.H., Saporta, S., Boulware, D., Moscinski, L., Alsina, M., et al. 2003. Genotypic and phenotypic comparisons of de novo and acquired melphalan resistance in an isogenic multiple myeloma cell line model. Cancer Res 63:7900–7906. O’Connor, P.M., and Kohn, K.W. 1990. Comparative pharmacokinetics of DNA lesion formation and removal following treatment of L1210 cells with nitrogen mustards. Cancer Commun 2:387–394. Olive, P.L., Banath, J.P., and Durand, R.E. 1990. Heterogeneity in radiation-induced DNA damage and repair in tumor and normal cells measured using the "comet" assay. Radiat Res 122:86–94. Kent, C.R., Eady, J.J., Ross, G.M., and Steel, G.G. 1995. The comet moment as a measure of DNA damage in the comet assay. Int J Radiat Biol 67:655–660. Bradley, M.O., Erickson, L.C., and Kohn, K.W. 1978. Non-enzymatic DNA strand breaks induced in mammalian cells by fluorescent light. Biochim Biophys Acta 520:11–20.

Chapter 13 Analysis of Genomic Aberrations Using Comparative Genomic Hybridization of Metaphase Chromosomes Melanie Carless Abstract Comparative genomic hybridization (CGH) allows the global screening of copy number aberrations within a sample. Specifically, large (>20 mb) deletions and amplifications are detected, which are likely to indicate regions harboring tumor suppressor and oncogenes. CGH involves the extraction of test and reference (karyotypically normal) DNA. These samples are whole-genome amplified by DOP-PCR and then differentially labeled with fluorophores via nick translation. Test and reference samples are competitively hybridized to normal metaphase chromosomes. The relative amount of each DNA that binds to a chromosomal locus is indicative of the abundance of that DNA. Thus, if a chromosomal region is amplified, the test DNA will outcompete the reference DNA for binding and fluorescence will indicate amplification. Conversely, if a region is deleted, more reference DNA will bind and fluorescence will indicate a deletion. The following chapter outlines the protocols used for CGH analysis of metaphase chromosomes. These protocols include metaphase chromosome slide preparation, DNA extraction (from blood, cell lines and microdissected formalin-fixed paraffin-embedded tissue), DOP-PCR, nick translation, in situ hybridization and fluorescence microscopy and image analysis. Key words: Comparative genomic hybridization (CGH), Cancer, Microdissection, Degenerate oligonucleotide-primed polymerase chain reaction (DOP-PCR), Nick translation, Fluorescence microscopy.

1. Introduction Comparative genomic hybridization is a molecular cytogenetic technique that enables the global screening of DNA sequence copy number aberrations (CNAs) within a sample in a single experiment. Pioneered by Kallioniemi and colleagues, CGH is used primarily to investigate tumors and detects large copy number changes such as amplifications and deletions that are typically greater than 20 Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_13 Springerprotocols.com

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megabases (mb) in size (1, 2). The principle of this technique is that genomic regions of deletion or amplification are likely to harbor tumor suppressor genes and oncogenes, respectively, which may contribute to a malignant phenotype. Other applications of CGH include prenatal, postnatal and pre-implantation screening (for detection of trisomies, monosomies, etc.) and detection of aberrations related to mental retardation (3). A more recent application of array-based CGH has allowed increased mapping resolution for the detection of locus-by-locus DNA copy number changes (4). CGH analysis involves DNA extraction from both test (usually tumor) and reference (karyotypically normal) samples, followed by universal amplification of the DNA (2). Amplified products then undergo nick translation whereby the test and reference DNA are differentially labeled with fluorescent dyes and are hybridized to normal metaphase chromosomes in the presence of COT-1 blocking DNA (2, 5). The amounts of test and reference DNA that are bound at a chromosomal locus are dependent on the relative abundance of those particular sequences within the two samples (2). Differential fluorescent hybridization signals emitted by the metaphase spread are therefore indicative of chromosomal gains and losses of the test DNA relative to the reference DNA (5). Ratios of the fluorescent signals can be quantified along the length of each chromosome and a copy number karyotype (ratio profile) of the test DNA can be generated (5). CGH can be performed on fresh tissue and also formalin-fixed paraffin-embedded (FFPE) tissue, making it an ideal cytogenetic technique to study solid tumors. The following chapter outlines various protocols used for the detection of genomic aberrations by comparative genomic hybridization. These include metaphase chromosome preparation, DNA extraction methods, degenerate oligonucleotide-primed polymerase chain reaction, nick translation, hybridization and image generation and analysis.

2. Materials All reagents used throughout each of the CGH-related protocols should be of molecular biology grade where applicable. Sterile techniques should always be used in preparation of reagents to prevent contamination during the protocol. Ensure that water used is MilliQ (autoclaved) or DNase/RNase-free. 2.1. Metaphase Chromosome Preparation

1. Sodium heparin vacutainer tube (to collect 10 ml blood), syringe, alcohol wipe (Becton, Dickinson and Company [BD]) 2. 75 cm2 cell culture flasks with vented caps (Corning)

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3. 50 ml centrifuge tubes (Corning, BD) 4. 37C incubator with 5% CO2, biological safety cabinet 5. 100 ml lymphocyte media (store at 4C; warm to 37C prior to addition of blood): 86.5 ml 1 RPMI media with L-glutamine (2 mM) (Gibco; store at 4C); 2.5 ml 1 M HEPES buffer (25 mM final concentration) (Gibco; store at 4C); 10 ml fetal calf serum (10% final concentration) (Gibco; store in aliquots at –20C); 1 ml 10,000 U penicillin/10,000 mg streptomycin (the final concentration is 100 U/100 mg) (Gibco; store in aliquots at –20C) 6. Phytohemagglutinin (M-form) (PHA, unknown) (Gibco; store at –20C)

concentration

7. 1 Hank’s buffered saline solution (HBSS) (Gibco) 8. Methotrexate (Sigma-Aldrich; powder, soluble in 0.1 M NaOH; toxic); prepare a stock solution using a minimum volume of 1 M NaOH and then dilute to 10–5 M with HBSS (store in aliquots at –20C in dark) 9. Thymidine (Sigma-Aldrich; powder); prepare a stock solution using a minimum volume of 1 M NaOH and then dilute to 10–3 M with HBSS (store at room temperature) 10. 10 mg/ml KaryoMAX-Colcemid1 (in HBSS) (Gibco; store at 4C) 11. Water bath (set to 37C) 12. Fixative (3:1 methanol/glacial acetic acid; prepare fresh; reagents available from Sigma-Aldrich) 13. SuperFrost slides (Fisher) 14. DAPI/Vectashield mix: add 35 ml 1 mg/ml DAPI to 965 ml Vectashield (0.35 mg/ml final concentration) (store 4C in the dark up to 2 months, after this time additional DAPI can be added to the solution); 1 mg/ml DAPI (40 ,6-diamidino2-phenylindole dihydrochloride) (Sigma-Aldrich; powder; when in liquid form store in aliquots at –20C in dark; irritant); Vectashield (Vector Laboratories; store at 4C in dark; irritant) 15. 25 25 mm coverslip 16. Fluorescent microscope (Olympus, Nikon, Leica) 17. Coplin jars (Laboratory Equipment World) 18. 20 SSC (Sigma-Aldrich); or make solution: 0.3 M sodium citrate, 3 M NaCl 19. 2 SSC: dilute 100 ml of 20 SSC with 900 ml MilliQ water 20. Denaturation solution (70% formamide in 2 SSC, pH 7.0); 35 ml formamide (Sigma-Aldrich; store at 4C; toxic); 15 ml 2 SCC buffer

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2.2. Sample Preparation and DNA Extraction

1. Lithium heparin vacutainer tube (to collect 10 ml blood), syringe, alcohol wipe (BD) 2. 50 ml centrifuge tubes (Corning, BD) 3. 15 ml centrifuge tubes (Corning, BD)

2.2.1. DNA Extraction from Blood/ Cell Lines

4. NKM buffer: 140 mM NaCl, 30 mM KCl, 3 mM MgCl2 5. RSB buffer: 10 mM Tris–HCl pH 7.5, 1 mM NaCl, 3 mM MgCl2 6. Plastic transfer pipettes 7. Lympholysis buffer: 25 mM Tris pH 7.5, 25 mM EDTA, 75 mM NaCl, 1% SDS 8. Proteinase K buffer: 2 mg/ml proteinase K, 10 mM CaCl2, 10 mM Tris pH 7.5 9. 6 M NaCl 10. Ethanol 11. Plastic inoculating loops 12. Microfuge tube 13. TE buffer: 10 mM Tris pH 7.2, 1 mM EDTA 14. ND-1000 spectrophotometer (or similar) (NanoDrop) 15. Complete cell culture media (e.g., 1 RPMI containing 10% FBS and 1% antibiotic); media (RPMI, DMEM, other) (Gibco; store at 4C); fetal bovine serum (FBS) (Gibco; store in aliquots at –20C); penicillin/streptomycin (10,000 U/10,000 mg) (Gibco; store in aliquots at –20C) 16. 75 cm2 flasks (Corning) 17. Phosphate-buffered saline (PBS) (Gibco; store at 4C or room temperature) 18. 0.05% trypsin–EDTA (Gibco; store temporarily at 4C or at –20C for longer periods) 19. Inverted phase-contrast microscope (Nikon, Olympus) 20. Positive control cell line such as MCF7 or MDA-MB-231

2.2.2. DNA Extraction from Formalin-Fixed Paraffin-Embedded Tissue

1. Formalin-fixed paraffin-embedded tissue sections, 6–12 sections, 5 mm thick on Superfrost slides (Fisher) (samples obtained from a pathologist) 2. Xylene (Sigma-Aldrich; toxic, possible carcinogen) 3. Ethanol (Sigma-Aldrich; toxic) 4. Mayer’s hematoxylin (Sigma-Aldrich; toxic) 5. Eosin Y solution (Sigma-Aldrich; toxic) 6. 22 60 mm coverslips (Fisher Scientific) 7. Histomount (Invitrogen; toxic, possible carcinogen; cap after use to prevent drying)


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8. Methyl green (Sigma-Aldrich; irritant); add 0.5 g of methyl green to 50 ml MilliQ water 9. Stereomicroscope (e.g., SMZ800 from Nikon) 10. Scalpel blade (#11) (Fisher) 11. PCR tubes (Eppendorf) 12. 20 mg/ml proteinase K (from Tritirachium album; SigmaAldrich; toxic) 13. Proteinase K Tween buffer (PKTB): 100 ml Taq polymerase reaction buffer (used in Section 3.3, reagent 5.) (Roche Applied Science; store at –20C); 20 ml 20 mg/ml proteinase K (0.4 mg/ml final concentration); 5 ml Tween 20 (0.5% final concentration) (Sigma-Aldrich; toxic); 875 ml DNase/ RNase-free water. 14. Thermocycler (Applied Biosystems) 15. Mineral oil (autoclaved and filtered) (Sigma-Aldrich)

2.3. Degenerate OligonucleotidePrimed Polymerase Chain Reaction (DOP-PCR)

1. PCR tubes 2. Aerosol filter barrier tips 3. Extracted DNA (FFPE tissue – 1 ml; fresh tissue or blood – 50 ng in 1 ml) 4. PCR tubes (Eppendorf) 5. 32 U/ml Thermo Sequenase DNA polymerase (Amersham; store at –20C) Includes 10 reaction buffer: 260 mM Tris–HCL (pH 9.5), 65 mM MgCl2 6. 5 U/ml Taq DNA polymerase (Roche Applied Science; store at –20C) Includes 10 reaction buffer: 100 mM Tris–HCl (pH 8.3), 15 mM MgCl2, 500 mM KCl 7. 10 mM UN-1 degenerate primer (50 -CCGACTCGAGN NNNNNATGTGG-30 ) 8. 10 mM dNTP mix (Invitrogen) 9. UltraPure DNase/RNase-free water (Gibco) 10. Mineral oil (autoclaved and filtered) (Sigma-Aldrich) 11. Thermocycler (Applied Biosystems) 12. Parafilm (Fisher)

2.4. Nick Translation

1. 10 dNTP mix 10 ml each of 10 mM dATP, dCTP, dGTP (200 mM final concentration) (Promega, Invitrogen; store at –20C, avoid freeze/thaw cycles); 250 ml 1 M Tris, pH 7.8 (500 mM final concentration) (Sigma-Aldrich; require HCl for pH

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adjustment); 25 ml 1 M MgCl2 (50 mM final concentration) (Sigma-Aldrich); 3.5 ml 14.3 M b-mercaptoethanol (100 mM final concentration) (Sigma-Aldrich; toxic); 1 ml 50 mg/ml bovine serum albumin (BSA, 100 mg/ml final concentration) (Invitrogen; store at –20C); 190.5 ml UltraPure DNase/RNase-free water. 2. Fluoroscein-12-dUTP (FITC-dUTP) (Perkin–Elmer; store at –20C in dark) 3. Texas Red-dUTP (TR-dUTP) (Perkin–Elmer; store at –20C in dark) 4. Bionick enzyme mix (Invitrogen: Bionick DNA labeling system; store at –20C; DNase I is sensitive to physical denaturation, do not vortex) Contains: 0.5 U/ml DNA polymerase I; 0.007 U/ml DNase I; 50 mM Tris–HCl, pH 7.5; 5 mM magnesium chloride; 0.1 mM phenylmethylsulfonyl fluoride (PMSF); 50% (v/v) glycerol; 100 mg/ml nuclease-free BSA 5. DNA Polymerase I/DNase I (Invitrogen; store at –20C; DNase I is sensitive to physical denaturation, do not vortex); contains: 0.5 U/ml DNA Polymerase I; 0.4 mU/ml DNase I; 50 mM Tris–HCl, pH 7.5; 5 mM magnesium acetate; 1 mM 2-mercaptoethanol; 0.1 mM PMSF; 100 mg/ml BSA; 50% (v/v) glycerol 6. 10 U/ml DNA polymerase (Invitrogen; store at –20C) 7. UltraPure DNase/RNase-free water

2.5. Agarose Gel Electrophoresis

1. Analytical Grade Agarose (Promega, Sigma-Aldrich) 2. 1 TAE: add 975 ml MilliQ water to 25 ml 40 TAE (Promega); 40 TAE: 1.6 M Tris–acetate, 40 mM EDTA 3. Conical flask 4. 10 mg/ml ethidium bromide solution (Promega, SigmaAldrich; mutagenic, store in dark) 5. 6 gel loading dye 6. 100 bp molecular weight marker (containing loading dye) (NEB, Qiagen, Promega) 7. Gel tray and combs (7 cm 10 cm tray and 1 11 teeth comb is sufficient for 10 samples, if analyzing more samples than this you may need to run a larger gel or multiple small gels) 8. Electrophoresis tank and power source 9. UV light box and camera (ensure glasses/face shield is worn when using UV light box)


2.6. In Situ Hybridization

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1. Coplin jars (Laboratory Equipment World) 2. Test and reference DNA (labeled by nick translation) 3. 1 mg/ml COT-1 DNA (Invitrogen; store at –20C) 4. 3 M sodium acetate, pH 5.2 (Sigma-Aldrich; toxic irritant) 5. Ethanol (Sigma-Aldrich; toxic) 6. 20 SSC (Sigma-Aldrich); optional laboratory made: 0.3 M sodium citrate, 3 M NaCl 7. 2 SSC (dilute 100 ml of 20 SSC with 900 ml MilliQ water) 8. Hybridization buffer: 50% formamide, 10% dextran sulfate, 2 SCC, pH 7.0 500 ml formamide (Sigma-Aldrich; store at 4C; toxic); 100 mg dextran sulfate sodium salt (Sigma-Aldrich; possibly toxic); 2 SSC up to 1 ml (approximately 400 ml; ensure that 20 SCC is diluted with DNase/RNase-free water for this buffer) 9. Denaturation solution: 70% formamide in 2 SSC, pH 7.0; 35 ml formamide (Sigma-Aldrich; store at 4C; toxic); 15 ml 2 SCC buffer 10. Metaphase slides (made in Section 3.1 or purchased from Vysis; store at –20C) 11. Diamond-tipped scribe 12. Slide warmer (LabScientific, Cole-Parmer, Electron Microscopy Sciences) 13. Rubber cement (Fisher Scientific) 14. Chamber with rack – must be a dark box or kept in the dark Containing 2 SSC-soaked paper towel on bottom for humidity control (e.g., Thermo Scientific Hybaid Humidity Chambers from Fisher Scientific); alternatively these can be made from any box with a rack to fit slides horizontally. If the container is not completely air tight paper towels may have to be re-soaked with 2 SSC each day if evaporation occurs. 15. 37C oven (Fisher Scientific) 16. 3 50 ml wash solution (50% formamide in 2 SCC) (75 ml formamide [Sigma-Aldrich; store at 4C]; 75 ml 2 SSC) 17. 1 50 ml PN buffer (0.1 M NaH2PO4, 0.1 M Na2HPO4, 0.1% NP-40 pH 8.0) (5 ml 1 M NaH2PO4 (monobasic; Sigma-Aldrich; irritant, toxic); 5 ml 1 M Na2HPO4 (dibasic; Sigma-Aldrich; irritant, toxic); 50 ml nonidet P40 (NP40, Igepal1CA-630) (Sigma-Aldrich; toxic, irritant) 18. DAPI/Vectashield mix: add 35 ml 1 mg/ml DAPI to 965 ml Vectashield (0.35 mg/ml final concentration) (store 4C in

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the dark up to 2 months, after this time additional DAPI can be added to the solution) 1 mg/ml DAPI (40 ,6-diamidino-2-phenylindole dihydrochloride) (Sigma-Aldrich, powder form; once in liquid form store in aliquots at –20C in dark; irritant) Vectashield (Vector Laboratories; store at 4C in dark; irritant) 19. 25 mm 25 mm coverslip (Fisher) 2.7. Fluorescence Microscopy and Image Analysis

1. Immersion oil (Sigma-Aldrich; irritant) 2. Fluorescent microscope with mercury burner lamp (Olympus, Leica, Nikon) 3. Fluorescent filter set (specific for fluorophores used) (Chroma Technology, Nikon) 4. High-performance Optronics)

black/white

CCD

camera

(Cohu,

5. Software package (e.g., PowerGene MacProbe or CytoVision software from Applied Imaging)

3. Methods 3.1. Metaphase Chromosome Preparation

Metaphase chromosome preparation is crucial for CGH analysis; an adequate number of cells must be in metaphase and chromosomes must be well spaced (not overlapping) and of sufficient size. Metaphase chromosome spreads are obtained from short-term cultured blood samples, whereby cell division is stimulated by phytohemagglutinin (PHA), cells are synchronized (DNA synthesis is blocked in S phase by methotrexate and released by thymidine) and mitosis is halted in metaphase by addition of colcemid (6). This protocol typically yields an increased number of mitotic cells (in metaphase) and results in improved chromosome morphology (6). The following protocol is modified from those presented in The AGT Cytogenetics Laboratory Manual (7). Even when this protocol is optimized and followed, good-quality metaphase spreads suitable for hybridization are not guaranteed. Alternatively, metaphase chromosome spreads can be purchased from Vysis. Although these are also not guaranteed to give good results, the slides typically provide quality metaphase spreads and are often the preferred choice. This protocol should be carried out in a biological safety cabinet to prevent contamination. Ensure sterile cell culture techniques.


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1. Collect 10 ml blood into a sodium heparin tube. Blood must come from a healthy volunteer who is karyotypically normal. 2. Add 2 ml of blood to each of four 75 cm2 cell culture flasks containing 25 ml lymphocyte media and 750 ml PHA (concentration 3% v/v). 3. Incubate flasks for 48 h in a 37C incubator with 5% CO2. 4. Add 278 ml HBSS containing 10–5 M methotrexate (10–7 M final concentration); alternatively, you can leave one or two of the flasks unsynchronized. 5. Return flasks to incubator for a further 17 h. 6. Transfer cultures to 50 ml conical tubes and centrifuge at 800 rpm for 8 min. 7. Remove media, loosen cell pellet and resuspend in 25 ml prewarmed (37C) lymphocyte media containing 250 ml 1 10–3 M thymidine (10–5 M final concentration). 8. Return cultures to the incubator for a further 5 h. 9. Add 25 ml 10 mg/ml KaryoMAX-Colcemid1 (10 ng/ml final concentration) and incubate for 10–15 min at 37C. It is advised to use different times for each of the flasks, for example, 10 and 15 min each for synchronized and unsynchronized cells. 10. Transfer cultures to 50 ml conical tubes and centrifuge at 800 rpm for 8 min. 11. Discard all but 500 ml of the media and resuspend the cell pellet in this. 12. Add 10 ml pre-warmed (37C) 750 mM KCl solution (to swell cells) and incubate at 37C for 20 min (in water bath). 13. Centrifuge the tubes at 800 rpm for 5 min, discard the supernatant (except for 500 ml) and resuspend the cell pellet. 14. Add 10 ml fresh fixative (add the first 2 ml drop-wise while gently shaking the tube by hand). 15. Repeat steps 13–14 several more times (usually 3 or 4) until the fixative becomes clear and the cell pellet is opaque in color. 16. Resuspend the pellet in 5 ml fixative (see Note 1). 17. Microscope slides should be cleaned with 100% ethanol and dried vertically. 18. Prepare one slide for each of the four tubes: using a plastic pipette place two separate drops of cells/fixative on the slide (well spaced, these are the two target areas) and allow the cells to air-dry (about 30 s) (see Note 2).

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19. Stain each target area with 20 ml DAPI/Vectashield mix (0.35 mg/ml) and visualize with fluorescence and phasecontrast microscopy (under 10 , 40 and 60 objectives) (see Note 2). 20. Once the most appropriate condition has been identified, prepare all other slides like this, at least 50 slides should be prepared. Slides should be stored at room temperature for 2 weeks to age. 21. After aging, denature one of the slides in denaturation solution (in a coplin jar) for 5 min at 73–1C, stain with DAPI (step 19) and view banding pattern (see Note 3). Ensure that slides are appropriate for analysis. 22. Store aged slides at –20C (to prevent further aging). 3.2. Sample Preparation and DNA Extraction

CGH is useful for the detection of copy number aberrations in diseased samples from any tissue source. Blood provides a good source of DNA for analysis but often extraction of DNA from solid tissue is more difficult due to limited availability of tissue or the requirement of formalin-fixed paraffin-embedded tissue. Here are methods of DNA extraction from whole blood (required for reference DNA), cell lines (required for positive controls) and formalin-fixed paraffin-embedded tissue (test DNA).

3.2.1. DNA Extraction from Blood

Acquisition and testing of both male and female DNA is necessary for the generation of a ratio threshold and dynamic range. DNA therefore needs to be extracted from at least one male and one female (both karyotypically normal). The following protocol is a modification of that developed by Miller and colleagues (8). This is a salting-out procedure whereby red blood cells are lysed and removed using salt solutions, white blood cells are lysed and proteins are degraded and removed, and ethanol is used to precipitate the DNA. 1. Collect 5–10 ml blood into a lithium heparin tube. Blood must come from healthy volunteers (one male, one female), who are karyotypically normal. 2. Transfer the blood to a 50 ml tube and add 40 ml NKM buffer. 3. Mix vigorously, then centrifuge at 4,800 rpm for 25 min at room temperature. 4. Using a plastic pipette, carefully pipette off the supernatant (containing lysed red blood cells), making sure that the white cell layer remains intact. Tap the tube to loosen the cell pellet. 5. Add a small amount of RSB buffer, resuspend the cell pellet well and then add RSB buffer up to a final volume of 50 ml. This will lyse the remaining red cells.


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6. Mix well then centrifuge at room temperature (RT), 4,000 rpm for 15 min. 7. Carefully pipette off the supernatant and tap the tube to loosen the cell pellet. 8. Repeat steps 5–7 once or twice until most red blood cells have been lysed and removed and the pellet appears to be white in colour. Ensure that the cell pellet is resuspended well in a small amount of RSB buffer (about 500 ml). 9. Add 8 ml lympholysis buffer and 500 ml proteinase K buffer to lyse the white cells. 10. Shake vigorously at 37C overnight. 11. Add 4 ml of sterile saturated NaC1 (approx. 6 M), mix vigorously for 15 s. 12. Centrifuge at 2,500 rpm at 4C for 15 min to pellet cellular proteins. 13. Carefully decant the supernatant containing the DNA into a 15 ml centrifuge tube. 14. Centrifuge the 15 ml tube at 2,500 rpm at 4C for 15 min to pellet any remaining proteins. 15. Using a plastic pipette carefully transfer the supernatant to a 50 ml centrifuge tube, leaving the protein undisturbed as a pellet. 16. Measure the volume and add exactly two volumes of chilled absolute ethanol. 17. Gently swirl and observe the DNA strands precipitate. 18. Using a plastic loop, carefully transfer the DNA precipitate into a microfuge tube containing 500 ml DNase/RNase-free water and incubate at 37C 4–6 h or until DNA is dissolved. 19. Measure the concentration and quality of the DNA using a spectrophotometer (260/280 absorbance ratio should be about 1.7). The DNA quality can also be viewed via agarose gel electrophoresis (see Section 3.5). Adjust the concentration of samples to 50 ng/ml before storing. 3.2.2. DNA Extraction from Cell Lines

To ensure that your protocol is working and for quality control across experiments, DNA from cell lines can be used as a positive control in CGH. Two commonly used positive control cell lines are MCF7 and MDA-MB-231, although any cell line can be used if the copy number abnormalities are known. Most cell lines, including MCF7 and MDA-MB-231, can be purchased from the American Type Culture Collection (ATCC). Cells should be cultured in pre-warmed (37C) media using aseptic techniques, according to the vendors’ instructions. 1. In a 37C 5% CO2 incubator, grow cell lines to 80% confluency in one or more 75 cm2 flasks using complete media.

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For MCF7 and MDA-MB-231 1 RPMI media containing 10% FBS and 1% antibiotic can be used, although many other media formulations could be substituted. Other cell lines may require a different media. 2. Remove the media and wash the cells once with 8 ml prewarmed (37C) PBS. 3. Add 4 ml trypsin and put the flask in the 37C incubator for about 2–5 min. Remove the cells when they are rounded up and starting to lift from the flask (by viewing under a phase-contrast inverted microscope) and add 6 ml complete media. 4. Tap the flask to remove cells from the flask wall and put media (containing cells) into a 50 ml centrifuge tube. 5. Centrifuge at 1,000 rpm for 5 min to pellet the cells and remove media. 6. Extract the DNA as per instructions in Section 3.2.1, starting at step 9 (half volumes may be used if the pellet is small). 3.2.3. DNA Extraction from ParaffinEmbedded Tissue

Formalin-fixed paraffin-embedded tissue can be obtained from most pathologists. To define cancerous tissue (versus normal tissue), you should consult the pathologist. If contaminating normal tissue is microdissected along with the cancerous tissue the sample will be diluted and false negative results might be obtained. It is therefore very important that the samples are microdissected with care, and if possible photos should be taken before and after microdissection for comparison. Typically 6–12 slides of 5 mm thickness are collected. Of these, 2–3 are used as ‘‘guide’’ slides and are stained with hematoxylin and eosin, and the remaining slides are stained with methyl green for microdissection. The guide slides should be from either end of the sections and if 12 sections are used, a third guide slide from the middle of the sections should also be used. Following microdissection, DNA is extracted by digesting proteins in a proteinase K buffer. This protocol outlines staining techniques, microdissection and DNA extraction from paraffin-embedded, formalin-fixed samples. Some investigators also have access to a microscope capable of laser capture microdissection, which can isolate individual cells for analysis. 1. De-paraffinise and stain 2–3 slides with hematoxylin and eosin: Immerse in xylene twice (3 min each), an ethanol series (100, 100, 90 and 70% for 1 min each), rinse in running water, hematoxylin (5 min), rinse in running water, eosin (2 min), rinse in running water (1 min), xylene twice (2 min each); apply a coverslip with Histamount. 2. De-paraffinise and stain the remaining slides with methyl green:


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Immerse in xylene twice (3 min each), an ethanol series (100, 95, 70, 50 and MilliQ water for 2 min each), 1% methyl green (5 min), rinse in MilliQ water (1–5 min); do not apply a coverslip. 3. Using a stereomicroscope and the hematoxylin- and eosinstained slides as a guide, microdissect the tumor tissue by hand from the methyl green-stained slide using a scalpel blade (see Note 4). If possible, take photos of the slide before and after microdissection. 4. Place the microdissected tissue into a PCR tube containing 15–30 ml PKTB (volume dependent on the size of the tumor sample). 5. Overlay the samples with 50 ml mineral oil. 6. Incubate samples at 55C for 3 days, each day add 0.6 ml fresh 20 mg/ml proteinase K to each sample. 7. On day 3, denature the proteinase K by heating samples to 95C for 10 min. 8. Remove oil from the sample by rolling on Parafilm, store the samples at 4C. 3.3. Degenerate OligonucleotidePrimed Polymerase Chain Reaction (DOP-PCR)

Universal amplification of DNA is performed using degenerate oligonucleotide-primed polymerase chain reaction (DOP-PCR). Briefly this involves an initial low stringency pre-amplification using Thermo Sequenase DNA polymerase (5 ml volume) followed by a high-stringency amplification using Taq DNA polymerase (50 ml volume). The protocol below is a modification of the protocol provided by Kuukasja¨rvi et al. 1997 (9). To prevent contamination, these steps should be carried out in a laminar flow hood using aerosol filter barrier tips. As with any PCR, when making the bulk mixes you should always use DNase/RNasefree water, adding this first to the bulk mix tube and the enzyme last. Also, a negative control (containing no DNA) should be used for each experiment. 1. Aliquot 1 ml (see Note 5) of each of your extracted DNA samples into a suitable PCR tube. It is recommended to perform DOP-PCR on multiple samples at once but this should be limited to 12 or 13 at the most. For each DOP-PCR experiment various controls should be run, including a negative control (no DNA template), a positive control (cell line such as MDA-MB-231 or MCF7, with known chromosomal abnormalities) and sufficient reference DNA for your experiments (typically, one reference PCR sample is sufficient for 2 experiments – see Note 5). Reference DNA is considered to be DNA extracted from an individual (male or female) with no cytogenetic abnormalities.

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2. Prepare a pre-amplification bulk mix sufficient for all samples to be DOP-PCRed, including a little extra for pipetting inaccuracies (e.g., 13.5 bulk mix for 13 samples):

Final concentration

1 Bulk mix (ml)

10 thermo sequenase buffer

1

0.5

6.75

10 mM dNTPs

0.2 mM

0.1

1.35

10 mM UN-1 primer

1.2 mM

0.6

8.1

32 U/ml thermo sequenase

2U

0.0625

0.84

DNase/RNase-free water

–

2.3375

31.56

4

54

Reagent

Final volume

13.5 Bulk mix (ml)

3. Add 4 ml of your pre-amplification bulk mix to each of your DNA samples. 4. Overlay each sample with about 50 ml mineral oil. 5. Place samples into the thermocycler for low stringency preamplification using the following cycling conditions: initial denaturation at 96C for 3 min; five cycles of 94C for 1 min, 25C for 1 min, a slow ramp to 74C over 3 min, 74C for 2 min; final extension at 74C for 10 min. Prior to adding the amplification mix to your samples, you should denature your samples at 95C for 5 min. 6. While pre-amplification thermo-cycling is occurring prepare your amplification mix for high-stringency amplification, although make sure you add the Taq DNA polymerase enzyme just prior to completion of the preamplification cycling (i.e., during the 5 min denaturation step):

Reagent

Final concentration

1 Bulk mix (ml)

13.5 Bulk mix (ml)

10 Taq polymerase buffer

1

4.5

60.75

10 mM dNTPs

0.2 mM

0.9

12.15


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(continued) 1 Bulk mix (ml)

13.5 Bulk mix (ml)

Reagent

Final concentration

10 mM UN-1 primer

1.3 mM

5.85

78.975

5 U/ml Taq DNA polymerase

2U

0.4

5.4


–

29.75

401.625

45

607.5

Final volume

7. Add 45 ml of your amplification bulk mix to each of your DNA samples. Vortex briefly and pulse centrifuge (it is important to do this step as quickly as possible). 8. Return the PCR tubes to the thermocycler for high-stringency amplification using the following cycling conditions: initial denaturation at 95C for 5 min; 35 cycles of 94C for 1 min, 56C for 1 min, 72C for 3 min; final extension at 72C for 5 min. 9. Electrophorese 5 ml of each sample on a 2% agarose gel for 45–60 min to determine if contamination has occurred and that amplified DNA is of an appropriate length (see Note 6). See Section 3.5 for agarose gel preparation. Figure 13.1 shows an example of typical DOP-PCR products.

3.4. Nick Translation

Comparative genomic hybridization quality is highly dependent upon the size of DNA fragments. Nick translation is the process by which DNA is digested to produce smaller fragments while incorporating a fluorescent dye. DNase I cleaves the DNA, while DNA polymerase adds nucleotides to the strand, incorporating the fluorescently labeled dUTP. The following protocol is a modification of the nick translation method outlined by Kallioneimi et al. (1, 2). This protocol uses FITC and Texas Red fluorophores, although many other fluorophores can be used for labeling providing the microscope used for detection has the appropriate excitation and emission filters and that these do not overlap greatly. Some protocols use an indirect labeling protocol (incorporating biotin), which may increase the signal. However, direct labeling limits background signal and usually provides signals of adequate intensity for analysis. As such, this is the protocol that is given here.

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Fig. 13.1. Typical DOP-PCR results. Lanes 1 and 2 show a distinct band at about 400 bp, which is postulated to signify preferential amplification of repetitive sequences, DOPPCR products exhibiting this band can still be used for CGH analysis. Lane 3 is the negative control. Lanes 4, 5, 8, 9 and 10 show amplification of DNA extracted from fresh tissue. Lane 7 shows amplification of microdissected DNA. Lane 6 is the molecular weight marker.

1. Prepare sufficient bulk mix for your samples:

Reagent

1 Bulk mix (ml)

13.5 Bulk mix (ml)

10 dNTP mix

5

67.5

10 Bionick enzyme mix

1.5

20.25

DNA polymerase I/DNase I enzyme mix

1.5

20.25

DNA polymerase

1.0

13.5

dUTP (FITC- or Texas Redlabeled)

1.0

13.5

Final volume

10

135.0


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Fig. 13.2. Typical nick translation products. Lane 1 is the molecular weight marker. Lane 2 is labeled with Texas Red and lanes 3 and 4 are labeled with FITC. Both dyes fluoresce under UV light and as such can be identified in the gel picture.

2. For DOP-PCRed samples obtained from fresh tissue add 10 ml bulk mix and 17.5 ml DNase/RNase-free water to 22.5 ml sample. For DOP-PCRed samples obtained from microdissected tissue add 10 ml bulk mix to 40 ml sample. (see Note 7 for additional comments on nick translation). 3. Incubate at 15C for 60 min and halt the reaction by heating the samples to 75C for 15 min (see Note 7). Nick translation products must be stored in the dark at –20C to prevent degradation of both the product and the fluorophore. 4. Electrophorese 5 ml of each sample on a 2% agarose gel at 90 V for 45–60 min to ensure nick translated product is of appropriate length (see Note 7). See Section 3.5 for agarose gel preparation. Figure 13.2 shows an example of typical nick translated products. 3.5. Agarose Gel Electrophoresis

Agarose gel electrophoresis can be used to determine the quality of DNA, DOP-PCR amplified products and nick translated products used for analysis. Electrophoresis is carried out using 2% agarose gels. The following protocol is sufficient for one 7 cm 10 cm tray, if a larger gel is needed adjust the volumes accordingly. 1. Add 50 ml 1 TAE to 1.0 g of agarose in a conical flask and heat in microwave until agarose is well dissolved (about 2 min, ensure it does not boil over).

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2. When cooled, add 2.5 ml 10 mg/ml ethidium bromide (0.5 mg/ ml final) and mix gently. 3. Pour agarose solution into gel tray and insert comb, allow gel to set before loading samples. 4. While gel is setting, add 1 ml of 6 loading dye to 5 ml of your sample. 5. Remove comb from gel when set: load 5 ml 100 bp molecular weight marker (containing loading dye) in the first lane then load all samples and your negative control. 6. Place gel into electrophoresis tank and run the gel at 80–90 V for about 45–60 min or until you can clearly see the size range of your DNA smears using the UV light box. To test for DNA quality 20–30 min is sufficient. 7. Capture an image of your gel for future records.

3.6. In Situ Hybridization

Labeled test and reference DNA are precipitated with COT-1 DNA to generate a probe for the hybridization. The probe and metaphase chromosomes are both denatured prior to hybridization. Following hybridization, slides are washed to remove excess probe and DAPI is added in the presence of an antifade solution. All denaturation and washing of the slides is done in coplin jars and no more than four slides should be processed at a time. Metaphase slides can be made in the laboratory or purchased from Vysis. Each slide has two target areas and thus two experiments can be performed per slide. An initial experiment should be performed by hybridizing differentially labeled male and female DNA (normal karyotype) to generate a ratio threshold. Also, positive control hybridizations should be performed incorporating a cell line with known cytogenetic aberrations and a reference DNA. This will indicate how sensitive your technique is and can also be used as a quality control hybridization across experiments. 1. Pre-warm denaturation solution in a coplin jar to 73–1C. 2. Precipitate the labeled test and reference DNA with COT-1 DNA by combining 22.5 ml of the test DNA and 40 ml of the reference DNA (labeled by nick translation) with 30 ml of COT-1 DNA, 9.5 ml 3.0 M sodium acetate (pH 5.2) and 237.5 ml cold 100% ethanol. 3. Vortex the mix briefly, freeze in liquid nitrogen and store at –80C for 2–3 h. 4. Centrifuge at 11,000 rpm at 4C for 30 min, remove ethanol and air-dry the pellet in the dark. 5. Resuspend the pellet/probe in 10 ml hybridization buffer. The probes will need to be heat denatured at 73C for 5 min just prior to hybridization to the target metaphase


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slide (while metaphase slides are drying – step 8 later in this protocol). 6. Label target metaphase slides (using pencil, not pen) and circle the target areas (underneath the slide) using a diamond-tipped scribe. 7. Soften target metaphase slides in 2 SCC at 37C for 5 min then dehydrate them using an ethanol series of 70, 85 and 100% for 2 min each (RT). 8. Place slides in the denaturation solution at 73–1C for 5 min (– 1 min) (see Note 8) then dehydrate the slides using an ethanol series of 70, 85 and 100% for 2 min each (RT). Slides are air dried vertically on paper towel for 10 min. 9. Place dried slides on a slide warmer set to 37C, then drop the entire denatured probe (10 ml) onto a target area (two probes per slide). Cover each target area with a 22 22 mm square coverslip (two per slide), ensuring that no air bubbles are formed (see Note 9). Seal all edges using rubber cement. 10. Place slides in a sealed humidified chamber and into a hybridization oven set to 37C for 3 days. 11. Following hybridization, remove coverslips and rubber cement from slides with a pair of tweezers (the rubber cement will be soft and will come off easily). 12. Wash slides three times in wash solution (50% formamide) at 45C for 10 min each (to denature poorly bound probe). 13. Wash slides in 2 SCC for 10 min at 45C then for 10 min at RT. 14. Wash slides in PN buffer for 10 min at RT (to remove poorly bound probe). 15. Rinse slides in DNase/RNase-free water for 5 min and dry slides vertically in the dark for 5–10 min or until dry. 16. Counterstain metaphase chromosomes by adding 20 ml DAPI/Vectashield mix and placing 22 mm 22 mm coverslips on each of the target areas. Store slides at 4C in the dark until ready for digital analysis. (see Note 10). 3.7. Fluorescence Microscopy and Image Analysis

Images of the DAPI, FITC and Texas Red stains can be captured using a fluorescence microscope and a CCD camera. An initial experiment, performed by hybridizing differentially labeled male and female DNA (normal karyotype), should generate a ratio threshold between 0.8 and 1.2, as seen on all chromosomes except for the X chromosome. This means that normal variation will fall between 0.8 and 1.2 (intensity is normalized to 1.0). Deviations from this ratio threshold indicate deletions (those below 0.8) or amplifications (those above 1.2). A male versus female

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hybridization should clearly show an amplification of the X chromosome (if female is considered as test). Analysis of positive control cell lines should reveal cytogenetic aberrations typical of that cell line. The ATCC usually contains some information or references to information on cytogenetic abnormalities associated with each cell line. However, it is important to remember that cytogenetic anomalies will vary (typically increasing in number) with increasing passage number of the cell line. Each metaphase spread contains two homologues of each chromosome, therefore the ratio profile generated for each metaphase spread is an average of these. An average (mean) ratio profile is generated from 6 to 12 homologous chromosomes derived from 5 to 10 metaphase spreads across each hybridization. The average fluorescence intensity –1 standard deviation is displayed for each region of all chromosomes. Copy number changes are assumed to be real if both the mean and one standard deviation are either above or below the set ratio threshold, with the other standard deviation above or below a ratio of one, respectively. Results need to be interpreted with caution in repetitive regions (e.g., centromeres) and also in regions affected by fluorophore artifacts (see final comments in Note 7). 1. All digital analysis should be performed in a dark room to prevent degradation of the fluorophores. Turn on the mercury burner lamp and allow it to stabilize (about 5 min) then place the slide on the microscope. 2. Using the 10 objective lens first, locate metaphase spreads using the DAPI excitation filter (see Note 11). Once located, increase the magnification to 40 and then 60 (using oil immersion). Each time the magnification changes, the metaphase spread of interest needs to be re-centered and focused. Before capturing the images, ensure that the metaphase spread is of good quality (see Note 11). Figure 13.3 gives an example of good quality metaphase spread.

Fig. 13.3. Example of a good-quality metaphase spread. Chromosomes should be long and banding should be distinct.


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3. Ensure the microscope is in camera mode. Using your chosen software package, a live preview can be used to finely focus and center the metaphase spread image. (see Note 12). 4. Using the computer software, capture an image using the DAPI excitation filter (approximately 0.5 s exposure time), this will be pseudo-colored blue. Capture additional images with the FITC (about 6 s exposure, pseudo-colored green) and Texas Red (about 2 s exposure, pseudo-colored red) excitation filters. (see Note 13) Although the images are analyzed in gray scale, a red/green/blue (RGB) composite image can be displayed at this point. 5. Image processing (by the software package) allows background flattening and subtraction, standardization of the red and green signal intensities to 1.0 (to limit differences in hybridization intensities across metaphase spreads) and grayscale conversion. 6. The DAPI stain produces a weak G-banding pattern that can be enhanced (by the user) for karyotyping. After enhancement, chromosomes (and often non-chromosomal material) are automatically selected. De-selection of non-chromosomal material can be done manually. Although bad quality or overlapping chromosomes can be de-selected, it is often best to leave them as they can aid in karyotyping of other chromosomes. Also, touching chromosomes or single chromosomes that have been selected as two (due to light banding) can be outlined manually. 7. An automated karyotype is generated based on pattern recognition, however this usually requires further manual manipulation. Correction of mis-called chromosomes and adjustment of chromosome orientation is done manually and any chromosomes that cannot be reliably interpreted are excluded from analysis. For help in karyotypic analysis, it is best to use established ideograms and guidelines such as the one provided at http://amba.charite.de/cgh/protocol/02/class.html. It may also be appropriate or necessary to get advice from a cytogeneticist on karyotypic analysis. 8. This process is repeated for 5–10 metaphase spreads to ensure that at least 6–12 copies of each chromosome are used for interpretation. 9. A ratio profile is generated from each karyotype (by overlaying normalized red and green images) (see Note 14). A ratio of 1 indicates equal copy number, whereas ratios deviating from this indicate gains and losses of chromosomal material. Due to the genetic heterogeneity of tumor tissue cutoff values of 1.2 for gains are recommended.

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Fig. 13.4. Example of an average ratio profile. If the reference sample is labeled with Texas Red and the test sample is labeled with FITC, amplifications will be indicated by a green line to the right of each ideogram (shown in this figure as a light grey line, such as that seen in 14q) and deletions will appear as a red line to the left of each ideogram (shown in this figure as a dark grey line, such as that seen in 13q). For example, in this figure chromosomal location 8q is amplified and 1p is deleted.

10. An average ratio profile –1 standard deviation is generated from 5 to 10 metaphase spreads to indicate regions of chromosomal gain or loss. Figure 13.4 demonstrates a typical average ratio profile generated.

4. Notes 1. Cells can be stored at –20C in 10 ml of fixative. If cells are stored, the fixative must be replaced before preparing the slides (i.e., centrifugation, removal of supernatant, re-suspension in 5 ml fixative). Although 5 ml of fixative is recommended, the amount actually required may need to be adjusted if there are too few or too many (crowded) metaphase spreads on the slide.


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2. Metaphase chromosome spreads within the target areas need to be well separated. Viewing a target area with a DAPI stain will indicate if metaphase spreads are appropriate for CGH analysis. You will see many interphase cells (which appear as small circles) as well as metaphase spreads (appear as larger circles at 10 and individual chromosomes at 40 and 60 ). Chromosomes should be long enough for karyotyping and chromosomes that are medium to dark in color (phase-contrast microscopy, not fluorescence) may give better hybridization. As four different conditions are tried (for the four flasks), one of these is likely to produce quality metaphase spreads, however, the preparation may need to be repeated with different Colcemid incubation times (step 9). If the cells are too crowded within the target areas, then more fixative can be added to dilute the cells. If there are too few cells on the slide, then centrifuge the samples and resuspend in a smaller volume of fixative to concentrate the cells. If metaphase spreads are not spread out enough (i.e., too many chromosomes are overlapping) then try dropping the cells from a height (drop from 1–5 cm above the slide) or try adding a second drop of fixative on top of the drop of cells to spread them out further. Remember that you want at least 5–10 good metaphase spreads, this means that you may require up to 50 spreads or more within a target area. If suitable metaphase spreads cannot be established then use commercially available slides from Vysis (although these are not guaranteed, quality control usually means good metaphase spreads). 3. Banding patterns seen by the DAPI stain should be distinct so that the chromosomes can be easily karyotyped. Typically, chromosomes should have sharp boundaries and weak DAPI banding. 4. To make microdissection easier, it is best to put a small amount of PKTB on the scalpel blade to moisten the tissue. If the tissue is not moistened, it is likely to flake off and is difficult to recover. It is also best to remove the tissue surrounding the tumor first and then remove the isolated tumor sample. 5. Do not add more than 1 ml of microdissected DNA as it may inhibit the reaction (possibly due to products from the proteinase K reaction). If using fresh DNA for a positive control or reference sample, use 50 ng in a 1 ml reaction. During nick translation we typically use 40 ml of microdissected DOPPCRed DNA but only 25 ml of fresh genomic DOP-PCRed DNA. Thus, if using fresh DNA for reference DNA (recommended), a single DOP-PCRed sample is sufficient for two nick translation reactions.

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6. There should be no DNA smear present in the negative control. DOP-PCRed DNA obtained from fresh tissue is typically 200–2,000 bp in size, from microdissected tissue it is about 100–1,000 bp in size (at least some of the DNA should be greater than 300 bp in size). If the DNA fragments obtained from microdissected samples are less than 400 bp or are low in intensity, multiple (usually two to four) DOPPCRed samples can be pooled together and the DNA precipitated. Precipitation is performed by adding one-tenth the pooled sample volume of 3.0 M NaAc (pH 5.2) and 2.5 the volume of ice cold 100% ethanol. Samples are stored at –80C overnight and centrifuged the following day for 30 min at 11,000 rpm at 4C. The supernatant is removed and the pellet is washed with 70% ethanol and centrifuged for 15 min at 11,000 rpm at 4C. The supernatant is removed; the pellet air dried for about 10 min and then resuspended in 50 ml DNase/RNase-free water. Once again, 5 ml of the product is electrophoresed to check for quality. 7. The combination of enzymes used (Bionick enzyme mix, DNA polymerase I/DNase I enzyme mix and DNA polymerase) will need to be optimized for each new batch of enzymes. The volumes given here are an example and can be used as a good starting point. The size of the nick translated product for DNA extracted from fresh tissue should be 300–2,000 bp and from microdissected tissue 100–1,000 bp (with at least some of the product greater than 300 bp). This protocol uses a 75C incubation to halt the reaction, which allows additional nick translation if the resulting product is too large in size. An alternative approach is to halt the reaction by adding EDTA, although if this is used additional nick translation cannot be performed. If the product is too small in size, the ratio of DNase I and polymerase I can be adjusted (a decrease in the DNase I concentration should result in a larger product). Although nick translated products can be stored at –20C, better results often ensue when hybridization proceeds immediately after. Typically, test DNA is labeled with FITC-dUTP and reference DNA is labeled with Texas Red in the initial experiment, however reverse experiments also have to be performed as fluorophores often give misleading results at specific chromosomal regions. FITC is known to give inaccurate results at 1p32-pter, 16p, 17p, 19, 22, 7q21, 9q34, 16q and 17q, whereas 4q13-21, 11q21-23, 13q21qter, Xq21-q22 and 19p need to be interpreted with caution when using Texas Red for labeling. 8. If metaphase chromosomes are denatured for too short a time, the probe will not bind easily and signal intensity will be low. Also, if metaphase chromosomes are over-denatured


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they will appear distorted. If chromosomes do appear distorted either decrease the denaturation time (usually by 30 s – 1 min) or decrease the temperature of the solution to 72C. 9. It is important to avoid air bubbles when placing coverslips on the slides as the metaphase spreads will not be exposed to the probe. As a maximum number of metaphase spreads is required, the entire target area should be exposed to the probe and therefore air bubbles should be avoided. 10. During the first analysis if you notice that the DAPI counterstain is too strong or too weak you can adjust the concentration of DAPI in the DAPI/Vectashield mix. Although Vectashield protects the fluorophores (FITC, Texas Red and DAPI), they are light sensitive and it is best to proceed to digital analysis quickly. It is recommended to leave the slides at 4C in the dark overnight (or at least for 1 h) and then analyze the metaphase spreads or if this is not possible, analysis should occur within a few days. 11. Although the conditions under which the slides are made allow for an increased number of cells in metaphase, the number of interphase nuclei will heavily outweigh the number of metaphase spreads. Under the 10 objective metaphase spreads look larger and less distinct than interphase nuclei. Although the 10 magnification can give an idea of whether a metaphase spread is of good quality, the chromosomes will have to be viewed at 60 to ensure this. The metaphase spread must be intact (46 chromosomes present), chromosomes must be minimally overlapping, chromosome arms should be of appropriate length (not too long or short) and distinct and the metaphase spread should not be too close to other interphase nuclei or artifacts. Artifacts, such as dust, are often stained with the fluorophore and emit a very strong signal under the different excitation filters. It is therefore recommended that each metaphase spread is viewed briefly under each of the excitation filters to make sure that there is no artificial fluorescence that would interfere with analysis of the metaphase spread. Viewing the metaphase spread with the different excitation filters at this point will degrade the fluorophores slightly and should therefore not last more than a few seconds. It is very rare that a metaphase spread will have well-defined chromosomes that do not overlap at all, for this reason multiple metaphase spreads are analyzed to ensure good coverage of all chromosomes. 12. As various computer software packages are available for image capture and analysis, detailed instructions on the use of any one software package are not provided. Instructions will vary

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according to the software used and should be provided with the software itself. This section focuses on the types of images to be captured and the analysis that is required. When choosing a software package, ensure that it meets the requirements for image capture and CGH analysis. 13. These exposure times are suggestions only and will typically be different for each metaphase spread on a slide and will vary across experiments. Most software packages should allow each image (i.e., blue, green, red) to be captured independently so that different exposure times can be optimized for each of the filters without interfering with the previously captured image. To prevent degradation of the fluorophores, after each exposure the filter should be removed/blocked so that the slide is not exposed to light while viewing the captured image. 14. The software package may allow a color image to be overlaid on the karyotype before the generation of a ratio profile. This is an additional measure to check for correct karyotyping as gains and losses should be displayed on both chromosomes in the same position. If there is a discrepancy between homologous chromosomes, then karyotyping is likely incorrect or one of the chromosomes may be in close proximity of an artificial fluorescent spot.

References 1. Kallioniemi, A., Kallioniemi, O. P., Sudar, D., Rutovitz, D., Gray, J. W., Waldman, F., and Pinkel, D. (1992) Comparative genomic hybridization for molecular cytogenetic analysis of solid tumors. Science. 258, 818–21. 2. Kallioniemi, O. P., Kallioniemi, A., Piper, J., Isola, J., Waldman, F. M., Gray, J. W., and Pinkel, D. (1994) Optimizing comparative genomic hybridization for analysis of DNA sequence copy number changes in solid tumors. Genes Chromosomes Cancer. 10, 231–43. 3. Xu, J., and Chen, Z. (2003) Advances in molecular cytogenetics for the evaluation of mental retardation. Am J Med Genet C Semin Med Genet. 117, 15–24. 4. Pollack, J. R., Perou, C. M., Alizadeh, A. A., Eisen, M. B., Pergamenschikov, A., Williams, C. F., Jeffrey, S. S., Botstein, D., and Brown, P. O. (1999) Genome-wide analysis of DNA copy-number changes using cDNA microarrays. Nat Genet. 23, 41–6.

5. Houldsworth, J., and Chaganti, R. S. (1994) Comparative genomic hybridization: an overview. Am J Pathol. 145, 1253–60. 6. Gallo, J. H., Ordonez, J. V., Brown, G. E., and Testa, J. R. (1984) Synchronization of human leukemic cells: relevance for highresolution chromosome banding. Hum Genet. 66, 220–4. 7. Barch, M., Knutsen, T., and Spurbeck, J. (1997) The AGT Cytogenetics Laboratory Manual, Lippincott-Raven Publishers, Philadelphia, PA. 8. Miller, S. A., Dykes, D. D., and Polesky, H. F. (1988) A simple salting out procedure for extracting DNA from human nucleated cells. Nucleic Acids Res. 16, 1215. 9. Kuukasja¨rvi, T., Tanner, M., Pennanen, S., Karhu, R., Visakorpi, T., and Isola, J. (1997) Optimizing DOP-PCR for universal amplification of small DNA samples in comparative genomic hybridization. Genes Chromosomes Cancer. 18, 94–101.

Chapter 14 In Vitro Replication Assay with Mammalian Cell Extracts Wasia Rizwani and Srikumar P. Chellappan Abstract Regulatory mechanisms for DNA replication are crucial to the control of the cell cycle in eukaryotic cells. One of the widely used assays to understand the complex mammalian replication system is the cell-free in vitro replication assay (IVRA). IVRA can provide a snapshot of the regulatory mechanisms controlling replication in higher eukaryotes by using a single plasmid, pEPI-1. This chapter outlines the general strategies and protocols used to perform IVRA to study the differential recruitment of replication factors either independently or in combination, based on the experience in studying the role of prohibitin in replication as well as other published protocols. This method can be employed to identify not only proteins that assist replication but also proteins that inhibit replication of mammalian genome. Key words: Replication assay, pEPI-1 plasmid, nuclear and cytosolic extracts.

1. Introduction Genome replication in prokaryotes and some eukaryotic viruses such as simian virus 40 (SV40) initiates with the binding of specific initiator proteins to genomic sites, termed ‘‘origins’’ and results in the localized unwinding of the DNA duplex to form replication forks (1). SV40 expresses a single initiator protein, T antigen, which binds with high specificity to the viral origin, a specific 60 bp long DNA segment that functions as a helicase separating the complementary DNA strands (2). Contrary to this, the replication of cellular genomes is strictly coordinated within the cell cycles and efficiently controlled to complete the replication only once per cell cycle (3). This is accomplished by several levels of control starting with the step-wise assembly of pre-replication complexes (pre-RC), in the early G1 phase of the cell cycle, consisting of the DNA-bound multisubunit origin recognition Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_14 Springerprotocols.com

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complexes (ORC with its components Orc1–Orc6) that serves as a platform for the association of additional proteins such as Cdc6, Cdt1 and the minichromosome maintenance proteins (Mcm2–Mcm7) (4, 5). This is followed by the activation of specific protein kinases to begin the transition into S phase, which is marked by the conversion of pre-RC to initiation complexes. Partial degradation of the pre-RC, including the loss of Mcm proteins, prevents re-replication during the S phase (6–10). Dissociation of the Orc1p subunit of ORC could also protect mammalian cells from undergoing re-replication (11). This process so far was best understood in yeast, Saccharomyces cerevisiae. Presence of autonomously replicating sequences (ARSs) in yeast that direct the extrachromosomal replication of ARS-bearing plasmids made it possible to understand how specific eukaryotic replication is and the significance of defined origin elements (12, 13). Also in yeast, the replication origins are genetically determined which is not the case with higher eukaryotes. Therefore, it is believed that replication start sites in mammalian genomes are not totally determined by the underlying DNA sequences but by epigenetic factors such as the presence of specifically bound transcription factors or the conformation of the chromatin (14). This entitled for an in vitro assay system that is comparable to the ARS system in yeast to investigate the importance of origin specificity and replication proteins in mammalian cells. For this purpose, there was a need for a biochemically amenable plasmid that not only replicated episomally in cultured mammalian cells but also contained an authentic origin sequence to prevent rapid incorporation into the host genome or complete loss, and completely soluble proteins (15, 16). Two regions appeared to be important for extrachromosomal replication, an active CMV promoter with the adjacent downstream sequences and at least partial S/MAR coding region (17, 18). Hence, plasmid pEPI-1 possessing these two features was constructed by Piechaczek, C. et al. (19) to study in vitro replication (Fig. 14.1). The plasmid pEPI-1 of size 6.7 kb replicates autonomously in a once-per-cell-cycle manner and is stably transmitted over many generations (100), as observed in CHO cells (19) and HeLa cells (20) without selection. pEPI-1 has a unique combination of genetic elements including a strong eukaryotic promoter (CMV) and a scaffold/ matrix attachment region (S/MAR). It probably associates with mitotic chromosomes via an interaction of its S/MAR element with the scaffold attachment factor, SAF-A in cellular chromosomes (21, 22), and is therefore correctly transmitted from parental to progeny cells. To mimic in vivo replication system, a mixture of proteins with chain-elongating functions and replication initiation properties was necessary (5). For a long time, Xenopus egg extracts were used to study replication and were successful in investigations

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Fig. 14.1. A map of pEPI-1 plasmid. S/MAR represents the human scaffold/matrixattached region cloned into the multiple cloning site (MCS) of the vector pGFP-C1. The plasmid has SV40 origin, which is a T antigen target site to initiate in vitro replication. Adapted from Piechaczek et al. 1999 (19).

into the events leading to pre-RC formation (23–27) and to characterize important replication factors such as Cdt1 and Mcm8 (28, 29). However, the Xenopus egg extract contains high amounts of initiator proteins such as ORC and Mcm proteins, stored for subsequent rounds of cell divisions in early embryogenesis that differs from the situation in proliferating adult mammalian cells with their limited amounts of ORC. For this reason, the Xenopus extracts have been replaced with cytosolic mammalian cell extracts rich in replicative chain elongation factors and either T antigen or high salt nuclear extract for replication initiation (30, 31), thereby, providing an in vitro replication system that is sufficiently robust to investigate questions related to replication initiation in mammalian cells. This chapter will describe in detail the protocols to be followed for conducting an in vitro replication assay. It involves the preparation of cytosolic and nuclear extracts from mammalian cells and purification of the pEPI-1 plasmid followed by the assay. For the replication assay, as a positive control, T antigen may be used in place of nuclear extract to initiate replication. Dpn I digestion is an optional step to confirm the amount of replication quantitatively. Replication products are resistant to Dpn I digestion only when hemimethylated or unmethylated thereby indicating that DNA has undergone at least one round of replication in vitro (31). Hence, for the Dpn I digestion assay to work, the plasmid has to be purified either from the pEPI-1 transfected

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mammalian cells or from the pEPI-1 transfected methylationdeprived bacterial strains. This is because bacterial plasmids undergo methylation on both the strands and are susceptible to Dpn I digestion. This assay is extensively employed to examine the formation of functional pre-replication complexes under biochemical conditions.

2. Materials 1. 30% acrylamide solution (0.8% bis): 60 g acrylamide and 1.6 g bis. Heat to dissolve in approximately 100 ml of water. QS to 200 ml with water and filter with Whatman #1 filter paper into a foil-wrapped bottle. 2. 40% acrylamide solution (19:1 acrylamide:bis): 80 g acrylamide and 4.21 g bis. Heat to dissolve in approximately 100 ml of water. QS to 200 ml with water and filter with Whatman #1 filter paper into a foil-wrapped bottle. 3. 50% acrylamide solution (19:1 acrylamide:bis): 237.5 g (97%) acrylamide and 12.5 g bis. Dissolve in 250 ml hot water. Filter to remove debris and wrap the bottle in foil. 4. Buffer A: 20 mM HEPES pH 7.4, 5 mM KCl, 1.5 mM MgCl2, 0.1 mM dithiothreitol. Stored at room temperature. 5. Buffer A containing 250 mM sucrose. Stored at room temperature. 6. Buffer A containing 450 mM potassium acetate. Stored at room temperature. 7. Buffer NE1: 10 mM HEPES pH 8.0, 1.5 mM MgCl2, 10 mM KCl, 1 mM DTT and 1 complete, EDTA-free protease inhibitor. 8. Buffer NE2: 20 mM HEPES pH 8.0, 1.5 mM MgCl2, 25% glycerol, 420 mM NaCl, 1 mM DTT, 0.5 mM PMSF and 1 complete, EDTA-free protease inhibitor. 9. Creatine kinase BB fraction, human. (Sigma, cat # C9983100UG). Dissolve in 1 ml water. Store at –20C in aliquots. 10. Creatine phosphate disodium salt tetrahydrate. (Sigma, cat # 27920). Dissolve 163 mg in 1 ml water to get a stock of 500 mM concentration. Store in aliquots at –20C. 11. Deoxynucleotide phosphates (100 mM stock solutions from Promega): dATP (cat # U120-B ), dCTP (cat # U121-B), dGTP (cat # U122-B), dTTP (cat # U123-B).


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12. Dpn I restriction enzyme. (Promega, cat # R0176S). 13. 50% glycerol: 25 ml 100% glycerol and 25 ml water. Autoclave. 14. NEB4 buffer (Promega, cat # B7004S, 10 concentrate). 15. Nucleotide phosphates (100 mM stock solutions from Promega)-ATP (cat # E601B), UTP (cat # E602B), GTP (c at # E603B), CTP (cat # E604B). 16. Phosphate-buffered saline, pH 6.5 (20 ): 160 g NaCl, 4 g KCl, 28.8 g Na2HPO4, 4.8 g KH2PO4 in 1 l of water. It can be stored at room temperature. For 1 PBS, 50 ml of 20 PBS is made up to 1 l, pH adjusts to 7.2 upon dilution with water. The buffer has to be ice-cold when used. 17. Protease inhibitor (complete, EDTA-free from Roche Applied Sciences, cat # 04693132001). One tablet is dissolved in 2 ml of water and aliquoted into 200 ml and stored at –20C. 18. Proteinase K (Promega, cat # V302A): dissolve 10 mg powder in 1 ml of TE buffer and store at –20C in aliquots. Thaw on ice before use. 19. RNase A (Sigma, cat # R-5503): dissolve 10 mg in 1 ml of 1 TE, pH 7.5. Boil for 30 min. 20. 3 M sodium acetate, pH 5.2 21. Stop mix: 60 mM EDTA, 2% SDS. 22. 50 TAE (1 M Tris–HCl, 57.1 ml acetic acid, 50 mM EDTA pH 8.3): 242.2 g Tris, 57.1 ml acetic acid, 18.6 g EDTA. Adjust pH to 8.3. QS to 1 l. Autoclave (optional, refer to Note 1). 23. 10 TBE (1 M Tris–HCl, 1 M boric acid, 20 mM EDTA pH 8.3): 242.2 g Tris, 123.66 g boric acid, 14.89 g EDTA. Adjust pH to 8.3. QS to 2 l. Autoclave (refer to Note 1). (Biotechniques 10:182, 1991 claims that filtering up to a 20 TBE solution through 0.2–0.45 mM cellulose acetate or cellulose nitrate filters prevents formation of precipitants during long-term storage. The solution may be re-autoclaved to dissolve precipitates that form). 24. 1 TE buffer, pH 7.5: 10 mm Tris–HCl and 1 mM EDTA. 25. S.O.C media (Invitrogen, cat # 15544-034). Stored at room temperature. 26. Kanamycin –30 mg/ml in water. Store at –20C. 27. [–32P]dATP-10 mCi/ml. 28. Water – molecular biology grade from Sigma. 29. QIAprep Spin Miniprep Kit – (Qiagen, cat # 2714). 30. Recipe for electrophoresis of DNA in polyacrylamide gels.

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Gel sizes 75 100 mm

Small: Medium:

100 150 mm

Large:

160 180 mm

5% analytical gels

1 mm Small

1 mm Medium

1 mm Large

10 TBE buffer (ml)

1.0

1.75

2.5

40% bis/acrylamide (ml)

1.25

2.2

3.1

Water (ml)

5.75

10.05

50% glycerol (ml)

2.0

3.5

5.0

11.0

20.0

28.5

7.5

10.7

Reagent

APS (mg) TEMED (ml) Total volume (ml)

4.27 10

17.5

14.4

25

With other concentrations of a bis/acrylamide stock substitute:

Reagent

1 mm Small

1 mm Medium


2.0

3.5

Water (ml)

5.0

9.25


1.67

2.9

Water (ml)

5.33

9.25

13.35


1.0

1.75

2.5

Water (ml)

6.0

10.5

1 mm Large 5.0 12.5 4.15

15.0


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3. Methods 3.1. Cell Culture and Preparation of Cytosolic and Nuclear Extracts 3.1.1. Protocol 1

The extracts can be isolated by two different methods as described below: 1. 293T cells (contain endogenously expressing T antigen) or breast cancer MCF7 cells are cultured in DMEM containing 1% antibiotic–antimycotic solution and 10% fetal bovine serum until 80–90% confluency in a 100 mm culture dish. One plate should be sufficient to make cell extracts for a few assays. 2. Wash the plates with cells twice with 10 ml of ice-cold 1 PBS, collect them in eppendorf with 1 ml of buffer A containing sucrose and wash twice with buffer A (see Note 3). 3. Keep the cells on ice for 10 min in 100–200 ml of buffer A (depending on the cell pellet size, preferable to keep the buffer volume small to obtain concentrated protein) and disrupt in a type S dounce homogenizer. 4. Pellet the nuclei by centrifugation at 16,000 g for 10 min, which are processed further as described later. 5. The supernatant is then cleared by centrifugation at 100,000 g for 1 h to prepare the S100 extract. Approximately 7–10 mg protein/ml is obtained. The protein is stored at –70C in aliquots. 6. Wash the nuclear pellet once in buffer A and incubate in approximately 100 ml of buffer A containing potassium acetate for 90 min on ice. 7. After preclearing at 12,000 g for 5 min, the supernatant is centrifuged at 300,000 g for 1 h and collected as S300 extracts (2–3 mg protein /ml). Store at –70C in aliquots until further use.

3.1.2. Protocol 2

1. Cells should be 80% confluent for extraction. 10 cm 10 cm dishes of cells will yield around 50 ml of nuclear extract (see Note 4). 2. Wash the plates with ice-cold PBS twice, scrape and collect cells in 1 ml PBS per plate. Place in eppendorf and spin at 800 g for 2 min at 4C. Resuspend all pellets into 1 ml of PBS and spin again under same conditions (see Note 3). 3. Aspirate PBS completely and estimate packed cell volume (PCV). Resuspend in 1 PCV of NE1 buffer. Leave on ice for 15 min. 4. Take up the suspension into a 1 ml syringe that has been washed with buffer NE1. In one stoke, squirt the cell suspension through a 23-gauge needle into an eppendorf. Repeat this step four more times. (Place syringes in freezer before use.)

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5. Microfuge at high speed (20,000 g) at 4C for 20 s to collect the supernatant, which is the cytosolic fraction. 6. Resuspend the nuclear pellet in two-third PCV of NE2 buffer. Leave on ice with regular stirring for 30 min. 7. Pellet the nuclear debris at high speed for 5 min in cold microfuge. 8. Collect the supernatant, which is the nuclear fraction.

3.2. Preparation of pEPI-1 Plasmid from Transfected Mammalian Cells

3.2.1. By Hirt Extraction Protocol (32)

1. Culture pEPI-1 transfected mammalian cells in 100 mm dishes (106–107 cells/plate). Wash the plate with ice-cold PBS twice, scrape the cells in 1 ml PBS and collect the pellet in an eppendorf tube by centrifuging for 2 min at 800 g at 4C. 2. Resuspend the pellet in 0.5 ml of 1 TE buffer, and then add another 0.5 ml of TE containing 2% SDS. Gently invert the tube several times to mix. Pipetting is to be avoided to prevent shearing of the DNA. 3. The volume will now be approximately 1.1 ml. Hold at room temperature for 10 min. 4. Add 5 M NaCl such that the final NaCl concentration will come to 1 M (i.e., around 220 ml of 5 M NaCl). Slowly invert the tube several times to mix and keep at 4C for 8 h to overnight. 5. Microfuge at 17,000 g for 1 h at 4C and transfer the supernatant to a separate 2 ml eppendorf tube carefully avoiding any remnants of the pellet to appear in the supernatant. Squirt 1 ml of 10 mM EDTA pH 8.0 containing 0.6% SDS. 6. Carefully transfer it to a new eppendorf which now contains the plasmid along with RNA and subject to proteinase K and RNase A treatment. The pellet contains genomic DNA and can be processed similarly in case it needs to be assessed. 7. For this, add 10 ml of 10 mg/ml proteinase K (final concentration will be 100 mg/ml) and 1 ml of 10 mg/ml RNase A. Leave at 37C for 1.5 h to digest the pellet. 8. Add equal volume of phenol–chloroform and extract the plasmid. Repeat this step once followed by extraction with chloroform or isopropanol alone. 9. Transfer the aqueous phase to a fresh eppendorf tube(s) and add two volumes of 100% ethanol and leave at –20C for 1 h to overnight or at –80C for 30 min. 10. Spin at high speed for 30 min at 4C in a microfuge. Decant the supernatant and air dry the pellet (mark the region where pellet is visible before drying because it becomes invisible after drying).


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11. Re-dissolve the pellet in 50 ml sterile water (see Note 2). 12. Plasmid can be checked by running on 0.8% agarose gel or analyzed by Dpn I digestion (see Section 3.3.2).

3.2.2. By Using QIAprep Spin Miniprep Kit

This is a modified protocol (33) basically designed for the purification of high-copy plasmid DNA from bacterial cultures (http://www.qiagen.com/literature/plklit.asphashplasmidmini). It is now being used to purify low-molecular-weight DNA such as SV40 containing plasmids from transfected mammalian cells (see Notes 6, 7 and 8). 1. Culture pEPI-1 transfected mammalian cells in 60 mm dish. Rinse the plate once with ice-cold PBS. 2. Add 250 ml each of buffers P1 and P2 directly to the plate and incubate for 5 min at room temperature. 3. Gently transfer the cell lysate to a microfuge tube using a rubber policeman and digest with proteinase K at a final concentration of 800 mg/ml at 55C for 1 h. 4. Add 350 ml of buffer N3 to precipitate cellular DNA and cell debris by mixing gently, incubate on ice for 5 min and spin at 16,000 g for 10 min. 5. Load the supernatant onto the spin column and centrifuge for a minute at 16,000 g. Wash the spin column once each with buffers PB and PE. Give an additional 1 min spin to remove residual wash buffer and dry the column for 5 min to evaporate residual ethanol, which may inhibit subsequent enzymatic reactions. 6. Elute the plasmid with 50 ml of water (see Note 2) by adding it to the center of the column, incubate at 37C for 5 min and centrifuge at 16,000 g for 1 min. Repeat this step, if desirable.

3.2.3. By Transformation into DAM or DCM Minus Bacterial Strains Followed by Purification Using QIAprep Spin Miniprep Kit

1. Thaw 50 ml DAM– or DCM– competent cells on ice in an eppendorf, add 1 ml plasmid and mix gently with a pipette. 2. Incubate on ice for 30 min. 3. Give heat shock by holding the tube in 37C water bath for 20 s. 4. Keep on ice for 2 min, and then add 950 ml S.O.C. media inside a sterile biosafety cabinet. 5. Allow the cells to grow for an hour at 37C with shaking conditions of 225 rpm. 6. Spread 50 ml culture on LB plate containing 30 mg/ml Kanamycin and incubate at 37C overnight. 7. The plate with bacterial colonies can be stored at 4C. Glycerol stocks can be made for long-term storage.

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8. Inoculate a single colony into 3 ml LB broth containing 30 mg/ml Kanamycin and incubate at 37C overnight rotating at 250 rpm in a bacterial shaker. The culture is pelleted at 800 g for 5 min and subjected to plasmid purification using QIAprep spin miniprep kit.

3.3. In Vitro Replication Assay

1. In a total volume of 35 ml with S300 nuclear extract (16–32 mg, preferably more nuclear extract to increase the probability of pre-RC formation), add pEPI-1 DNA (160–320 ng) at 2 mM ATP and 80 mM potassium acetate in buffer A plus 1 ml of Complete, EDTA-free protease inhibitor (see Note 2). Pre-incubate this mix at 37C for 60 min. 2. Initiate the replication by the addition of the S100 extract (45 mg), 40 mM creatine phosphate; 0.6 mg/ml of creatine kinase; 30 mM potassium acetate; 80 mMCTP, GTP and UTP; 100 mM dGTP, dCTP and dTTP; 30 mM dATP; 0.5 mg – T antigen and 10 mCi of [–32P] dATP in a total volume of 50 ml (see Note 5). 3. Incubate at 37C for 60 min and then stop the reaction by 30 ml of stop mix. 4. Extract the DNA by proteinase K digestion and phenol– chloroform treatment, followed by ethanol precipitation as described below (Section 3.3.1).

3.3.1. DNA Extraction Following Replication Assay

1. After the replication assay, adjust the reaction volume to 200 ml with water (molecular biology grade). 2. To this, add 10% of 3 M sodium acetate pH 5.2 and 1 ml ethanol and vortex for a few seconds to mix. Allow the DNA to precipitate by incubating at –80C for 15 min, centrifuge at high speed (20,000 g) for 15 min at 4C. 3. Decant the solvent and air dry the pellet. An optional 70% ethanol wash can also be done before air drying the DNA pellet. 4. Re-dissolve the pellet in 200 ml of water and perform proteinase K digestion (50 mg) at 37C for 30 min. Following which, phenol–chloroform extraction is done by adding 200 ml of buffered phenol, mix by vortexing for a minute and centrifuge at high speed for 10 min at room temperature. Collect the aqueous phase and repeat steps 2 and 3. 5. Resuspend the pellet in 20 ml of sterile water.

3.3.2. Dpn I Digestion

1. Reaction mix (total volume 20 ml) contains plasmid obtained from step 3.3.1 (10 ml), NEB 4 (2 ml), Dpn I (2 ml), 10 BSA (2 ml) and water (4 ml). Incubate the mix for 4 h to overnight at 37C and follow with phenol:chloroform extraction and ethanol precipitation.


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2. Re-dissolve the DNA pellet in 20 ml of water. Run it either on 0.8% agarose at 80–100 V or 5% polyacrylamide gel at 200 V. Ethidium bromide staining will help in visualizing the bands. If testing for replication, dry the gel for 1 h and expose to autoradiography anywhere from 2 days to a week. 3. If running an agarose gel, before drying, it is preferable to wash the gel in 7% TCA in water for 30 min with gentle rocking (see Note 1).

4. Notes 1. It is advisable to autoclave buffers used in making the plasmid, carrying out IVRA or running gel to prevent plasmid degradation by nucleases. 2. A very important point to remember is to use EDTA-free solutions and water in order to prevent unwanted termination of IVRA because EDTA is used to stop the replication assay. 3. It is essential to perform all steps of cell lysis on ice or at 4C to prevent unwanted degradation of proteins. Likewise, addition of protease inhibitors wherever mentioned will preserve the integrity of proteins. 4. Precautions during nuclear and cytosolic extract preparation. Make sure the homogenization is proper in order to eliminate cytosolic protein contamination in the nuclear fraction. Usually the nuclear extract obtained is dialyzed before use, but not in this assay because it needs a high salt nuclear extract that contains many proteins known or believed to be involved in mammalian genome replication such as Mcm’s, Orc’s, Cdt1 or Cdc6 among others. 5. Under similar biochemical conditions, it was found that replication of plasmid was only 2–3% when cytosolic and nuclear extracts were used from MCF7 cells, much better for 293T cell extracts and best achieved by large T antigen. This is quite plausible because the initiator T antigen is added in high amounts to the incubation mixtures and does not need to interact with other proteins for the recognition and the unwinding of the viral origin. In contrast, the dozen or so cellular initiator functions are inevitably diluted when extracted from the nuclei of cultured cells and assemble on one of many possible sites on the plasmid template where they must interact in a highly complex manner before they are ready to initiate a replication round. Hence, as a positive control large T antigen could be used since pEPI-1 plasmid has SV40 origin to assess replication in in vitro conditions. T antigen can be purchased from the

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company, ChimerX, USA (cat # 5800–01). Another reason why replication efficiency is low when nuclear extracts from cells are used, instead of T antigen, could be the presence of limiting factors such as proteins that negatively control replication in the nuclear extracts. Immunodepletion or silencing of potential replication-inhibiting proteins could help in accelerating the replication efficiency. Reversibly, the effect of these proteins can be seen by addition of excessive amounts of inhibiting proteins. 6. It is preferable to extract pEPI-1 plasmid directly from transfected mammalian cells by Hirt extraction protocol or to amplify the plasmid in bacterial strains – DAM– or DCM– strains, which prevents methylation of plasmids. The reason for observing these preventative measures is parental pEPI-1 plasmid obtained from bacteria is potentially methylated on both strands and hence susceptible to Dpn I digestion. 7. Hirt extraction of plasmid from cells could be a time-consuming and labor-intensive procedure. Qiagen has a kit available to isolate these plasmids called as QIAprep Spin Miniprep Kit with some modifications as described in the methods, Section 3.2.2. The advantage of this kit over Hirt extraction is phenol–chloroform step can be omitted and experimental inconsistencies such as pipetting errors can be avoided. 8. The spin kit uses silica gel membrane for DNA binding, hence care should be taken to prevent overloading of the sample on the columns to obtain high DNA yield of good quality.

Acknowledgments Work in the Chellappan lab is supported by the grants CA63136, CA77301 and CA127725 from the NIH. We wish to thank Mark Alexandrow, Moffitt Cancer Center, for providing the pEPI-1 plasmid and for helpful discussions.

References 1. Dean, F. B., Bullock, P., Murakami, Y., Wobbe, C. R., Weissbach, L. & Hurwitz, J. (1987) Simian virus 40 (SV40) DNA replication: SV40 large T antigen unwinds DNA containing the SV40 origin of replication, Proc Natl Acad Sci U S A. 84, 16–20. 2. Simmons, D. T. (2000) SV40 large T antigen functions in DNA replication and transformation, Adv Virus Res. 55, 75–134.

3. Coverley, D. & Laskey, R. A. (1994) Regulation of eukaryotic DNA replication, Annu Rev Biochem. 63, 745–76. 4. Bell, S. P. & Dutta, A. (2002) DNA replication in eukaryotic cells, Annu Rev Biochem. 71, 333–74. 5. Blow, J. J. & Hodgson, B. (2002) Replication licensing – defining the proliferative state?, Trends Cell Biol. 12, 72–8.


6. Alexandrow, M. G., Ritzi, M., Pemov, A. & Hamlin, J. L. (2002) A potential role for mini-chromosome maintenance (MCM) proteins in initiation at the dihydrofolate reductase replication origin, J Biol Chem. 277, 2702–8. 7. Aparicio, O. M., Weinstein, D. M. & Bell, S. P. (1997) Components and dynamics of DNA replication complexes in S. cerevisiae: redistribution of MCM proteins and Cdc45p during S phase, Cell. 91, 59–69. 8. Labib, K., Tercero, J. A. & Diffley, J. F. (2000) Uninterrupted MCM2-7 function required for DNA replication fork progression, Science. 288, 1643–7. 9. Schaarschmidt, D., Ladenburger, E. M., Keller, C. & Knippers, R. (2002) Human Mcm proteins at a replication origin during the G1 to S phase transition, Nucleic Acids Res. 30, 4176–85. 10. You, Z., Komamura, Y. & Ishimi, Y. (1999) Biochemical analysis of the intrinsic Mcm4Mcm6-mcm7 DNA helicase activity, Mol Cell Biol. 19, 8003–15. 11. DePamphilis, M. L. (1999) Replication origins in metazoan chromosomes: fact or fiction? Bioessays. 21, 5–16. 12. Bell, S. P. & Stillman, B. (1992) ATP-dependent recognition of eukaryotic origins of DNA replication by a multiprotein complex, Nature. 357, 128–34. 13. Lee, D. G. & Bell, S. P. (1997) Architecture of the yeast origin recognition complex bound to origins of DNA replication, Mol Cell Biol. 17, 7159–68. 14. Mechali, M. (2001) DNA replication origins: from sequence specificity to epigenetics, Nat Rev Genet. 2, 640–5. 15. McWhinney, C. & Leffak, M. (1990) Autonomous replication of a DNA fragment containing the chromosomal replication origin of the human c-myc gene, Nucleic Acids Res. 18, 1233–42. 16. Price, G. B., Allarakhia, M., Cossons, N., Nielsen, T., Diaz-Perez, M., Friedlander, P., Tao, L. & Zannis-Hadjopoulos, M. (2003) Identification of a cis-element that determines autonomous DNA replication in eukaryotic cells, J Biol Chem. 278, 19649–59. 17. Bode, J., Kohwi, Y., Dickinson, L., Joh, T., Klehr, D., Mielke, C. & Kohwi-Shigematsu, T. (1992) Biological significance of unwinding capability of nuclear matrix-associating DNAs, Science. 255, 195–7.

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18. Stehle, I. M., Scinteie, M. F., Baiker, A., Jenke, A. C. & Lipps, H. J. (2003) Exploiting a minimal system to study the epigenetic control of DNA replication: the interplay between transcription and replication, Chromosome Res. 11, 413–21. 19. Piechaczek, C., Fetzer, C., Baiker, A., Bode, J. & Lipps, H. J. (1999) A vector based on the SV40 origin of replication and chromosomal S/MARs replicates episomally in CHO cells, Nucleic Acids Res. 27, 426–8. 20. Schaarschmidt, D., Baltin, J., Stehle, I. M., Lipps, H. J. & Knippers, R. (2004) An episomal mammalian replicon: sequence-independent binding of the origin recognition complex, Embo J. 23, 191–201. 21. Baiker, A., Maercker, C., Piechaczek, C., Schmidt, S. B., Bode, J., Benham, C. & Lipps, H. J. (2000) Mitotic stability of an episomal vector containing a human scaffold/matrix-attached region is provided by association with nuclear matrix, Nat Cell Biol. 2, 182–4. 22. Jenke, B. H., Fetzer, C. P., Stehle, I. M., Jonsson, F., Fackelmayer, F. O., Conradt, H., Bode, J. & Lipps, H. J. (2002) An episomally replicating vector binds to the nuclear matrix protein SAF-A in vivo, EMBO Rep. 3, 349–54. 23. Gilbert, D. M., Miyazawa, H. & DePamphilis, M. L. (1995) Site-specific initiation of DNA replication in Xenopus egg extract requires nuclear structure, Mol Cell Biol. 15, 2942–54. 24. Hyrien, O. & Mechali, M. (1992) Plasmid replication in Xenopus eggs and egg extracts: a 2D gel electrophoretic analysis, Nucleic Acids Res. 20, 1463–9. 25. Mahbubani, H. M., Paull, T., Elder, J. K. & Blow, J. J. (1992) DNA replication initiates at multiple sites on plasmid DNA in Xenopus egg extracts, Nucleic Acids Res. 20, 1457–62. 26. Newport, J. (1987) Nuclear reconstitution in vitro: stages of assembly around proteinfree DNA, Cell. 48, 205–17. 27. Newport, J. & Spann, T. (1987) Disassembly of the nucleus in mitotic extracts: membrane vesicularization, lamin disassembly, and chromosome condensation are independent processes, Cell. 48, 219–30. 28. Maiorano, D., Cuvier, O., Danis, E. & Mechali, M. (2005) MCM8 is an MCM27-related protein that functions as a DNA helicase during replication elongation and not initiation, Cell. 120, 315–28.

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29. Maiorano, D., Moreau, J. & Mechali, M. (2000) XCDT1 is required for the assembly of pre-replicative complexes in Xenopus laevis, Nature. 404, 622–5. 30. Alexiadis, V., Halmer, L. & Gruss, C. (1997) Influence of core histone acetylation on SV40 minichromosome replication in vitro, Chromosoma. 105, 324–31. 31. Baltin, J., Leist, S., Odronitz, F., Wollscheid, H. P., Baack, M., Kapitza, T., Schaarschmidt, D. & Knippers, R.

(2006) 32. DNA replication in protein extracts from human cells requires ORC and Mcm proteins, J Biol Chem. 281, 12428–35. 32. Hirt, B. (1967) Selective extraction of polyoma DNA from infected mouse cell cultures, J Mol Biol. 26, 365–9. 33. Ziegler, K., Bui, T., Frisque, R. J., Grandinetti, A. & Nerurkar, V. R. (2004) A rapid in vitro polyomavirus DNA replication assay, J Virol Methods. 122, 123–7.

Chapter 15 Investigation of Genomic Methylation Status Using Methylation-Specific and Bisulfite Sequencing Polymerase Chain Reaction Melanie Carless Abstract Epigenetic modification plays a central role in the regulation of gene expression and therefore in the development of disease states. In particular, genomic methylation of cytosines within CpG dinucleotides is crucial to development, gene silencing and chromosome inactivation. Importantly, aberrant methylation profiles of various genes are associated with cancer and potentially autoimmune disease, brain-related disease, diabetes and heart disease. Various methods are available for the detection and quantification of methylation in a given sample. Most of these methods rely upon bisulfite conversion of DNA, which converts unmethylated cytosines to uracil, while methylated cytosines remain as cytosines. Methylationspecific amplification of DNA can be used to detect methylation at one or more (typically up to about 4) CpG sites by using primers specific to either methylated or unmethylated DNA. Alternatively, amplification of both methylated and unmethylated DNA followed by sequencing can be used to detect methylation status at multiple CpG sites. The following chapter provides protocols for bisulfite conversion of DNA, methylation-specific PCR and bisulfite sequencing PCR. Key words: Methylation, bisulfite, PCR, MSP, BSP.

1. Introduction Various mechanisms exist by which gene expression is regulated, including both genetic and epigenetic modifications. In recent years it has become evident that epigenetic modifications are essential for regular cellular function as well as for the development of various disease states, of which the most frequently studied is cancer. One such epigenetic mechanism is genomic methylation, which occurs at the C(5) position of cytosine in Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_15 Springerprotocols.com

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cytosine–guanine dinucleotides (CpG) and is regulated by DNA methyltransferases (1–3). In normal cells, CpG dinucleotides are generally methylated throughout the genome, except for in regions known as CpG ‘‘islands’’, which are hypomethylated (3). CpG islands are regions of about 0.5–2.5 kb in size that are rich in CpG dinucleotides (1). These islands are usually located in promoter regions or 50 exonic regions of genes and are present in about half of all human genes (1, 3). Methylation of cytosine residues within the DNA affects the binding of transcription factors to that DNA sequence (2). Thus, when CpG islands within the promoter or 50 exonic regions become methylated, transcription can be inhibited and gene expression is silenced. As such, DNA methylation plays a critical role in allele-specific expression, X chromosome inactivation, genomic stability and embryonic development (2). In addition to this, silencing of genes critical to normal cell growth and development can have detrimental consequences, leading to the development of various diseases. Aberrant methylation plays an important role in the development of cancer (1, 4) and potentially in autoimmune diseases (lupus, multiple sclerosis) (4), brainrelated disorders (bipolar disorder and schizophrenia) (5), atherosclerosis (6) and diabetes (7). Hypermethylation of CpG islands in specific genes is usually accompanied by an overall decrease in methylation of the genome, which is believed to lead to chromosomal instability and increased mutation events (4). The potential importance of genomic methylation in a number of diseases calls for techniques to assess gene methylation status. Early methylation detection techniques included employment of methylation-sensitive restriction enzymes (qualitative) and hydrazine or potassium permanganate DNA modification for genomic sequencing (8). The introduction of a method that uses sodium bisulfite-mediated conversion of cytosine (not 5methylcytosine) to uracil (9) has heralded a breakthrough in genomic methylation analysis. This allows methylated and unmethylated DNA to be differentiated as only methylated cytosines will remain as cytosines after conversion. Since then, various assays for the assessment of methylation status within specific regions have been developed, including methylationspecific PCR (MSP), methylation-sensitive single-nucleotide primer extension (MS-SNuPE), combined bisulfite restriction analysis (COBRA), bisulfite sequencing PCR (BSP), spectrometry-related assays, real-time quantitative-based assays, enzymatic regional methylation assay (ERMA) and oligonucleotide-based microarrays; for a review of these methods see (10). Global methylation techniques such as high-performance liquid chromatography (HPLC), restriction landmark genomic scanning (RLGS), differential methylation hybridization (DMH) and bacterial artificial chromosome (BAC) arrays can be employed to

Detection of Genomic Methylation

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examine the amount/ratio of methylated DNA in the whole genome or regions of the genome, such as repetitive regions or CpG islands (10). This chapter focuses on methylation-specific PCR and bisulfite sequencing PCR to investigate the methylation status of specific loci. Both of these techniques use bisulfite-modified DNA to interrogate one or more CpG sites within a CpG island. Briefly, MSP uses two independent primer sets that span one or more CpG sites that are specific for either methylated or unmethylated cytosine. BSP uses a single primer set that flanks a region containing a CpG island, so that all cytosines within the island can be interrogated for methylation status by sequencing. Following is some commonly used methods for the assessment of methylation in CpG sites.

2. Materials 2.1. Sodium Bisulphite Conversion

1. Microfuge tubes (1.5 ml or similar, need to be amber or kept in dark) 2. Pipette tips (2, 20, 200 ml, 1 ml or similar) 3. 1 mg (or less) DNA 4. Optional: salmon sperm DNA (Sigma), this is only required if you have less than 1 mg DNA 5. 1.0 M NaOH 6. 10 mM hydroquinone (Sigma): this must be made fresh the day of the experiment; light sensitive; toxic (hazard class 6.1) 7. 3.0 M sodium bisulfite, pH 5.0 (Sigma): this must be made fresh the day of the experiment, adjust pH with NaOH 8. Mineral oil (autoclaved, filtered into screw cap tubes and reautoclaved) 9. Heatblock or water bath (set to 50C, must fit microfuge tubes) 10. Vacuum manifold (a Luer-Lok system can be used instead but the vacuum gives higher reproducibility and DNA quality) 11. Wizard DNA Clean-up system (Promega, A7280): store at room temperature (RT) resin should be protected from light 12. 80% isopropanol 13. 3 M NaOH 14. 20 mg/ml glycogen (Roche)

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15. 10 M ammonium acetate (NH4Ac) 16. Ethanol (100 and 70% in DNase/RNase-free water) 2.2. PCR

1. 10 bisulfite PCR buffer (alternatively use buffer provided with Taq polymerase): 1.66 ml 1 M ammonium sulfate (166 mM final) (Sigma); 6.7 ml 1 M Tris–HCl, pH 8.8 (670 mM final); 670 ml 1 M MgCl2 (67 mM final); 69.9 ml 14.3 M b-mercaptoethanol (100 mM final) (Sigma: toxic, class 6.1, keep tightly sealed); 900.1 ml DNase/RNase-free water 2. 25 mM dNTPs (Promega or other; store at –20C) 3. 10 mM forward and reverse primers MSP: two primer sets specific to methylated and unmethylated DNA BSP: one primer set that does not discriminate methylated from unmethylated DNA 4. DNase/RNase-free water 5. Taq polymerase (Sigma, Invitrogen, Qiagen) 6. Thermocycler 7. PCR tubes or 96 well plate

2.3. Methods of Detection

2.3.1. MethylationSpecific PCR

1. Analytical grade agarose (Promega, Sigma) 2. 1 TAE: add 975 ml MilliQ water to 25 ml 40 TAE (Promega) (40 TAE: 1.6 M Tris–acetate, 40 mM EDTA) 3. Conical flask 4. 10 mg/ml ethidium bromide solution (Promega, Sigma; mutagenic, hazard class 6.1, store in dark) 5. 100% glycerol solution or 6 gel loading dye (if it does not interfere with band size) 6. 100 bp molecular weight marker (NEB, Qiagen, Promega) 7. Gel tray and combs (7 cm 10 cm tray and 2 11 teeth combs are sufficient for the PCR described; wide teeth are required for the combs to hold up to 60 ml sample) 8. Electrophoresis tank and power source 9. UV light box and camera (ensure glasses/face shield is worn when using UV light box)

2.3.2. Bisulfite Sequencing PCR

1. 1% (or 2%) agarose gel 2. UV light box (ensure glasses/face shield is worn when using UV light box) 3. Scalpel/razor for cutting 4. Gel extraction/purification kit (Qiagen, Invitrogen, Promega, Millipore)


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5. BigDye1 Terminator Ready Reaction Mix v3.1, includes 5 sequencing buffer (Applied Biosystems; store at –20C in the dark, avoid freeze/thaw more than 5–10 times, keep thawed reagents on ice) 6. 10 mM forward and reverse primers (same as those used in Section 2.2) 7. DNase/RNase-free water 8. 125 mM EDTA (Sigma) 9. Ethanol: 100 and 70% (in DNase/RNase-free water) 10. Hi-DiTM formamide (Applied Biosystems; store at –20C) 11. 96 well plate 12. Polymer (Pop-7TM for 3730, others available; Applied Biosystems; store at 4C) 13. 10 running buffer (Applied Biosystems; store at 4C) 14. 3730 (or 310, 370, 373, 377, 3100, 3130) genetic analyzer and required accessories (capillary, plate tray, septa seal) (Applied Biosystems)

3. Methods 3.1. Sodium Bisulfite Conversion

Sodium bisulfite converts unmethylated cytosines into uracil so that following PCR amplification all unmethylated cytosines within a sequence will be replaced with a thymine. Methylated cytosines will remain as cytosines following PCR amplification. Various kits are available for bisulfite conversion and are often used as some shorten the procedure time (as little as 3–5 h). These kits are available from companies such as Qiagen, Invitrogen and Sigma-Aldrich. Applied Biosystems, Zymo Research, Chemicon and Active Motif provide kits which require longer preparation times (10–16 h). If a kit is being used for bisulfite conversion, the manufacturers’ instructions should be followed. Alternatively, a standard protocol can be used for bisulfite conversion of methylated DNA, this takes about 17 h. The following protocol is a modification of that provided by Frommer and colleagues (9). 1. Denature 1–2 mg DNA (in 40 ml DNase/RNase-free water) with 10 ml 1.0 M NaOH (final concentration 0.2 M) at 37C for 10–30 min (see Note 1). 2. Add 30 ml 10 mM hydroquinone and 520 ml 3.0 M sodium bisulfite (pH 5.0), vortex to mix and briefly centrifuge. Keep samples in dark (use amber tubes or foil).

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3. Add 50–100 ml sterile mineral oil (to prevent evaporation) and incubate at 50C for 16 h (see Note 2). Remove the mineral oil prior to purification. 4. Purify the sample using the Wizard DNA Clean-up system according to the protocol provided with the kit. Due to the large volume, two columns will be used per sample. Briefly, 1 ml of clean-up resin will be added to 300 ml of sample, mixed and added to a mini-column attached to a vacuum manifold, which is used to draw the mix through. The resin is washed with 2 ml 80% isopropanol. Extract the DNA by incubating resin with 50 ml pre-warmed (65–70C) DNase/RNase-free water for 1 min and elute by centrifugation at 10,000g for 20 s. 5. Add 5.5 ml 3 M NaOH and incubate at room temperature for 10 min 6. Add 1 ml 20 mg/ml glycogen, 33 ml 10 M ammonium acetate and 268.5 ml (3 volumes) cold ethanol 7. Precipitate DNA at –20C for 2 h (or overnight) then centrifuge at 10,000g for 30 min. 8. Remove supernatant and wash pellet with 70% ethanol and centrifuge at 10,000g for 20 min. 9. Remove supernatant, air-dry pellet and re-suspend in 20 ml DNase/RNase-free water. At this point you should have two tubes of purified DNA (generated by using two columns of the Wizard clean-up kit). Typically 2 ml is required per PCR reaction, although 1 ml may also be sufficient. DNA should be stored at –20C but freeze/thaw cycles should be avoided, therefore it may be best to aliquot samples. 3.2. Primer Design

Methylation-specific PCR and bisulfite sequencing PCR rely heavily upon complete bisulfite conversion of DNA. Primer design has to be carefully executed to account for the conversion of DNA as well as the degradation of DNA that accompanies this conversion. A number of free software programs are available for the design of matched methylated and unmethylated MSP primer sets and for BSP primer sets. It is also possible to design primers without the use of these programs but in any case there are important considerations involved in the design of primer sets.

3.2.1. MethylationSpecific PCR Primer Considerations

Methylation-specific PCR works on the premise that methylated DNA and unmethylated DNA will differ slightly in their constitution after bisulfite conversion. All unmethylated cytosines will be converted to uracil after bisulfite treatment and primers should therefore be designed with thymines in place of cytosines. However, methylated cytosines will remain as cytosines and therefore the methylated primer set will include cytosines at CpG sites. For


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example, if a DNA sequence is unmethylated all cytosines will be converted to uracil and a corresponding primer should be designed that incorporates thymine at all cytosine positions. However, if that same DNA sequence was methylated at one or more CpG sites then only the unmethylated cytosines will be converted to uracil and the corresponding primer will incorporate cytosines at the CpG sites. The following table shows examples of these sequences. Unmethylated DNA

Methylated DNA

Initial sequence:

AAC TGA CGT ACT ACG

AAC TGA CmGT ACT ACmG

Converted sequence:

AAU TGA UGT AUT AUG

AAU TGA CGT AUT ACG

Primer sequence:

AAT TGA TGT ATT ATG

AAT TGA CGT ATT ACG

Note that the primer sequence designed for unmethylated DNA is not complementary to the methylated sequence and therefore will not bind. As such, an unmethylated primer set will amplify unmethylated DNA but not methylated DNA and a methylated primer set will only amplify methylated DNA. In this way, if a sample of DNA undergoes two separate PCR amplifications for the methylated and unmethylated primer sets, the presence of an amplified product in one of these reactions will indicate the methylation status of the DNA. 1. As with all PCR, primer specificity is an important issue. Primers should therefore be 20–35 bp in length with an optimum length of 25–30 bp. To keep a similar annealing temperature to standard PCR reactions, the primer is generally longer as GC content is low. All primers that are designed should be interrogated for specificity. It is recommended to assess each of the four primers using methBLAST (http://medgen.ugent.be/methBLAST/), which is based on the NCBI tool BLAST (http://www.ncbi.nlm.nih.gov/ BLAST/) but aligns sequences to the human (or rat or mouse) genome based on conversion of predicted methylation sites (11). It tests for alignment to bisulfite-converted unmethylated DNA, bisulfite-converted methylated DNA and their reverse complements. 2. The primer sequence must contain at least one (and up to three or four) CpG sites. Some investigators consider it best to incorporate as many CpG sites into the primer as possible, allowing for good discrimination between

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methylated and unmethylated DNA. If a CpG island is small (i.e., spanning 30 bp), primers can be designed to cover the entire island. Although this gives information on the methylation status of multiple CpG sites, there are potential problems. If a DNA strand is only partially methylated (i.e., only some CpG sites are methylated) this approach could present problems. Partial methylation and also methylation of only one DNA strand are major drawbacks of MSP. For this reason BSP is often the method of choice, although it is more time consuming. Some prefer to use MSP but try to design primers that incorporate only one CpG site to avoid issues with partial methylation. If the primers contain only one CpG site, this should be located at the very 30 end so that discrimination between the methylated and the unmethylated DNA is maximal. For example:

Methylated primer:

AAT GTT GGA GTA TAG TTT ATG ATT GTC

Unmethylated primer:

AAT GTT GGA GTA TAG TTT ATG ATT GTT

3. Investigation of a heterogeneous population can be performed using MSP. If some cells show methylation and others do not, both methylated and unmethylated primer sets will amplify and a ratio can be determined. This does require full methylation of all CpG sites within the primer set though. 4. It is ideal to run concurrent amplifications with methylated and unmethylated primers and they should therefore be designed to have similar annealing temperatures. However, since the methylated and unmethylated primer sets will have different GC contents, to keep a similar annealing temperature the primers may need to be designed at slightly different positions or be of slightly different lengths. This is true for both the forward and the reverse primers. In this case, it is imperative that the methylated and unmethylated primer sets cover all the same CpG sites, so that the same sites are tested for methylation status. For example: Methylated primer:

GAT CGT AGA TTG TCG ATG TTG ACG TTA

Unmethylated primer:

T TGT AGA TTG TTG ATG TTG ATG TTA GGT GA


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5. It is important to include multiple non-CpG cytosines in the primer to ensure that only bisulfite-converted DNA is amplified. If these are not incorporated, it is possible to amplify DNA that may not have been converted by the sodium bisulfite, which could overestimate the abundance of methylated DNA. For example: No non-CpG sites

Including non-CpG

Unconverted methylated DNA:

AAT CGT AGG TCG

ACT CGT AGC TCG

Unconverted unmethylated DNA:

AAT CGT AGG TCG

ACT CGT AGC TCG

Converted methylated DNA:

AAT CGT AGG TCG

ATT CGT AGT TCG

Converted unmethylated DNA:

AAT TGT AGG TTG

ATT TGT AGT TCG

If there are no non-CpG cytosines the primer designed for the methylated DNA will detect all unconverted DNA as well as the converted methylated DNA and hence indicates that a sample may be methylated when it is not. However, when non-CpG cytosines are included in the primer set, primers will not amplify unconverted DNA. 6. Amplicon size should ideally be between 100 and 500 bp. It is possible to analyze fragments up to about 1 kb, but this may require further optimization. The process of bisulfite conversion is harsh on DNA (50C for 16 h) and it therefore becomes degraded. This means that there will be less DNA fragments of an appropriate size to cover the entire amplicon if it is large. Therefore, if possible maintain a smaller amplicon size. 7. A nested PCR can be performed to increase the sensitivity of detection and is often the method of choice, particularly if DNA is quite degraded after the bisulfite conversion. Nested MSP involves dual PCR amplification whereby the initial PCR amplifies all methylated and non-methylated DNA and a second PCR is used to differentiate the two (using the initial PCR as a template). Design of the first primer set is not specific to methylation status and should follow the guidelines of BSP primer design (Section 3.2.2). Following this, MSP primer sets should be designed within the amplicon generated in the first PCR that are specific to either methylated or unmethylated DNA.

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3.2.2. Bisulfite Sequencing PCR Primer Considerations

Bisulfite sequencing PCR is based on the indiscriminant amplification of a section of methylated or unmethylated DNA, which contains CpG sites within the amplicon but not the primer sequence. It requires only one set of primers to amplify both methylated and unmethylated DNA, which can then be distinguished by subsequent sequencing methods. This method is ideal for the detection of methylation at multiple CpG sites, even if only partial methylation is present in a CpG island. 1. It is important that the primers contain no CpG sites. As bisulfite modification of CpG sites will result in differential nucleotides for methylated and unmethylated DNA, incorporation of a CpG site in the primer will bias amplification of the DNA. If it is unavoidable to include a CpG within the primer sequence, then the primer sequence should be degenerate and have a ‘‘Y’’ at the cytosine site (for detection of C or T), this site should preferably be at the 50 end. This will allow for amplification of both methylated and unmethylated DNA indiscriminantly. For example: Methylated DNA:

ACmG TCC ATG CAT TGA

Unmethylated DNA:

ACG TCC ATG CAT TGA

Primer sequence:

AYG TTT ATG TAT TGA

2. Primers must be specific. Once primers are designed they should be tested for alignment to a bisulfite-converted genome that could be methylated or unmethylated (methBLAST). See point number 1 in Section 3.2.1 for more information. 3. There should be multiple non-CpG cytosines within the primer sequence to ensure that only bisulfite-converted DNA is amplified. If unconverted DNA is also amplified, this will result in a heterogeneous amplicon. If bisulfite-converted and non-converted DNAs are sequenced together, all unmethylated cytosine residues will display a thymine and a cytosine. Depending on the ratio of converted/non-converted DNA this may appear as a heterozygote or as a homozygote, whichever the case the sequence may be difficult to read and analyze. If this occurs at a CpG site, the results may be difficult to interpret. 4. It is best to try to include the entire CpG island within the amplicon as this will be most informative. However, as with MSP, amplicon size is important. Therefore, primers should be designed so that the majority of the CpG island (at least 2/3) is covered. Amplicons up to about 1 kb can be analyzed, sometimes this will require a second round of amplification using the same primer set and 1–2 ml of the first round PCR product as template. Islands which are larger than this may require


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multiple primers to be designed. Designing primers within a CpG island can be difficult as there may not be a long run of nucleotides without a CpG site. In this case, degenerate primers may need to be used (see point 1.). 3.2.3. Primer Design Software

Both MSP and BSP have advantages and disadvantages associated with their use. Choosing one of these methods depends strongly upon the type of experiment being performed. It may even be necessary or preferable to try multiple methods for detection of methylation status. Whichever method is chosen by an investigator, good primer design is crucial. The previous two sections have described some of the major considerations and this information may be sufficient for manual primer design. However, various programs exist that design matched methylated and unmethylated primer sets for MSP or primers for BSP. Some of these are listed in this section. Each of these websites has tutorials or help options, in particular MethPrimer (recommended for MSP) and PerlPrimer (recommended for BSP) are very user friendly. 1. MethPrimer: http://www.urogene.org/methprimer/index. html(12). Designs primers for MSP and BSP. Access is also available to databases containing information on PCR primers for popular DNA methylation analyses, methylated cytosine occurrence, genes affected by promoter CpG island methylation and software tools for handling generated data. A moderated discussion forum is also available through this website. 2. PerlPrimer: http://perlprimer.sourceforge.net/(13). Designs primers for BSP and allows the user to input PCR conditions to give an accurate prediction of primer melting temperature for maximum yield of PCR product. This application is also suitable for standard PCR and QPCR and sequencing. 3. BiSearch: http://bisearch.enzim.hu/(14). Designs primers for MSP and BSP as well as standard PCR. Also allows the user to search various genomes for primer specificity. 4. Methyl Primer Express v1.0: https://www2.appliedbiosystems.com/ (Applied Biosystems – search products – DNA sequencing – software). Designs primers for MSP and BSP.

3.3. Amplification of Bisulfite-Converted DNA

Methylation-specific PCR is performed to directly determine methylation status of one or more CpG sites. It requires amplification of a bisulfite-converted DNA sample using two sets of primers – one specific for DNA methylated at the CpG site/s (referred to as ‘‘M’’) and one specific for DNA unmethylated at the CpG site/s (referred to as ‘‘U’’). When preparing the PCR reactions, ensure that you include two positive controls (if possible include a sample that you know is methylated and one that is not for the gene/CpG island of interest), a no template negative control (water) and an unconverted DNA negative control (this

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will show certain specificity problems with your primer). Bisulfite sequencing PCR typically amplifies a region flanking a CpG island and therefore one set of primers are used. If the island is particularly large, more than one set of primers may be used. In either case, the methylation status of the CpG sites is detected with additional methods (sequencing; cloning and sequencing). As with all PCR, optimization is required for both MSP (methylated and unmethylated primer sets) and BSP for each gene of interest. Typically, optimization may include performing a magnesium titration, a primer titration (forward and reverse primer concentrations could be adjusted and do not need to be equal), adjusting the annealing temperature and/or altering the Taq polymerase and buffer used. The following protocol is a modification of that provided by Herman and colleagues (15). Note that this is only one example, PCR conditions will be dependent on your primers selected. If your gene/CpG island of interest has been amplified before, you may find optimized protocols within journals or in databases such as that provided on the MethPrimer website (see Section 3.2.3 point 1). Also, if a nested PCR approach is required, two separate PCR amplifications will need to be performed and both reactions will need to be optimized. Each PCR reaction should follow standard PCR guidelines for optimization. 1. Prepare one (BSP) or two (MSP) bulk mixes ensuring that you have sufficient bulk mix for all samples and controls. The following table gives an example of a bulk mix prepared for six samples and four controls. Remember, for MSP this has to be made for both a methylated primer set and an unmethylated primer set. Note that at this point, no Taq polymerase is included (see Note 3), due to the need for a hot start reaction and the final volume is 39 ml (although concentrations are based on a 50 ml reaction volume). Methylated/ unmethylated bulk mix Reagent

Final concentration

1

10.5

10 Bisulfite PCR buffer

1

5 ml

52.5 ml

25 mM dNTPs

1.25 mM

2.5 ml

26.25 ml

10 mM forward primer (either U or M)

800 nM

4.0 ml

42.0 ml

10 mM reverse primer (either U or M)

800 nM

4.0 ml

42.0 ml


23.5 ml

246.75 ml

Total

39.0 ml

409.5 ml


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2. Add 39 ml of bulk mix to 1 ml of sample (or control) and denature the sample by heating to 95C for 5 min (see Note 3). 3. While samples are being denatured, make up a Taq polymerase bulk mix containing 1.25 U Taq in 10 ml water per sample. (See Note 3). 4. After 5 min at 95C add 10 ml of water containing 1.25 U Taq polymerase to each sample while keeping the temperature at 95C. 5. Perform the amplification using the following thermocycling conditions: 35 cycles of denaturation at 95C for 30 s, annealing at 30 s (usually 50–60C, depending on primer Tm), extension at 72C for 30 s; followed by a final extension at 72C for 5 min. For larger amplicons (such as 1 kb), extension can occur for 1 min and if amplification is not optimal, annealing time can also be extended to 1 min. 3.4. Methods of Detection

3.4.1. MethylationSpecific PCR

Because MSP uses two sets of primers that amplify DNA with CpG sites that are either methylated or unmethylated, the PCR products can be directly visualized on a gel. Many published protocols use polyacrylamide-based gels but often agarose gels give results that are of equal quality or better. If the size of the amplicon is less than 80 bp, polyacrylamide may be the best method, otherwise agarose is recommended as it is cheaper and easier to handle. As such, this is the method I have included in this chapter; the following protocol is sufficient for detection of methylated and unmethylated primer sets for six samples, two positive controls and two negative controls. If more samples are run a larger gel will need to be made, in this case adjust weights/volumes accordingly. If a polyacrylamide gel is required or preferred, it should be a 6–8% non-denaturing gel and run at 10 V for 1 or more hours. 1. Add 70 ml 1 TAE to 1.4 g of agarose (2% gel) in a conical flask and heat in microwave until agarose is well dissolved (about 2 min, ensure it does not boil over). 2. When cooled, add 2.5 ml 10 mg/ml ethidium bromide (0.5 mg/ml final, wear gloves), mix gently. 3. Pour agarose solution into gel tray and insert comb, allow gel to set before loading samples. 4. While gel is setting, add 3 ml of 100% glycerol solution to each sample (total volume 53 ml) or 10 ml 6 loading dye (total volume 60 ml). 5. Remove comb from gel when set: load 5 ml 100 bp molecular weight marker (with loading dye) in the first lane; load all samples (containing glycerol) for methylated and unmethylated primer sets, positive and negative controls (6 loading dye can be used in place of glycerol if the dye size is not similar to that of your expected band).

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6. Place gel into electrophoresis tank and run the gel at 80–90 V for about 30 min or until you can clearly see your bands of interest using the UV light box. 7. Visualize PCR fragments with a UV light box and capture an image: Methylated positive control: product should only be present with the M primer set Unmethylated positive control: product should only be present with the U primer set No template and non-converted negative controls: no bands should be present A band present in the M primer set indicates the sample is methylated A band present in the U primer set indicates the sample is unmethylated A band present in both primer sets indicated a heterogeneous population (provided there is no contamination and complete conversion, as shown by the negative controls) No bands present in the sample likely indicates mixed CpG methylation within the primer (BSP could be more suitable) or bad sample preparation. 3.4.2. Bisulfite Sequencing PCR

Samples that have undergone bisulfite treatment and amplification using primers that are not specific for methylation status can be sequenced. The sequencing results will indicate methylation status of each CpG dinucleotide in the amplicon. In most cases direct sequencing is sufficient, however if problems occur (e.g., difficult primer optimization for sequencing, heterogeneous population) it is possible to clone the amplified product and then sequence DNA from 8 to 10 clones. As direct sequencing is easier, cheaper and less time consuming, it is the preferred method for analysis and is the protocol given here. As with all sequencing, a forward and a reverse reaction should be carried out independently for each sample, usually using the same primers that were designed for your PCR. 1. Separate the bands by electrophoresis on a 1–2% agarose gel (see Section 3.4.1). Ensure that your negative controls are free from contamination. 2. Using a scalpel or razor, cut the band of interest out of the gel (for each sample, including positive controls) making sure to cut out as little agarose as possible. There should only be a small margin of agarose surrounding the band. 3. Purify the agarose gel using a DNA purification column (available from various companies) according to the manufacturers instructions (see Note 4). Typical elution volumes should be 20–30 ml in DNase/RNase-free water. Run 5 ml of sample on an agarose gel to ensure DNA is still present.


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4. Optional: If a heterogeneous population is present it is possible to clone your DNA, and then sequence 8–10 individual clones. 5. Prepare a bulk mix for your sequencing reaction (prepare one using the forward primer and one using the reverse primer), this should be sufficient for each of your samples and the two positive controls: Reagent

1 forward

1 reverse

8.5

5 Running buffer

2 ml

2 ml

17 ml

BigDye1 Terminator Ready Reaction Mix v3.1

4.0 ml

4.0 ml

34 ml

10 mM forward primer

0.32 ml

10 mM reverse primer Total

6.32 ml

2.72 ml 0.32 ml

or 2.72 ml

6.32 ml

53.72 ml

6. Add 6.32 ml of your bulk mix to 3.68 ml of your purified DNA (see Note 5). 7. Run the sequencing reaction at 96C for 1 min; followed by 25 cycles of 96C for 10 s, 50C for 5 s and 60C for 4 min, if necessary the products can be stored at 4C or –20C. 8. Add 2.5 ml 125 mM EDTA to each tube and then add 25 ml 100% ethanol and invert several times to mix. Leave at room temperature in the dark for 15 min to precipitate the DNA. 9. Centrifuge at 1600–2000 g for 45 min at 4C. 10. Remove supernatant by decanting onto a paper towel (tap tubes upside down on paper towel to remove residual ethanol). 11. Add 30 ml 70% ethanol to each tube and centrifuge at 1600–2000g for 15 min at 4C. 12. Remove supernatant by decanting (as in step 10) and dry the tubes. Tubes can be air dried for 15 min to 1 h or dried in a speed vacuum for 5 min without heat. It is important to protect the pellet from light during this drying stage. At this point products can be stored at –20C in the dark if sequencing is to be done at a later date. 13. Add 5 ml Hi-DiTM formamide to the samples and proceed immediately with capillary electrophoresis using the Applied Biosystems 3730 genetic analyzer (or similar instrument suitable for sequencing). The 3730 instrument requires samples

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Fig. 15.1. Methylated versus unmethylated cytosine sequence.

to be in a 96-well or 384-well plate. Ensure that fresh (every 2–3 days) 1 running buffer and DNase/RNase-free water are in the appropriate trays when using the genetic analyzer. 14. Review your sequencing results using SeqScape software (v2.5) or Sequence Scanner Software (v1.0; free software, limited capabilities). You can differentiate methylated cytosines (C) from unmethylated cytosines (T), as shown in Fig. 15.1.

4. Notes 1. If you have less than 1 mg DNA you can add 1 mg Salmon sperm DNA as a carrier. Although more than 1–2 mg DNA could be used (some protocols suggest 4–10 mg), incomplete bisulfite conversion (due to a higher concentration of DNA) is possible and could lead to false-positive results. If this is a concern, it is recommended to limit the amount of DNA to 1 mg and perform replicate conversions if more DNA is required. DNA needs to be fully denatured in NaOH but to prevent degradation limit incubation time to no more than 30 min. 2. Avoid incubating samples for longer than 16 h as methylated cytosines will start to convert to thymines, leading to falsenegative results. 3. Taq polymerase can be purchased from a number of companies and most will be successful in amplification. This PCR requires a hot start, if using a Taq polymerase such as Platinum Taq from Invitrogen then this protocol can be followed. However various Taq polymerases such as HotStar Taq (Qiagen) and


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JumpStart Taq (Sigma) are designed to be active only when a particular reaction temperature (e.g., 70C) is reached. If using a polymerase such as this, a bulk mix can be made up including the Taq polymerase and water up to 49 ml; steps 3–4 in this protocol can then be omitted (5 min at 95C is still required). If there is difficulty in amplification, JumpStart (or JumpStart Red) Taq polymerase is recommended. Note that each of the polymerases will come with a 10 reaction buffer. This is likely to be more suitable for the PCR reaction than the one provided in this protocol and will also require optimization. I would recommend first trying the PCR with the provided buffer using a magnesium titration (1–4 mM) and annealing temperature optimization, the amount of primer (200 nM final) and dNTPs (200 mM) could also be lowered in this case. 4. Some gel extraction/purification kits may require a specific grade of agarose such as low-melt agarose and often work best when low-percentage gel is used (e.g., 1%). Also, some kits may only be suitable for fragments larger than 100 bp (e.g., Invitrogen), while others are suitable for smaller fragments (70 bp for Qiagen). It is important to check the details of a kit before choosing one, including reagents and equipment needed. 5. For sequencing reactions, as little as 0.5 ml of BigDye can be used in a 10 ml reaction. Optimization of this protocol for each gene can reduce the amount of BigDye used and increase the amount of DNA used in the sequencing reaction if DNA quantity is low. If using smaller volumes of BigDye more 5 sequencing buffer may be required. This can be made in the laboratory with 2 ml 2 M Tris (pH 9.0), 100 ml 1 M MgCl2, and 7.9 ml DNase/RNase-free water (final concentration 400 mM Tris, 10 mM MgCl2).

References 1. Park, S. Y., Kim, B. H., Kim, J. H., Cho, N. Y., Choi, M., Yu, E. J., Lee, S., and Kang, G. H. (2007) Methylation profiles of CpG island loci in major types of human cancers. J Korean Med Sci 22, 311–7. 2. Ducasse, M., and Brown, M. A. (2006) Epigenetic aberrations and cancer. Mol Cancer 5, 60. 3. Moss, T. J., and Wallrath, L. L. (2007) Connections between epigenetic gene silencing and human disease. Mutat Res 618, 163–74. 4. Wilson, A. S., Power, B. E., and Molloy, P. L. (2007) DNA hypomethylation and human diseases. Biochim Biophys Acta 1775, 138–62.

5. Abdolmaleky, H. M., Cheng, K. H., Faraone, S. V., Wilcox, M., Glatt, S. J., Gao, F., Smith, C. L., Shafa, R., Aeali, B., Carnevale, J., Pan, H., Papageorgis, P., Ponte, J. F., Sivaraman, V., Tsuang, M. T., and Thiagalingam, S. (2006) Hypomethylation of MBCOMT promoter is a major risk factor for schizophrenia and bipolar disorder. Hum Mol Genet 15, 3132–45. 6. Kim, J., Kim, J. Y., Song, K. S., Lee, Y. H., Seo, J. S., Jelinek, J., GoldschmidtClermont, P. J., and Issa, J. P. (2007) Epigenetic changes in estrogen receptor beta gene in atherosclerotic cardiovascular tissues and

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7.

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in-vitro vascular senescence. Biochim Biophys Acta 1772, 72–80. Maier, S., and Olek, A. (2002) Diabetes: a candidate disease for efficient DNA methylation profiling. J Nutr 132, 2440S–43S. Munson, K., Clark, J., Lamparska-Kupsik, K., and Smith, S. S. (2007) Recovery of bisulfite-converted genomic sequences in the methylation-sensitive QPCR. Nucleic Acids Res 35, 2893–903. Frommer, M., McDonald, L. E., Millar, D. S., Collis, C. M., Watt, F., Grigg, G. W., Molloy, P. L., and Paul, C. L. (1992) A genomic sequencing protocol that yields a positive display of 5-methylcytosine residues in individual DNA strands. Proc Natl Acad Sci U S A 89, 1827–31. Brena, R. M., Huang, T. H., and Plass, C. (2006) Quantitative assessment of DNA methylation: Potential applications for disease diagnosis, classification, and prognosis in clinical settings. J Mol Med 84, 365–77.

11. Pattyn, F., Hoebeeck, J., Robbrecht, P., Michels, E., De Paepe, A., Bottu, G., Coornaert, D., Herzog, R., Speleman, F., and Vandesompele, J. (2006) methBLAST and methPrimerDB: web-tools for PCR based methylation analysis. BMC Bioinformatics 7, 496. 12. Li, L. C., and Dahiya, R. (2002) MethPrimer: designing primers for methylation PCRs. Bioinformatics 18, 1427–31. 13. Marshall, O. J. (2004) PerlPrimer: crossplatform, graphical primer design for standard, bisulphite and real-time PCR. Bioinformatics 20, 2471–2. 14. Tusnady, G. E., Simon, I., Varadi, A., and Aranyi, T. (2005) BiSearch: primer-design and search tool for PCR on bisulfite-treated genomes. Nucleic Acids Res 33, e9. 15. Herman, J. G., Graff, J. R., Myohanen, S., Nelkin, B. D., and Baylin, S. B. (1996) Methylation-specific PCR: a novel PCR assay for methylation status of CpG islands. Proc Natl Acad Sci U S A 93, 9821–6.

Chapter 16 Biochemical Analysis of Arginine Methylation in Transcription Marc Tini, Hina Naeem, and Joseph Torchia Abstract Protein arginine methylation has emerged as an important mechanism for regulating the functions of proteins involved in diverse aspects of gene regulation such as transcriptional activation and repression, mRNA processing and nuclear-cytoplasmic shuttling. This modification is catalyzed by the PRMT family of enzymes which utilize intracellular S-adenosyl methionine as a cofactor to dimethylate-specific arginines found within many target proteins. The establishment of in vitro biochemical assays as well as the development of modification-specific antibodies, and more recently mass spectrometry, have increased our understanding of the mechanism of catalysis of the PRMT family of enzymes. In the following discussion, we present some of the more commonly used in vivo and in vitro techniques which can be utilized to study the mechanism of arginine methylation and its role in transcription. Key words: Arginine methylation, chromatin, nuclear hormone receptors, PRMT, ChIP.

1. Introduction Protein arginine methylation is an enzymatic-dependent process that is catalyzed by the protein arginine methyltransferase (PRMTs) family of enzymes and plays an integral role in modulating the biological activities of proteins involved in transcription, RNA processing DNA repair, signal transduction and cell differentiation and proliferation (1, 2). PRMTs utilize as a cofactor the methyl donor S-adenosy-l-methionine (AdoMet) – a naturally occurring compound present in all living cells. Arginine can be methylated on either or both of its two terminal guanidino nitrogen atoms resulting in the formation of asymmetric or symmetric dimethylarginines, respectively. There are currently nine Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_16 Springerprotocols.com

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human PRMTs which have been identified (2, 3). Eight have been shown to possess catalytic activity and are classified as either type I or type II enzymes, based on their methylation reaction products. The Type I PRMTs consisting of PRMT1, PRMT2, PRMT3, PRMT4/CARM1 PRMT6, and PRMT8 catalyze the formation of asymmetric dimethylarginine residues, whereas the Type II PRMTs consisting of PRMT5, PRMT7 and PRMT9 catalyze the formation of symmetric dimethylarginine residues. Both types generate a monomethylated arginine as a reaction intermediate. The discovery that CARM1/PRMT4 is a histone methyltransferase associated with the steroid receptor coactivator (SRC) family of proteins provided the first indication of a role of methylation in transcriptional activation (4). Consequently, numerous studies have demonstrated that recruitment of both PRMT1 and CARM1 plays an important role in nuclear receptor-mediated gene transcription. CARM1 interacts with all three steroid receptor coactivator (SRC) family members (4–7) and can enhance the hormone-dependent activation of reporter genes by multiple NRs when coexpressed with SRC coactivators. In the absence of SRC coactivators, PRMTs have no effect on receptor function. This has led to a model suggesting that PRMTs acts as secondary coactivators for NRs by binding to SRC coactivators resulting in chromatin modification of target genes. In support of this, chromatin immunoprecipitation (ChIP) studies have shown that CARM1 is recruited to hormone-responsive promoters in response to hormone stimulation (6). In vitro methylation assays have revealed clear specificity differences among the various PRMT proteins. For example, CARM1 specifically methylates histone H3 at Arg 2, 17 and 26 (8, 9) whereas PRMT1, recognizes Arg 3 on histone H4. In addition to histones, PRMTs have been shown to methylate other transcriptional regulatory proteins. For example, CARM1 specifically methylates the transcriptional coactivator CBP/p300 and mutation of the methylation sites abolishes steroid-dependent transcriptional activation (10, 11). More recently, CARM1-dependent methylation of the transcriptional coactivator p/CIP has been shown to be involved in both protein stability and coactivator complex dissociation, thus establishing arginine methylation as a critical regulatory signal in nuclear receptor-mediated transcription (12). In general, PRMT1 methylates arginines found within glycine-rich regions, referred to as GAR motifs, found in many proteins possessing RNA-binding properties (13–15). In contrast, CARM1 displays a higher degree of specificity and methylates arginines found within proline and arginine-rich domains known as PGM motifs which have been identified in proteins involved in chromatin remodeling and in RNA splicing (16).

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In the following discussion we describe in vitro and in vivo techniques employed to study arginine methylation in nuclear receptor signaling events. For the most part, the techniques have been adapted from methods used previously to characterize other covalent modifications such as phosphorylation and acetylation. Arginine methylation is commonly analyzed in vitro using purified recombinant methyltransferase proteins to radiolabelspecific substrates with S-adenosyl-L-[methyl-3H]-methionine. Alternatively, cell extracts may be used as a source of both substrate and methyltransferase activity. In this case, cells may be grown in the presence of adenosine dialdehyde (AdOx), an indirect methyltransferase inhibitor, resulting in the accumulation of hypomethylated proteins that may be immunoprecipitated and used as substrates for in vitro methylation reactions. Regardless of the protein source, it is critically important to demonstrate that methylation occurs in the context of the full-length protein which can be generated by bacterial expression, or for larger proteins, using a baculovirus expression system. Detailed molecular mapping of the methylation sites may be performed using recombinant fusion proteins or synthetic peptides. Generally, recombinant protein fragments are initially employed to roughly map methylated regions; subsequently, peptide substrates may be used for further mapping. However, caution should be exercised when using recombinant proteins or peptides as potential substrates because improper folding may mask or expose methylation sites that are not recognized by the enzyme in the native protein. We have found that biotinylated peptides provide a rapid and highly reproducible method to test for substrate specificity once the approximate location of the methylated arginine has been determined. Furthermore, in this context, amino acid substitutions can be used to assess the requirement for specific amino acids in substrate recognition. The addition of the biotinylated moiety allows the capture of the peptide on a solid support matrix containing covalently bound streptavidin which facilitates washing and removal of unwanted contaminants prior to analysis. Once the relevant arginines have been identified, site-directed mutagenesis is used to generate amino acid substitutions in the full-length protein in order to confirm that these residues are indeed methylated in this context and to permit functional analysis. In vitro studies can provide evidence that a protein is a substrate for covalent modification but this does not necessarily imply that the modifications occur in living cells. The traditional method of detecting in vivo methylation involves metabolic labeling of intact cells with radiolabeled methionine in the presence of translation inhibitors, followed by immunoprecipitation of the protein of interest and analysis by SDS-PAGE and fluorography (17). Alternatively, methyl-arginine-specific antibodies can also be

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used to detect argine methylation by immunoblotting of crude cell lysates or immunoprecipitated proteins. This approach has been used to successfully identify methylated arginines in histones as well as non histone proteins. Several commercially available antibodies which have been raised against synthetic peptides of GAR motifs (Sym10, Sym11 from Upstate biotechnology) can be used for this purpose. In addition, we have used the H3R17me2 antibody available from Upstate biotechnology which recognizes dimethylated arginine 17 on histone H3, a major substrate for CARM1. By immunoprecipitating the transcriptional coactivator p/CIP from cell lysates, followed by Western blotting using the H3R17me2 antibody we have been able to demonstrate that p/CIP is specifically methylated in intact cells in a CARM1-dependent manner. More recently, mass spectrometry (MS) is becoming a more popular method to characterize post-translational modifications. Upon methylation, a 14 Da increase per incorporated methyl group can be used to identify methylation of purified endogenous proteins (18). In addition, MS has been used successfully to monitor in vitro methylation of recombinant proteins (19). A major limitation of all these approaches is that, depending on the protein, the relative abundance of the methylated protein pool may be limiting. In the case of MS which requires enzymatic degradation of the protein, the entire protein sequence may not be covered completely by the detected peptides or may be below the detection limit. Therefore potential methylation sites may be missed. To circumvent this problem epitope-tagged substrate and methyltransferase proteins can be overexpressed in order to obtain sufficient quantities of the methylated substrate. This can be performed in mammalian cells or other expression systems. We have overexpressed the transcriptional coactivator p/CIP in insect cells, in the presence and absence of CARM1, to identify functionally relevant methylation sites. Finally we describe a standard ChIP protocol that may be used to assess the promoter occupancy of CARM-1 and p/CIP on estrogen-regulated genes. The chromatin immunoprecipitation assay (ChIP) is a powerful tool to assess the promoter occupancy of transcription factors as well as for monitoring chromatin modifications at specific target genes.

2. Materials 2.1. Purification of Recombinant Proteins

1. Sf-900 II SFM (1 ), liquid (Invitrogen # 10902161) 2. Cellfectin (Invitrogen # 10362010) 3. Benzonase (Sigma # E1014)


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4. Lysis buffer: 20 mM Tris–HCl pH 7.9, 300 mM KCl, 1.5 mM MgCl2,0.5 mM DTT, 0.5 mM EDTA, 20% glycerol, 0.1% NP40, 0.2 mM PMSF, 5 ug/ml leupeptin, aprotinin, pepstatin. 5. Wash buffer: 20 mM Tris–HCl pH 7.9, 100 mM KCl, 1.5 mM MgCl2, 0.5 mM DTT, 0.5 mM EDTA, 0.1 % NP40, 10% glycerol, 0.1 mM PMSF, 6. Elution buffer: 20 mM Tris–HCl pH 7.9, 100 mM KCl, 0.2 mM EDTA, 1.5 mM MgCl2,10 % glycerol, FLAG peptide 0.2 mg/ml 7. ANTI-FLAG M2 agarose (SIGMA # A2220) 8. FLAG peptide (SIGMA # F3290) 9. PGEX-4T1 vector (GE Healthcare # 27-4580-01) 10. IPTG (isopropyl b-D-thiogalactopyranoside):(Sigma I5502): dissolve 25 mg/ml IPTG in double distilled water and store at –20C 11. Glutathione sepharose 4B (GE Healthcare # 27-4574-01) 2.2. In Vitro Methylation Reactions

1. Reaction buffer: 50 mM Tris–HCl pH 8.5, 20 mM KCl, 10 mM MgCl2, 1 mM b-mercaptoethanol, 0.1 M sucrose 2. Wash buffer: 50 mM Tris–HCl pH 8, 150 mM NaCl, 1 mM EDTA, 0.5% NP-40 3. Peptides: resuspend peptides in water at a concentration of approximately 1 mg/ml, aliquot and stored at –20C 4. MagnaSphere streptavidin beads (Promega)

2.3. Chromatin Immunoprecipitation Assay

1. Charcoal-stripped fetal bovine serum 2. 37% formaldehyde 3. 1 PBS 4. Cell lysis buffer: 50 mM Tris–HCl pH 8.0, 10 mM EDTA, 1% SDS, protease inhibitors 5. Protease inhibitor cocktail (Roche) 6. Dilution buffer: 10 mM Tris–HCl pH 8.0, 1% Triton X-100, 2 mM EDTA, 150 mM NaCl protease inhibitors 7. Wash buffer A: 20 mM Tris–HCl pH 8.0, 150 mM NaCl, 2 mM EDTA, 0.1% SDS, 1% Triton X-100, 10% glycerol 8. Wash buffer B: 20 mM Tris–HCl pH 8.0, 500 mM NaCl, 2 mM EDTA, 0.1% SDS, 1% Triton X-100, 10 % glycerol 9. LiCl–detergent buffer: 10 mM Tris–HCl pH 8.0, 0.25 M LiCl, 1% NP-40, 1% deoxycholate, 1 mM EDTA 10. Elution buffer: 1% SDS–0.1 M NaHCO3

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3. Methods 3.1. Expression of Recombinant CARM1 and p/CIP Protein Using Baculovirus

To generate recombinant proteins using baculovirus, we routinely use the Bac-To-Bac Baculovirus Expression System available from Invitrogen. 1. The cDNA for CARM1, or other methyltransferases, is subcloned into pFastBac plasmid vector. To facilitate purification of recombinant CARM1 we have incorporated a FLAG epitope at the amino terminus which is in-frame with the initiating methionine. 2. Transform competent DH10 Escherichia coli cells containing the bacmid with a mini-attTn7 target site. The transposition between the Tn7 element on the pFastBac donor plasmid and the Tn7 target site on bacmid results in colonies containing the recombinant bacmid identified by disruption of the lacZ gene which can be identified as white colonies on agarose plates (Note 1). 3. Allow 48 h incubation (at 37C) so that positive colonies can be clearly identified. Pick several white colonies and incubate overnight in 5 ml LB containing the appropriate selection markers. 4. The following day, Sf9 cells are seeded at a density of 9 105 cells/35-mm well in 2 ml of SF-900 II medium containing 50 units/ml penicillin, 50 mg/ml streptomycin. Cells are allowed to adhere to the plates at 27C for 2 h. During this time, mini-prep DNA is prepared from the positive colonies grown overnight. The bacmid DNA is high molecular weight so care should be exercised during preparation to prevent shearing (avoid vortexing the sample). Carefully resuspend the DNA in 50 ml of TE buffer and transfect 5–20 ml into each well using Cellfectin reagent. A mock transfection may also be included containing no DNA. 5. The cells are incubated for 72 h at 27C and the supernatant corresponding to each sample is removed and stored at 4C. Approximately 200 ml of 1 SDS-PAGE sample buffer containing 5 mg/ml benzonase (added directly to the sample buffer immediately prior to use) is added to the cell monolayer and incubated at room temperature with shaking. The resulting lysate is directly loaded onto an SDS-PAGE gel and the expression of the recombinant protein analyzed by Western blotting using FLAG antibody. 6. Once protein expression is confirmed, the viral stock is amplified by inoculating a 50 ml Sf9 insect cell culture with a 100 ml aliquot of the corresponding supernatant. The suspension culture is then incubated for 48 h with shaking at 27C.


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After 48 h the amplification process is repeated. Depending on the virus, we have found that two consecutive amplifications usually yield a sufficiently high titer virus (> 109 pfu) to produce ample protein. 7. To express the desired protein, a 100 ml suspension culture is established (1.5 106 cells/ml) 24 h prior to inoculation. The next day the cells are counted and the culture is inoculated with the amplified virus at a multiplicity of infection (M.O.I) of 5–10. 8. At 48 h post-infection, cells are pelleted by centrifugation at 1000 g for 5 min, washed with ice-cold PBS and resuspended in 5 ml of lysis buffer, disrupted by 10 strokes with a dounce homogenizer and incubated on ice for 15 min. The cell lysate is then clarified by centrifugation at 100,000 g for 30 min. 9. To purify CARM1 or p/CIP, the supernatant is combined with 200 ml of a 50% slurry of FLAG M2 sepharose and the slurry is mixed for 2 h at 4C on a rocking platform. The beads are washed once with 5 ml lysis buffer and three times with 5 ml wash buffer. 10. The bound proteins are eluted with three consecutive 200 ml washes (5 min each wash with rocking at 4C) with elution buffer containing 0.2 mg/ml FLAG peptide. 11. Protein expression is confirmed by SDS-PAGE and Coomassie blue staining (Note 2). The purified protein is then dialyzed against elution buffer without peptide and stored at –80C. 3.2. Coexpression of p/CIP and CARM1 for Analysis by Mass Spectrometry

1. Expression and purification of p/CIP and CARM1 are performed exactly as described in Section 3.1 except that two suspension cultures are established, a 100 ml suspension culture inoculated with virus for p/CIP alone, and a second culture inoculated with viruses for both p/CIP protein and CARM1. 2. The complexes are immunopurified using FLAG M2 agarose as described in Section 3.1. The eluants are then separated by SDS-PAGE on a large preparative gel and stained with Coomassie blue. The 160 kDa band corresponding to p/CIP is excised and analyzed by mass spectrometry (Note 3).

3.3. Expression of Recombinant p/CIP Fusion Proteins in Bacteria

1. Glutathione S-transferase (GST) fusion proteins are generated by cloning various regions of p/CIP into the pGEX bacterial expression vector in-frame with the GST moiety and the resulting plasmids were transformed into the Escherichia coli strain BL-21. 2. A single colony of transformed BL21 was grown overnight at 37C in 5 ml LB media containing 100 mg/ml ampicillin. The following day, the overnight culture is diluted 1:100 in 100 ml of LB media and grown at 30C with shaking until an optical density of 0.5 at 600 nm is reached, at which time protein

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expression was induced with 0.5 mM of isopropyl-b-Dthiogalactopyranoside (final concentration) for 5 h at room temperature. 3. Cells are pelleted by centrifugation at 4C for 20 min at 10,000 g and resuspended in 10 ml of lysis buffer followed by incubation on ice for 30 min with occasional mixing. The resuspended cells are then lysed by sonication (4 15 s bursts @ 50% intensity) and centrifuged at 4C for 1 h at 30,000 g to pellet cell debris. 4. A total of 500 ml of a 50% slurry of glutathione-sepharose beads prewashed in lysis buffer is added to the clarified supernatant, and the samples are incubated at 4C with rocking for 1 h. The beads are washed twice with lysis buffer and three times with wash buffer. The bound proteins are eluted with 250 ml of elution buffer containing 10 mM reduced glutathione in three successive elutions with 10 min rocking at 4C. 5. Protein expression is confirmed by SDS-PAGE and Coomassie blue staining. The eluted proteins are dialyzed against elution buffer without glutathione and stored at –80C. 3.4. In Vitro Methyltransferase Assay Using Cell Extracts

1. Wild-type 13 dpc mouse embryonic fibroblasts (MEFs) or CARM1 knockout MEFs are grown on 150 mM plates to approximately 90% confluency, washed twice with 10 ml PBS and harvested (Note 4). 2. Centrifuge cells at 1000 g for 5 min and the cell pellet is resuspended in 500 ml of lysis buffer and left on ice for 10 min with occasional pipetting. The extracts are centrifuged for 15 min at 15,000 g and the supernatant is transferred to a fresh eppendorf tube. 3. One microgram of FLAG-tagged p/CIP is added to 50 ml of a 50% slurry of FLAG-sepharose in 200 ml PBS. The mixture is incubated for 1 h at 4C. The beads are then spun down at 1000 g for 30 s and washed four times with PBS. 4. The FLAG-sepharose containing the bound p/CIP is incubated in 10 ml of 5 reaction buffer, 10–40 ml cell extract (10 mg/ml) from wild-type or CARM1 knockout mouse embryonic fibroblasts (MEFs) and 0.5 uCi S-adenosyl-L[methyl -3H] methionine ({3H}AdoMet). 5. The reaction is incubated at 30C for 1 h and the beads are washed three times with methylation wash buffer. 6. The substrate bound beads are then suspended in SDS sample buffer, boiled and separated by 10% SDS-PAGE. The separated proteins are transferred to poly(vinylidene difluoride) membrane, sprayed with EN3HANCE (Perkin–Elmer) and exposed to film for approximately 48 h.


3.5. Assay for Arginine Methylation Using Purified Recombinant Proteins

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1. In an eppendorf tube on ice add 4 ml of 5 reaction buffer, 0.5–1.0 ug of recombinant protein, 100 ng of purified recombinant CARM1 (or other methyltranferase proteins) and 0.5 uCi{HAdoMet}. The mixture is taken up to a final volume of 20 ml with water 2. Reactions are allowed to proceed at 30C for 1 h. The reactions are mixed 1:1 (v/v) with SDS sample buffer and the methylation of recombinant proteins is analyzed by SDSPAGE and fluorography. The gel is then washed in 5% (v/v) acetic acid–10% (v/v) methanol for 30 min at RT and treated with EN3HANCE according to the manufacturer’s instructions. The gel is dried and methylation is analyzed by fluorography.

3.6. Assay for Arginine Methylation Using Biotinylated Peptides as Substrates

1. The assay conditions are identical to those described in the previous section except that the recombinant substrate protein is replaced with 0.5–1 mg of the biotinylated peptide. 2. After the methylation reaction is complete, the reaction products are diluted to 200 ml with 1 methylation reaction buffer. 3. To capture the biotinylated peptide, the mixture is added to an eppendorf tube containing 25 ml MagnaSphere streptavidin beads (Promega). Biotin–streptavidin interactions are performed for 1 h at 4C with rotation. The beads are then washed three times with wash buffer for 10 min with rocking at 4C and radioactivity is measured by liquid scintillation counting.

3.7. Using MethylSpecific Antibodies to Assess Arginine Methylation

1. MCF7 cells are grown on 150 mM plates in the presence and absence of 50 mM AdOx for at least 48 h. 2. Wash cells twice with PBS, harvest and centrifuge at 1000 g for 5 min. The cell pellet is resuspended in 1 ml lysis buffer and centrifuged at 10,000 g for 10 min. The supernatant is transferred to a fresh tube and precleared by adding 50 ml of a 50% slurry of protein A-sepharose. The mixture is then centrifuged at 1000 g for 1 min and protein concentration is determined. 3. Approximately 6 mg of p/CIP antibody is added to 1 mg of MCF7 cell extract for 2 h on ice. Fifty microliters of a 50% slurry of protein A-sepharose is added for 1 h at 4C with rocking. The antigen antibody complex is then washed four times with lysis buffer and the beads are then resuspended in SDS sample buffer and the complex is separated by SDSPAGE. Western blotting is performed using both the p/CIP antibodies to validate immunoprecipitation from both control cells and AdOX-treated cells, and the H3R17me2 antibody (Note 5).

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3.8. In Vivo Detection of Arginine Methylated Protein by Metabolic Labeling of Cells

The following procedure is a modified version of the method described by Desrosiers and Tanguay (20). 1. Five 100 mm plates of adherent MCF7 cells are grown to approximately 75 % confluency. Depending on the nature of the experiment (e.g., to determine the effects of hormone stimulation on methylation) cells may be grown in charcoal-stripped media for approximately 48 h prior to metabolic labeling. 2. On the day of the experiment, the cells are washed twice with PBS and protein translation is inhibited by adding cyclohexamide 50 and 10 mg/ml chloramphenicol in medium for approximately 30 min. 3. Wash cells twice with PBS and replace with methionine deficient DMEM and containing 5% dialyzed FBS, cyclohexamide and chloramphenicol for 60 min. 4. Prepare a cell-labeling mixture consisting of DMEM supplemented with 5% dialyzed FBS, 50 mg/ml cyclohexamide and containing 20 mCi/ml of L-[methyl-3H]methionine. The amount of labeling mix depends on the number of plates. For 100 mm plates we add an amount needed to just cover the cells (approximately 1–2 ml/plate). Continue metabolic labeling for 3 h at 37C. 5. Wash cells twice with PBS, harvest and resuspend the pellet in 1 ml lysis buffer. The pellet is left on ice for 15 min and then centrifuged at 15,000 g for 30 at 4C and stored at –80C. 6. p/CIP is then immunoprecipitated with specific antibody as described in Section 3.7.

3.9. Chromatin Immunoprecipitation of CARM1 Target Genes

1. To remove molecules from the medium which may potentially stimulate the estrogen receptor, MCF-7 cells are grown on 150 mm plates in phenol red-free DMEM containing 5%–10% charcoal-stripped FBS for at least 48 h prior to stimulation with 10–7 M b estradiol. Stimulation for 30 min is usually sufficient to detect promoter occupancy of target genes by the estrogen receptor and its associated coactivators. We normally combine chromatin from two 150 mm plate at approximately 90% confluency for three immunoprecipitation. 2. Cells are then cross-linked with 1% formaldehyde (added directly to the media) at room temperature for approximately 10 min depending on the cell type. Optional: Cross-linking can be terminated by adding ice cold glycine to a final concentration of 0.125 M and incubated for 5 min. 3. Wash cells twice with ice-cold PBS containing 0.5 mM EDTA, harvest and pellet at 1000 g for 5 min. The cell pellets are lysed in 0.4 ml of cell lysis buffer and incubated on ice for 10 min.


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4. Cell lysates are sonicated to yield DNA fragments ranging in size from 300- to 1000-bp (Note 6). Dilute supernatants 10-fold with dilution buffer and preclear with 60 ml of 50% slurry of protein A-sepharose containing 2.5 mg of sheared salmon sperm DNA for 1 h at 4C. Pellet beads by centrifugation and transfer supernatant to a clean eppendorf tube and immunoprecipitation is performed overnight at 4C with 1.0–5 mg of specific antibody. 5. After antibody incubation, 60 ml of protein A-sepharose containing 2.5 mg of salmon sperm DNA per ml is added and incubated for 2 h at 4C. The beads are washed once with 1 ml wash buffer A, two times with 1 ml wash buffer B and one time with 1 ml LiCl–detergent buffer. The samples are then washed two times with 1 ml TE buffer. 6. Immuncomplexes are then extracted twice with 200 ml elution buffer (incubate for 15 min while shaking at room temperature during each incubation). The supernatants are pooled and NaCl is added to a final concentration of 200 mM and the cross-linking was reversed by heating at 65C 6 hr to overnight. 7. The DNA is purified using Qiagen PCR purification spin columns and eluted in a final volume of 60 ml. 8. The DNA is resuspended in TE buffer and analyzed by PCR using gene-specific primers that should ideally amplify fragments between 200 and 400 bp.

4. Notes 1. The number of positive colonies obtained varies depending on the insert and quality of the DNA. In order to confirm successful recombination, PCR may be used with primers corresponding to the insert. Generally however the false positive rate is very low. When plating out transformed DH10 cells it is important to plate out several concentrations of the transformed cells. Plating the transformed cells at too high a concentration increases the ambiguity between the blue negative colonies and the white colonies containing the bacmid. We have found that a concentration of approximately 50–100 colonies per plate is optimal. 2. The total yield obtained using this method will vary depending on the protein. Generally however for most proteins we have found that a 100 ml suspension culture will be sufficient to obtain approximately 20–100 mg of total protein.

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3. The amount of protein required for mass spectrometry depends on a number of factors and should be discussed with the facility performing the analysis. First the relative abundance of the methylated protein is critical as the digestion process is not 100% efficient and the amount of methylated protein may be limiting. As a general rule we have found that if the protein is visible by Coomassie staining then the probability of a positive identification will be high. 4. The amount of cell extract required to obtain enough enzymatic activity will depend on the cell type. Protein concentration should be kept high and several concentrations should be tested in order to detect enzymatic activity. 5. The H3R17me2 antibody sold by Upstate is raised against a peptide containing dimethylated arginine 17 on histone H3. We and others have found that it does recognize dimethylated arginines in several proteins. However, whether this antibody recognizes a specific dimethylation must be experimentally determined for each individual protein. 6. Correct sonication is critical to the overall success of the procedure and is a major source of variability. All sonication should be performed on ice and an appropriate tip should be used so that it can immerse deep enough into the solution to prevent any frothing.

References 1. Lake, A.N. and M.T. Bedford. 2007. Protein methylation and DNA repair. Mutat Res 618:91–101. 2. Bedford, M.T. and S. Richard. 2005. Arginine methylation an emerging regulator of protein function. Mol Cell 18:263–272. 3. Lee, Y.H., S.A. Coonrod, W.L. Kraus, M.A. Jelinek and M.R. Stallcup. 2005. Regulation of coactivator complex assembly and function by protein arginine methylation and demethylimination. Proc Natl Acad Sci U S A 102:3611–3616. 4. Chen, D., H. Ma, H. Hong, S.S. Koh, S.M. Huang, B.T. Schurter, D.W. Aswad and M.R. Stallcup. 1999. Regulation of transcription by a protein methyltransferase. Science 284:2174–2177. 5. Koh, S.S., D. Chen, Y.H. Lee and M.R. Stallcup. 2001. Synergistic enhancement of nuclear receptor function by p160 coactivators and two coactivators with protein methyltransferase activities. J Biol Chem 276:1089–1098.

6. Ma, H., C.T. Baumann, H. Li, B.D. Strahl, R. Rice, M.A. Jelinek, D.W. Aswad, C.D. Allis, et al. 2001. Hormone-dependent, CARM1-directed, arginine-specific methylation of histone H3 on a steroid-regulated promoter. Curr Biol 11:1981–1985. 7. Chen, D., S.M. Huang and M.R. Stallcup. 2000. Synergistic, p160 coactivator-dependent enhancement of estrogen receptor function by CARM1 and p300. J Biol Chem 275:40810–40816. 8. Chen, S.L., K.A. Loffler, D. Chen, M.R. Stallcup and G.E. Muscat. 2002. The coactivator-associated arginine methyltransferase is necessary for muscle differentiation: CARM1 coactivates myocyte enhancer factor-2. J Biol Chem 277:4324–4333. 9. Schurter, B.T., S.S. Koh, D. Chen, G.J. Bunick, J.M. Harp, B.L. Hanson, A. Henschen-Edman, D.R. Mackay, et al. 2001. Methylation of histone H3 by coactivator-associated arginine methyltransferase 1. Biochemistry 40:5747–5756.


10. Xu, W., H. Chen, K. Du, H. Asahara, M. Tini, B.M. Emerson, M. Montminy and R.M. Evans. 2001. A transcriptional switch mediated by cofactor methylation. Science 294:2507–2511. 11. Chevillard-Briet, M., D. Trouche and L. Vandel. 2002. Control of CBP co-activating activity by arginine methylation. Embo J 21:5457–5466. 12. Naeem, H., D. Cheng, Q. Zhao, C. Underhill, M. Tini, M.T. Bedford and J. Torchia. 2007. The activity and stability of the transcriptional coactivator p/CIP/SRC-3 are regulated by CARM1-dependent methylation. Mol Cell Biol 27:120–134. 13. Boisvert, F.M., A. Rhie, S. Richard and A.J. Doherty. 2005. The GAR motif of 53BP1 is arginine methylated by PRMT1 and is necessary for 53BP1 DNA binding activity. Cell Cycle 4:1834–1841. 14. Boisvert, F.M., M.J. Hendzel, J.Y. Masson and S. Richard. 2005. Methylation of MRE11 regulates its nuclear compartmentalization. Cell Cycle 4:981–989. 15. Boisvert, F.M., J. Cote, M.C. Boulanger and S. Richard. 2003. A proteomic analysis of arginine-methylated protein complexes. Mol Cell Proteomics 2:1319–1330.

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16. Cheng, D., J. Cote, S. Shaaban and M.T. Bedford. 2007. The arginine methyltransferase CARM1 regulates the coupling of transcription and mRNA processing. Mol Cell 25:71–83. 17. Liu, Q. and G. Dreyfuss. 1995. In vivo and in vitro arginine methylation of RNA-binding proteins. Mol Cell Biol 15: 2800–2808. 18. Zou, Y. and Y. Wang. 2005. Tandem mass spectrometry for the examination of the posttranslational modifications of highmobility group A1 proteins: symmetric and asymmetric dimethylation of Arg25 in HMGA1a protein. Biochemistry 44: 6293–6301. 19. Kim, S., B.M. Merrill, R. Rajpurohit, A. Kumar, K.L. Stone, V.V. Papov, J.M. Schneiders, W. Szer, et al. 1997. Identification of N(G)-methylarginine residues in human heterogeneous RNP protein A1: Phe/Gly-Gly-Gly-Arg-Gly-Gly-Gly/Phe is a preferred recognition motif. Biochemistry 36:5185–5192. 20. Desrosiers, R and R.M. Tanguay, 1988. Methylation of Drosophila histones at proline, lysine, and arginine residues during heat shock. J Biol Chem. 263:4686–92.

Chapter 17 In Vitro Histone Demethylase Assays Kenji Kokura and Jia Fang Abstract Histone methylation plays important roles in chromatin structure, transcription, and epigenetic state of the cell. Tremendous discoveries recently demonstrated that methylation mark is not static but is dynamically regulated by both histone methyltransferases and the histone demethylases. Two families of histone demethylases have been identified to remove methyl groups from lysine side chain through different reaction mechanisms in presence of distinct cofactors. Amine oxidase LSD1 family requires flavin adenine dinucleotide (FAD) whereas dioxygenase Jmjc domain-containing proteins family relies on Fe(II) and alpha-ketoglutarate. Identification of these enzymes opened a new era in understanding how chromatin dynamic is regulated and further understanding the regulation of these enzymes will provide significant insights into fundamental mechanisms of many biological processes and human diseases. This chapter describes different assay conditions and detection methods for different family of histone demethylases. We also summarize step-by-step protocols for purification and preparation of various histone substrates for histone demethylase assays. Key words: Epigenetic regulation, histone methylation, histone demethylation, histone demethylase, oxidative hydroxylation, chromatin, nucleosomes, histone.

1. Introduction Post-translational modifications on histone tails have essential roles in regulating chromatin dynamics and transcription (1, 2). As one of the best studied modifications, histone methylation occurs on both lysine and arginine residues and regulates fundamental processes such as heterochromatin formation, transcription, and X-chromosome inactivation (3). Within the same residue, lysine side chain could be mono-, di-, or tri-methylated, whereas arginine side chain could be mono- or di-methylated by three families of histone methyltransferases (HMTases), the Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_17 Springerprotocols.com

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PRMT family, the SET domain-containing proteins, and the nonSET domain proteins DOT1/DOT1L. Given that different methylation sites and different methylation states on the same residue could lead to different functional outcomes, histone methylation provides numerous combinatorial potentials for the functional consequences (4). Although it was historically assumed that histone methylation is an irreversible epigenetic mark, numerous recent studies have revealed that methyl groups on histone tails can be actually removed through different enzymatic reactions. This reversibility obviously adds an additional layer of complexity to the chromatin regulatory functions of histone methylation. Seeking for putative histone demethylases lasted over 40 years since Paik and coworkers purified N6-methyl-lysine oxidase from rat kidney capable of demethylating free mono- and di-methyl lysine residues in 1964 (5). One of the major obstacles is lack of reliable and sensitive in vitro assays. Human arginine deiminase PADI4/PAD4 was the first enzyme identified to turn over the methyl group from histones by converting monomethylated arginine to citrulline (6, 7), but it is not a strict histone demethylase. In the same year, the nuclear amine oxidase LSD1 was reported as the first bona fide histone lysine demethylase to directly remove methyl groups from histone H3-K4 through flavin adenine dinucleotide (FAD)-dependent amine oxidation (8). However, LSD1 only mediates mono- and di-demethylation and there are only two LSD1 homologues in the mammalian genome, indicating that additional demethylases exist using different enzymatic mechanisms. Based on the reaction mechanism used by AlkB family of proteins to demethylate damaged DNA (9, 10), a novel in vitro histone demethylase assay was developed recently based on detection of release product formaldehyde (11, 12). And a large family of JmjC domain-containing histone demethylases was thus uncovered by an unbiased biochemical purification approach (11). Many proteins in this family have been demonstrated to use Fe(II) and alpha-ketoglutarate as cofactors to mediate a hydroxylation-based histone demethylation and mediate important functions (13, 14). Here, we describe a few histone demethylase assays based on different detection methods, as well the substrate preparations for the assays. Nucleosomes or core histone octamers are first purified from large-scale cultured HeLa-S3 cells through micrococcal nuclease (MNase) digestion followed by sucrose gradient ultracentrifugation and hydroxyapatite chromatography. Resulting histones can be directly used as substrates for the Western blotbased histone demethylase assays using site-specific methylhistone antibodies. These histones can be also radioactively labeled with S-adenosyl-L-[methyl-3H]-methionine by various HMTases. After purification, radioactive-labeled histones are

Histone Demethylase Assays

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incubated with different histone demethylase samples under two different demethylation assay conditions and employed for the formaldehyde release assays. The type I assay condition is designed for Jmjc domain-containing histone demethylases, which require Fe(II) ion and alpha-ketoglutarate as cofactors to catalyze radical-based oxidative hydroxylation. During this reaction, the hydroxyl–methyl groups are released spontaneously from lysine side chain as formaldehyde, while alpha-ketoglutarate is converted to succinate and carbon dioxide (11). Ascorbic acid is also included in the reactions to regenerate Fe(II) from Fe(III) (11, 15). The type II reaction is suitable for the LSD1 family of histone demethylases which rely on cofactor FAD. This reaction hydroxylates lysine methylamine group and releases formaldehyde, while FAD is reduced to FADH2 (8). Released radioactive [3H]-formaldehyde under both assay conditions can be purified by removal of all proteins through trichloro acetic acid (TCA) precipitation. To further improve the sensitivity, formaldehyde in supernatant is then converted to 3,5-diacetyl-1,4-dihydrolutidine (DDL) by modified NASH method (16). DDL stays in organic phase during following extraction step with 1-pentanol and can be separated from any other radioactive contaminations in the aqueous phase. And finally, the radioactivity of DLL is measured by liquid scintillation counting. As supplemental assays, we also describe a transfection-based immunostaining assay which can be used to test whether a particular protein has the histone demethylase activities.

2. Materials 2.1. Purification of Nucleosomes and Histone Octamers

1. Cultured HeLa-S3 cells (20 L), harvested at mid-log phase (5 105 cells/ml). 2. 40 ml dounce homogenizer with type B pestle. 3. Histone purification buffer I: 10 mM MES pH 6.5, 0.5 mM sodium metabisulfite, 0.5 mM benzamidine–HCl, 5 mM MgCl2, 1 mM CaCl2, 15 mM NaCl, 60 mM KCl, 0.25 M sucrose, 0.5% Triton X-100, 0.1 mM PMSF, 0.5 mM DTT. 4. Histone purification buffer II: 10 mM PIPES pH 6.5, 0.5 mM sodium metabisulfite, 0.5 mM benzamidine–HCl, 5 mM MgCl2, 1 mM CaCl2, 0.1 mM PMSF, 0.5 mM DTT. 5. Nuclease, micrococcal (MNase, Worthington Biochemical), dissolve at 10 units/ml concentration in 50 mM Tris–Cl pH 7.9, 5 mM CaCl2, aliquots are stored at –80C. 6. 0.5 M EDTA pH 8.0.

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7. 10% SDS. 8. 4 M NaCl. 9. Histone purification buffer III: 1 mM EDTA pH 8.0, 0.2 mM PMSF. 10. SG gradient makers (Hoefer SG100, GE Healthcare). 11. Sucrose gradient buffer: 5 or 30% (w/v) sucrose in 10 mM Tris–HCl pH 7.5, 1 mM EDTA, 0.75 M NaCl, 0.3 mM PMSF, stored at 4C. 12. 25 % TCA solution (w/v), stored at 4C. 13. Acetone, pre-chilled and stored at –20C. 14. 5 SDS loading buffer: 0.25 M Tris–HCl pH 6.8, 0.5 M DTT, 10% SDS, 0.25% bromophenol blue, 50% glycerol. 15. Dialysis tubing with a 3,500 Da molecular weight cut-off. 16. Amicon Ultra-15 Centrifugal Filter Devices (Millipore). 17. Histone storage buffer: 10 mM HEPES–KOH pH 7.5, 10 mM KCl, 1 mM EDTA, 0.2 mM PMSF, 10% glycerol. 18. Hydroxyapatite Bio-Gel1 HT Gel (BioRad). 19. Histone purification buffer IV: 40 mM Na2HPO4 pH 6.8, 1 mM DTT, 0.2 mM PMSF. 2.2. In Vitro Histone Methyltransferase Assays

1. 5 HMTase buffer: 100 mM Tris–HCl pH 8.0, 20 mM EDTA, 2.5 mM DTT. 2. S-Adenosyl-L-[methyl-3H]methionine (Perkin–Elmer, 5–15 Ci/ mmol). 3. Histone methyltransferases: active HMTases can be purified as endogenous proteins or complexes, purified recombinant proteins or reconstituted complexes from insect cell or purified recombinant proteins from bacteria as fusion proteins with GST (glutathione S-transferase), His-tag, or CBP (calmodulinbinding protein) tag (See Note 1). 4. EN3HANCE or ENLIGHTENING gel autoradiography enhancer (Perkin–Elmer). 5. BioRad Model 583 Gel Dryer. 6. Kodak BioMax MR X-ray film.

2.3. Histone Demethylase Assays

1. Buffer C: 40 mM HEPES–KOH pH 7.9, 50 mM KCl, 0.2 mM PMSF, 1 mM DTT, 10% glycerol. 2. 5 Histone demethylase buffer type I: 250 mM HEPES– KOH pH 8.0, 70 mM Fe(NH4)2(SO4)2, 5 mM alphaketoglutarate, 10 mM ascorbate acid. (See Notes 2, 12). 3. 5 Histone demethylase buffer type II: 0.5 M glycine pH 8.0, 250 mM KCl. (See Note 3).


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4. 25% TCA solution (w/v), stored at 4C. 5. NASH conversion buffer: 3.89 M ammonium acetate, 0.1 M acetic acid, 0.2 % 2,4-pentandione, stored at 4C. 6. 1-Pentanol (Sigma). 7. Scintillation cocktail (EcoscintTM H, National Diagnostics). 8. Site-specific methyl-histone antibodies (Abcam Inc or Upstate Inc.). 9. 4% paraformaldehyde (w/v) in PBS, freshly made. 10. Blocking buffer: 1% BSA in PBS. 11. Secondary antibodies: donkey anti-mouse, rabbit or chicken, conjugated with rhodamine or FITC (Jackson ImmunoResearch Laboratories, Inc). 12. 4,6-Diamidino-2-phenylindole dihydrochloride (DAPI, Sigma), dissolved in water at 1 mg/ml as stock solution (1,000 ). 13. Fluorescent mounting medium (Dako Cytomation).

3. Methods 3.1. Histone Octamer and Nucleosomes Purification from HeLa-S3 Cells

Suspension-cultured HeLa-S3 cells are good source to generate native histones for various applications. Both oligo- and mononucleosomes can be obtained from particular MNase digested nuclei and purified through following sucrose gradient ultracentrifugation. Core histone octamers can be further purified by hydroxyapatite chromatography. The following purification protocol is derived from previous published methods (17–19) and can be used for purification of native histone substrates from HeLa-S3 cells for HMTase assays and other purposes (see Note 4).

3.1.1. Nuclei Preparation and Digestion

1. Resuspend HeLa-S3 cell pellet (1 1010 cells) with 100 ml of histone purification buffer I and homogenize with dounce homogenizer (pestle B) for about 10 strokes (see Note 5). 2. Pellet nuclei by centrifugation for 10 min at 4,000 rpm (Beckman SX4750) at 4C. Wash one time with same buffer and resuspend in 12 ml of histone purification buffer II. 3. First, perform a test to optimize MNase digestion condition. Take out 500 ml nuclei suspension and warm up to 37C followed by adding CaCl2 to 5 mM final concentration and 50 units of MNase stock. Carry out the digestion at 37C and transfer 50 ml digested nuclei every 5 min (up to 45 min) into a micro tube containing 1 ml 0.5 M EDTA to stop the reaction.

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Add to each tube in order 75 ml H2O, 30 ml 10 % SDS, 25 ml 4 M NaCl and finally 200 ml phenol/chloroform with vortexing at each step. Extract DNA by centrifugation for 10 min on a microfuge and analyze 5–10 ml of aqueous phase from each time point on a 1 % agarose gel to determinate optimal digestion time, which gives majority of products around 2 kb of DNA size. 4. Warm up the bulk of nuclei for 10 min at 37C and measure the total volume. Add CaCl2 to 5 mM final concentration and 10 ml MNase stock (100 units) per 1 ml of nuclei suspension. 5. Proceed the digestion for determined amount of time. And stop the digestion by adding 0.5 M EDTA to final concentration of 4 mM. 6. Centrifuge for 5 min at 4C at 5,000 rpm (Sorvall SS-34) and save the supernatant as S1. Measure the A260 units by adding 10 ml sample into 1 ml of 1 M NaOH. 7. Resuspend the pellet with equal volume of histone purification buffer III. Incubate on ice for 15–30 min with extensive vortexing at every 5–10 min to extract chromatin. 8. Centrifuge for 15 min at 10,000 rpm (Sorvall SS-34) at 4C and save the supernatant as S2. Measure the A260 units as step 7. Repeat this extraction step if the A260 units are low in both S1 and S2. 9. Combine DNA enriched samples and add the 5 M NaCl dropwise to the final concentration of 0.75 M with vortexing to remove histone H1 (see Note 6) 3.1.2. Nucleosomes Purification by Sucrose Gradient

1. Prepare 35 ml linear sucrose gradients (5–30%) in 40 ml polyallomer tubes (Beckman) using a SG gradient maker with 5 and 30% sucrose gradient buffer. 2. Gently layer 4–5 ml of digested nuclei samples (up to 300 A260 units) to each gradient and balance gradients well. 3. Centrifuge for 16 h at 26,000 rpm (Beckman SW-28) at 4C. 4. Carefully fractionate gradients by manually siphoning 1 ml of fractions from top to bottom of each gradient. 5. Transfer 50 ml samples from each fraction to microtubes with 100 ml of 25% TCA and precipitate histones for 30 min on ice. Spin for 15 min at full speed at 4C. 6. Wash the pellet with 0.5 ml of cold acetone and centrifuge again. Remove the supernatant and air dry the protein pellet. 7. Dissolve the pellet with 30 ml 1 SDS loading buffer and analyze the purity of histone proteins by 18% SDS-PAGE. 8. Transfer another 50 ml samples from each fraction to determine the size of DNA (Step 3, Section 3.1.1.)


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9. Combine fractions with good purity of histone proteins and about 1–2.5 Kb of DNA size as oligo nucleosomes. Dialyze against histone storage buffer. (see Notes 7, 8) 10. Combine fractions with good purity of histone proteins and more than 2.5 kb size of DNA. Dialyze against Tris–Cl (pH 8.0) with 1 mM PMSF to remove EDTA and sucrose and concentrate by ultrafiltration (Ultrafree-15, Millipore). Mono-nucleosome can be purified with the complete MNase digestion followed by same sucrose gradient protocol as discussed above (see Notes 7–9). 3.1.3. Core Histone Octamers Purification by Chromatography

1. Combine all other histone-containing fractions and dialyze against TE buffer with 1 mM PMSF. 2. Prepare a certain volume of hydroxyapatite (BioRad) column based on total amount of proteins. Equilibrate the column with histone purification buffer IV with 300 mM NaCl. 3. Load the sample onto the hydroxyapatite column and wash the column with 6–8 column volumes of histone purification buffer IV with 500 mM NaCl. 4. Elute the histone proteins with histone purification buffer IV containing 2.5 M NaCl. Collect 0.5–1 ml elution in each faction. 5. Precipitate 5–10 ml samples from each fraction with 25% TCA and determine the purity of proteins and the ratio of different core histone proteins by 18% SDS-PAGE. Combine right fractions as histone octamers and dialyze against histone storage buffer. (see Notes 7, 9)

3.2. In Vitro Methylation of Histones

HMTase assays are adapted from a published protocol (20) and essentially carried out as described previously (18, 21). Various recombinant histone methyltransferases can be purified through standard recombinant protein purification protocols and incubated with their preferred histone substrate together with S-adenosyl-L-[methyl-3H]-methionine serving as the methyl donor. Methylated histones generated from following assays can be used as substrates for the following histone demethylase assays we will discuss later. Because of the variation among different batches of HMTases and histones, optimizing the HMTase/ substrate ratio is strongly suggested. 1. Incubate 3–5 mg core histone octamers or nucleosomes (visible in coomassie) and 1 ml S-adenosyl-L-[methyl-3H]-methionine, 1/5 volume of 5 HMT buffer with different amount of purified histone methyltransferases at 30C for 1 h. 2. Stop the reaction by adding 1/5 volume of 5 SDS loading buffer and analyze reactions by 18% SDS-PAGE. Stain the gel with Coomassie blue R-250 for 30 min and destain it very well.

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3. Treat the gel with gel autoradiography enhancer for 30–45 min with gentle shaking and wash it in deionized water for 5–10 min. 4. Dry the gel on the filter paper for 2.5 h at 70C with gel dryer. 5. Expose the gel to an X-ray film with intensifying screens for overnight at –80C. Develop the film and determine the optimized HMTase/histones ratio based on the activity. 6. Scale up the reaction 10- to 50-fold under optimized condition. Incubate reactions for 1–2 h at 30C. 7. Transfer the reaction into a dialysis tubing and dialyze against 4 l histone storage buffer in the absence of EDTA for overnight at 4C to remove free S-adenosyl-L-[methyl-3H]-methionine. 8. Analyze 1 ml sample by liquid scintillation counting. The radioactive-labeled histones can be stored at 4C and applied for histone demethylase assays. (see Note 10). 3.3. Histone Demethylase Assays

So far two major families of proteins, LSD1 and Jmjc domaincontaining proteins, have been identified to demethylate histones in the presence of different cofactors. Through two different oxidation reactions, these demethylases can remove methyl groups from histone proteins and release formaldehyde. The following two in vitro assays apply different reaction conditions for different families of histone demethylases. The type I reaction buffer is optimized for the Jmjc domain-containing histone demethylases which contains Fe(II), alpha-ketoglutarate and ascorbic acid as cofactors, whereas the type II reaction buffer is for LSD1 family of histone demethylases. We also describe different detection methods for these assays. The in vitro radioactive formaldehyde release assay is one of the most sensitive assays, but the methylation sites and methylation states of the radioactive substrates are limited by availability of HMTases. Since all sitespecific methyl-histone antibodies are commercial available, the Western blot based detection method can be employed on every methylation state but it has lower sensitivity. Besides, the transfection-based immunostaining method has been widely used in most of histone demethylase studies (11, 13, 22–34).

3.3.1. In Vitro Formaldehyde Release Assays

1. Dialyze recombinant histone demethylases or protein fractions against 4 l buffer C. 2. Incubate [methyl-3H]-histone octamers or nucleosome substrates (about 10,000 cpms, see Section 3.2) and enzyme samples with 1/5 volume of 5 histone demethylase buffer I or II for 1–2 h at 37C (see Notes 11–13). 3. Add water into reaction to bring the total volume to 100 ml. 4. Add 100 ml 25 % TCA into reactions and precipitate proteins for 20 min on ice.


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5. Centrifuge for 10 min at 4C with full speed and transfer 180 ml supernatant to microtubes with 200 ml NASH conversion buffer. 6. Incubate the conversion reactions for 50 min at 37C. 7. Add 0.4 ml 1-pentanol and vortex well. Centrifuge for 5 min at room temperature. 8. Transfer 350 ml organic phase to scintillation counting vials with 1 ml scintillation cocktail. Mix thoroughly by inverting the vials more than five times and measure counts the scintillation counter. 3.3.2. Western BlotBased Histone Demethylase Assays

1. Dialyze recombinant histone demethylases or protein fractions and purified native histone proteins against buffer C. 2. Incubate 1–5 mg purified native histone substrates (see Section 3.1) and enzyme samples with 1/5 volume of 5 histone demethylase buffer I or II for 1 h at 37C (see Notes 11–14). 3. Stop the reaction by adding 1/5 volume of 5 SDS loading buffer and analyze reactions by 18% SDS-PAGE. 4. Transfer proteins onto a nitrocellulose membrane and probe with site-specific methylated histone antibodies using standard Western blot procedures. Anti-histone H3 or H4 antibodies should be included as the loading control.

3.3.3. ImmunostainingBased Histone Demethylase Assays

1. Plate 293T or NIH 3T3 cells onto 0.2% gelatin (Sigma)coated glass coverslips in 12-well plates 1 day before the transfections (see Note 15). 2. Transfect cells with expression plasmids (Flag, HA, myc or GFP tagged) for enzyme candidates and control plasmids using appropriate transfection reagent. Wait for 24–48 h. 3. Wash cells once with PBS and fix cells for 20 min in 4% paraformaldehyde solution (see Note 16). 4. Permeabilize cells for 5 min with cold PBS buffer containing 0.5% Triton X-100. 5. Wash cells with blocking buffer for 3 5 min and block the coverslips for 30 min. 6. Incubated coverslips with primary antibodies for 1 h in a humidified chamber with gentle rocking motion. 7. Wash cells with blocking buffer for 3 5 min and incubate with fluorescent-labeled secondary antibodies for 1 h with gentle rocking motion. 8. Wash cells with PBS for 2 5 min and stained with DAPI. 9. Wash cells 3 5 min with PBS and 1 5 min with water and mount coverslips on slides. Dry the slides in the dark place for overnight and examine under fluorescent microscope.

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4. Notes 1. For histone lysine methylation, following recombinant HMTases can be used to generate different [methyl-3H]histone substrates for histone demethylase assays: GSTSET7/9 (H3K4me1, core histone octamers), GST-SET7/ 9-Y245A (H3K4me2&3, core histone octamers), GST-G9a (621-1000aa) (H3K9me1 and H3K9me2, core histone octamers), GST-SUV39H1(82-412)-H320R (H3K9me, core histone octamers), GST-Dim5 (N. crassa, H3K9me3, core histone octamers), Ezh2 complex (H3K79me, nucleosomes), CBP-Set2-Flag (S. pombe, H3K36me2, nucleosomes), GST-DOT1L (1-416) (H3K79me, nucleosomes), GST-SET8 (H4K20me1, core histone octamers and nucleosomes), GST-Suv4-20h1 and GST-Suv4-20h2 (H4K20me2&3, nucleosomes). 2. Type I histone demethylase buffer should be made freshly from 1 M HEPES–KOH (pH8.0), 0.5 M Fe(NH4)2(SO4)2, 0.5 M alpha-ketoglutarate and 0.5 M ascorbate acid stock solutions. Aliquot of stock solutions of cofactors should be stored at –20C. 3. Alternatively, another reaction buffer was initially reported for LSD1 family of histone demethylases: 50 mM Tris–HCl pH 8.5, 50 mM KCl, 5 mM MgCl, 0.5% BSA, 5% glycerol (8). 4. Core histone octamers and nucleosomes can be purified from chicken blood red cells following same protocols. 5. Generation of nuclei should be verified under microscope. 6. This step is to remove histone H1 from nucleosomes. Omit this step and remove 0.75 M NaCl in the sucrose gradient buffer if histone H1 is desired. Alternatively, 0.75 M NaCl can be included directly in histone purification buffer III during extraction steps (step 7–8) to simplify the procedure if histone H1 is not required. 7. Dialyze to histone storage buffer in the absence of 1 mM EDTA if histones will be directly applied to histone demethylase assays under type I reaction condition. 8. Purified nucleosomes can be stored for up to 6 months at 4C and for longer at –80C. 9. A couple of top gradient fractions contain mononucleosomes, however, these fractions are not recommended for HMTase assays directly because of impurity. 10. 3H-labeled methyl-histone octamers or nucleosomes should be stored at 4C for up to 1 month. Avoid freeze-thaw steps.


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Longer storage could result in higher background for the histone demethylase assays for some unknown reasons. 11. Avoid EDTA in type I histone demethylase reactions. EDTA does not affect the type II histone demethylase reactions. 12. 70 mM Fe(NH4)2(SO4)2 in 5 type I histone demethylase buffer is suitable for most of cases, however, titration of Fe(NH4)2(SO4)2 concentration is recommended to get optimized assay condition for different histone demethylases. 13. 10 mM MgCl2 is included in a similar type I reaction buffer for the core histone octamer substrates, but not for the nucleosome substrates (22–24). 14. Commercial or customer synthesized site-specific methylhistone peptides can be also applied as substrates using Western blot-based detection method. And mass spectrometry can be employed to detect reduction of histone peptide masses which corresponds to methyl groups if peptide substrates are applied (11, 12, 22, 23, 26–34). 15. Other cell lines have been also used for the histone demethylase assays including HeLa, U2OS, COS-7, iMEF, etc. 16. Alternatively, cells can be pre-extracted with Triton X-100 prior to fixation (35).

Acknowledgments Work in the Fang lab is supported by funds from the American Cancer Society Institutional Research Grant (ACS-IRG) Program and H. Lee Moffitt Cancer Center & Research Institute.

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Chapter 18 Histone Chaperone as Coactivator of Chromatin Transcription: Role of Acetylation Shrikanth S. Gadad, Jayasha Shandilya, Venkatesh Swaminathan, and Tapas K. Kundu Abstract Histone chaperones are a group of histone-interacting proteins, involved in several important cellular functions. These chaperones are essential to facilitate ordered assembly of nucleosomes, both in replication dependent and independent manner. Replication independent function of histone chaperone is necessary for histone eviction during transcriptional initiation and elongation. In this chapter we have discussed a method to evaluate the role of histone chaperone NPM1 (the only known chaperone to get acetylated with functional consequence) in the transcriptional activation which is acetylation dependent. Key words: NPM1, acetylation, nucleosomes, transcriptional co-activators, histone transfer assay.

1. Introduction In eukaryotes, the genome is organized into a highly compact nucleoprotein structure called chromatin, which in itself is repressive in nature. However, various cellular machineries act in conjunction to maintain the fluidity of the chromatin, mainly by post-translational modifications of histones and nonhistone chromatin-associated proteins as well as ATP-dependent chromatinremodeling complexes. Reversible post-translational modifications like acetylation–deacetylation and methylation–demethylation act as marks for recruiting various chromatin remodelers, which in turn make the chromatin template either permissive (H3K14, K18 acetylation/H3K4 methylation) or repressive (H4K16 The first three authors have contributed equally and Tapas K. Kundu is the corresponding author. Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_18 Springerprotocols.com

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deacetylation/H3K9 methylation) for facilitating various cellular processes like transcription, replication and repair. Increasing evidences point towards the obligatory requirement of a plethora of histone chaperones in transcriptional regulation. Histone chaperones are a group of histone-interacting proteins responsible for regulating diverse cellular functions like histone transport and storage (e.g., NAP1), replication-coupled histone deposition (e.g., CAF1), histone removal (e.g., Asf1), transcriptional coactivation (e.g., NPM1) and possibly stress response (1). With the recent advances in the histone chaperone field various assays have been developed to characterize proteins belonging to the histone chaperone family both in human and in yeast. The chaperoning activity of a protein can be monitored by histone transfer assay (HTA), wherein the chaperone mediated assembly of nucleosomes onto a relaxed DNA template is assessed by the introduction of negative supercoils (2). In vitro chromatin transcription assays have been modified to study the effect of histone chaperone on the remodeling events, which has been shown to facilitate transcription from a repressed chromatin in an acetylation dependent manner (3). Additionally, post-translational modifications of histones and the chaperone itself, mainly by acetylation can be studied in in vitro chromatin transcription assay by using specific inhibitors for HATs. Taken together these experimental approaches would be useful in the functional characterization of histone chaperone proteins. In this chapter, we shall describe the histone chaperone activity by histone transfer assay and the role of acetylation of histone chaperone in the regulation of chromatin transcription, taking NPM1 as a model histone chaperone.

2. Materials 2.1. CsCl-Purified pG5 ML Array

2.2. Purification of Core Catalytic Domain of Drosophila Topoisomerase I (dTopoI) (see Ref.(6))

The transcription template is composed of an adenovirus major late (ADML) core promoter in front of a 380 bp G-less cassette. Upstream of the core promoter are the activator (Gal4) binding sites, while the 50 and 30 flanking regions of this transcriptional cassette contains five repeats of the nucleosomal positioning sequence from the sea urchin 5S rRNA gene. (see Fig 18.1) (4). 1. BL21(DE3) competent cells 2. 100 ml Luria broth containing 50 mg/ml kanamycin 3. 1M IPTG 4. Lysis buffer: 50 mM sodium phosphate pH 7.0, 0.5 M NaCl, 15% glycerol, 15 mM imidazole, 0.1% (v/v) NP-40, 0.2 mM PMSF and 0.5 mM benzamidine. 5. Ni-NTA His-Bind Resin (Novagen)

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[

[

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5 Gal4 Sites

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Fig. 18.1. pG5ML array template.

6. End-to-end shaker 7. Econo-Fast column (Biorad) 8. Elution buffer: lysis buffer containing 0.5 M imidazole 9. Dialysis buffer: 25 mM HEPES pH 7.6 and 0.1 mM EDTA, 10% glycerol, 50 mM NaCl, 0.01% Nonidet P-40, 0.5 mM DTT, 0.2 mM PMSF and 0.5 mM benzamidine 2.3. Preparation of HeLa Nuclear Extract and Nuclear Pellet

1. HeLa S3 cell line 2. Spinner flasks 3. Joklik’s medium (Sigma) supplemented with L-glutamine (Sigma), 1 penicillin–streptomycin–amphotericin B solution and 10% fetal bovine serum 4. Hypotonic buffer: 10 mM Tris pH 7.5, 10 mM KCl, 1.5 mM MgCl2, 0.2 mM PMSF and 10 mM -mercaptoethanol 5. Dounce’s homogenizer (Wheaton) with pestle B 6. 100 mM NaCl and 5% glycerol 7. Low salt buffer: 20 mM Tris pH 7.5, 25% glycerol, 1.5 mM MgCl2, 0.2 mM EDTA, 20 mM KCl, 0.2 mM PMSF and 10 mM -mercaptoethanol 8. TGME buffer: 50 mM Tris–HCl pH 7.9, 25% glycerol, 5mM MgCl2, 0.1 mM EDTA pH 8.0, 0.2 mM PMSF and 10 mM -mercaptoethanol. 9. High salt buffer: 20 mM Tris–HCl pH 7.5, 25% glycerol, 1.5 mM MgCl2, 0.2 mM EDTA, 1.2 M KCl, 0.2 mM PMSF and 10 mM -mercaptoethanol 10. BC100: 1 M HEPES pH 7.9, 100 mM KCl and 20% glycerol, 0.2 mM EDTA pH 8.0

2.4. Purification of Core Histones

1. HeLa nuclear pellet 2. Buffer A: 100 mM potassium phosphate buffer pH 6.7, 0.1 mM EDTA, 10% glycerol, 0.1 mM PMSF, 0.1 mM DTT and 630 mM NaCl 3. Dounce’s homogenizer (Wheaton) with pestle B

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4. Hydroxyapatite BioGel HTP (Biorad) 5. 10 mM potassium phosphate, pH 6.7 6. Econo-Fast glass column 7. Buffer B: same as buffer A with 2 M NaCl 8. BC100: 20 mM Tris–HCl pH 7.4, 20% glycerol, 2 mM -mercaptoethanol, 0.2 mM EDTA, 2 mM PMSF, 0.1% nonidet P-40 (IGEPAL CA-630) and 100 mM KCl. 9. 6.6 mM trichostatin A 10. 100 mM sodium butyrate 2.5. Purification of Human NPM1, Mouse NAP1

1. BL21 (DE3) 2. 100 ml Luria broth with 50 mg/ml kanamycin (for NPM1) or 100 mg/ml ampicillin (for mNAP1) 3. 1 M IPTG 4. Lysis buffer: 20 mM Tris–HCl pH 7.4, 20% glycerol, 2 mM mercaptoethanol, 0.2 mM EDTA, 2 mM PMSF, 0.1% nonidet P-40, 30 mM imidazole and 300 mM KCl 5. Ni-NTA His-Bind Resin (Novagen) 6. End-to-end shaker 7. Wash buffer: same as lysis buffer with 300 mM KCl 8. Econo-Fast column (Biorad) 9. Elution buffer: same as lysis buffer in addition contains 250 mM imidazole and 100 mM KCl.

2.6. Reagents for In vitro Interaction Studies

1. NPM1 or acetylated NPM1 2. Core histones or acetylated core histones 3. Ni-NTA His-Bind Resin (Novagen) 4. Interaction buffer: 20 mM Tris–HCl pH 7.9, 20% glycerol, 0.2 mM EDTA pH 8.0, 0.1% Nonidet P-40, 2 mM PMSF, 300 mM KCl and 30 mM imidazole (see Note 1) 5. 2 SDS loading buffer: 100 mM Tris–HCl pH 6.8, 4% (w/ v) SDS electrophoresis grade, 0.2% (w/v) bromophenol blue, 20% glycerol and 200 mM -mercaptoethanol.

2.7. Histone Transfer Assay

1. pG5ML plasmid (see Note 2). 2. Core histones or acetylated core histones 3. dTopo I (catalytic domain) 4. NPM1 or acetylated NPM1 5. 2 assembly buffer: 10 mM Tris–HCl pH 8.0, 1 mM EDTA, 200 mM NaCl and 0.1 mg/ml bovine serum albumin (Sigma). (see Note 1)


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6. Proteinase K stop buffer: 200 mM NaCl, 20 mM EDTA pH 8.0, 1% SDS, 0.25 mg/ml glycogen and 0.125 mg/ml proteinase K 7. Agarose gel (1%) 8. 6 sample buffer: 0.25% (w/v) bromophenol blue, 0.25% (w/v) xylene cyanol, 30% (v/v) glycerol in water. 2.8. Buffers and Reagents for In Vitro Transcription Assay

1. BC 0: 20 mM Tris–HCl, pH 7.9, 20% glycerol, 0.2 mM EDTA pH 8.0. 2. BC 200: 20 mM Tris–HCl pH 7.9, 20% glycerol, 0.2 mM EDTA pH 8.0, 200 mM KCl. 3. 20 buffer: 0.4 M HEPES pH 8.4 and 100 mM MgCl2. 4. 2 HAT buffer: 20 mM HEPES, pH 7.8, 100 mM KCl, 10 mM DTT, 1 mM PMSF, 40 mM sodium butyrate, 0.5 mg/ml BSA and 10% glycerol. 5. 20 NTP mix: 12 mM ATP, 12 mM CTP, 0.5 mM UTP, and 2 mM 30 O-methylGTP (Amersham). 6. 10 mCi/ml 32pUTP (Board of Radiation and Isotope Technology, DAE, Govt. of India). 7. Transcription loading dye: 8.0 M urea in 1 TBE, xylene cynol and 0.005% bromophenol blue. 8. 2 transcription assembly buffer: 20 mM Tris–HCl pH 8.0, 2 mM EDTA, 300 mM NaCl and 0.2 mg/ml BSA.

2.9. Buffers and Reagents for MNase Assay

1. R buffer: 10 mM HEPES pH 7.5, 10 mM KCl, 1.5 mM MgCl2, 0.5 mM EDTA, 10% glycerol, 10 mM -glycerophosphate, 1 mM DTT and 0.2 mM PMSF. 2. MNase stop solution: 0.5 TE, 0.25 M EDTA 3. Proteinase K stop buffer: 200 mM NaCl, 20 mM EDTA pH 8.0, 1% SDS, 0.25 mg/ml glycogen and 0.125 mg/ml proteinase K

3. Methods 3.1. Purification of pG5ML

DNA templates for in vitro transcription and histone transfer assays are purified by plasmid maxi prep kit as recommended by the manufacturers (Qiagen) followed by CsCl equilibrium gradient purification (5).

3.2. Purification of dTopoI

His-6 tagged core catalytic domain of the Drosophila topoisomerase I is purified using the Ni-NTA column(6).

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1. Transform BL21 (DE3) cells with the expression vector and grow overnight in 100 ml LB medium containing 50 mg/ml kanamycin. 2. Inoculate the overnight culture into 1 l of LB medium containing 50 mg of kanamycin and grow at 37C, till the OD600 reaches 0.5. 3. Induce the culture with IPTG to a final concentration of 0.42 mM and grow at 30C for 5 h. 4. Harvest the cells by centrifugation at 5000 rpm (2980 g) for 10 min at 4C. 5. Resuspend the pellet in lysis buffer and sonicate with four burst of 30 s at a setting of 32% amplitude (Sonics-Vibra cell). 6. Clarify the lysate by centrifuging at 16,000 rpm (30,597 g) for 30 min at 4C. 7. Mix the supernatant with Ni-NTA beads (pre-equilibrated with the lysis buffer) and keep on an end-to-end shaker at 4C for 3 h. 8. Wash the resin five times with the lysis buffer and pack it onto a column. 9. Elute the protein with the elution buffer (lysis buffer containing 0.5 M imidazole) 10. Dialyze the purified protein in dialysis buffer (see Subheading 2.2) 3.3. Preparation of HeLa Nuclear Extract and Nuclear Pellet

HeLa nuclear extract is prepared from HeLa S3 cell line cultured in spinner flasks (7). 1. Grow the HeLa S3 cells in the Joklik’s medium (Sigma) supplemented with L-glutamine (Sigma), 1 penicillin– streptomycin–amphotericin B solution and 10% fetal bovine serum (FBS). 2. Pellet the cells at 3000 rpm (1800 g) for 10 min at 4C. 3. Wash the pellet with ice-cold 1 PBS containing 0.2 mM PMSF and 10 mM -mercaptoethanol. 4. Determine the packed cell volume (PCV). 5. Wash the cells with one PCV volume of the hypotonic buffer and pellet down at 2500 rpm (1250 g) for 5 min at 4C. 6. Decant the supernatant carefully and resuspend the cells in 2 PCV volumes of the hypotonic buffer and incubate in ice for 15 min. 7. After the incubation, homogenize the cell suspension by 15 strokes with the pestle B in a Dounce’s homogenizer (Wheaton) for 15 min. 8. Clarify the homogenate by centrifuging at 4000 rpm (1910 g ) for 15 min at 4C.


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9. The resultant supernatant is the cytoplasmic extract (S100 extract), which is stored in a buffer composed of 100 mM NaCl in 5% glycerol at –80C. 10. The pellet volume will be recorded as the nuclear pellet volume (NPV). 11. Process the nuclear pellet for nuclear extract preparation. 12. Homogenize the nuclear pellet with half the NPV volume of low salt buffer. 13. Pour the homogenate into a small beaker with a magnetic bead and add an equal volume of high salt buffer dropwise and stir continuously for 30 min. 14. Clarify the resultant homogenate by centrifugation at 14,000 rpm (23,360 g) for 20 min at 4C. 15. Dialyze the supernatant against BC100 for 3 h and centrifuge at 14,000 rpm (23,360 g) for 20 min at 4C. The supernatant will be the HeLa nuclear extract. The pellet in the previous step is the nuclear pellet which is homogenized in TGME buffer and is subsequently used for histone purification. 3.4. Purification of Core Histones and Hyperacetylated Core Histones

Human core histones are purified from the HeLa nuclear pellet (7). 1. Resuspend 5 ml HeLa nuclear pellet in 15 ml buffer A. 2. Homogenize the suspension in a dounce’s homogenizer (Wheaton) with Pestle B for 30 min on ice. 3. Clarify the suspension at 14,000 rpm (23,360 g) for 20 min at 4C. 4. Transfer the supernatant to a fresh tube and add 5 gm (per 2.4 of mg DNA) Hydroxyapatite BioGel HTP (Biorad) which is presoaked in 15 ml 10 mM potassium phosphate buffer pH 6.7. 5. Incubate this at 4C for 3 h on an end-to-end shaker. 6. Wash the resin twice with 25 ml of buffer A with 630 mM NaCl on a centrifuge. 7. Pack the hydroxyapatite into a column and wash overnight with 500 ml buffer A containing 630 mM NaCl (see Note 3). 8. Elute the core histones in buffer B containing 2 M NaCl. Collect 20 fractions of 0.5 ml each. 9. To detect the presence of histones in the collected fractions, 20 ml of eluted protein is precipitated with 25% trichloro acetic acid by keeping on ice for 30 min. Centrifuge at 13,000 rpm (15,700 g) at 4C for 30 min. 10. Wash once with ice-cold acetone and centrifuge again at 13,000 rpm (15,700 g) at 4C for 30 min.

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11. Pipette out the supernatant and dry the pellet in speed vac. 12. Dissolve the pellet in 5 ml of 1 SDS loading buffer and heat at 90C for 10 min. Resolve the histones on a 15% SDS–PAGE. Visualize the bands by staining with Coomassie brilliant blue. For purification of hyperacetylated core histones, the same procedure is followed, except the starting material will be HeLa cells treated with the histone deacetylase inhibitors, 150 nM trichostatin A and 5 mM sodium butyrate for 24 h (3). 13. Core histones and hyperacetylated core histones are dialyzed in BC100 (see Subheading 2.4). 3.5. Purification of NPM1 and mNAP1

His-6-tagged recombinant NPM1 or mNAP1 is purified using the Ni-NTA column (3). 1. Transform the BL21 (DE3) cells with either NPM1 or mNAP1expression vector and grow overnight in 100 ml LB medium containing 50 mg/ml kanamycin (for NPM1) or 100 mg/ml ampicillin (for mNAP1). 2. Inoculate the overnight culture into 1 l of LB medium containing appropriate antibiotics and grow at 37C till the OD600 reached 0.5. 3. Induce the culture with IPTG (0.5 mM) and grow at 37C for 3 h. 4. Harvest the cells by centrifugation at 5000 rpm (2980 g) for 10 min at 4C. 5. Resuspend the pellet in lysis buffer and sonicate with four burst of 30 s at a setting of 32% amplitude (Sonics-Vibra cell). 6. Clarify the lysate by centrifuging at 16,000 rpm (30,597 g) for 30 min at 4C. 7. Mix the supernatant with Ni-NTA beads pre-equilibrated with the lysis buffer and keep on an end-to-end shaker at 4C for 3 h. 8. Wash the resin five times with the wash buffer and pack onto an Econo-Fast column. 9. Elute the protein with the elution buffer (see Subheading 2.5). 10. The purified His-6-tagged recombinant NPM1 and mNAP1 are finally dialyzed in BC100 without imidazole before storing as small aliquots in –80C.

3.6. In Vitro Interaction of NPM1 or Acetylated NPM1 with Core Histones or Acetylated Core Histones

Interactions are carried out by in vitro Ni-NTA pulldown assay: 1. Mix 1 mg of the His-6-tagged protein (full length NPM1 or acetylated NPM1) with 4.0 mg core histones or acetylated core histones in the interaction buffer along with the Ni-NTA His-Bind Resin.(Reaction volume: 300 ml). 2. Incubate the mixture for 2 h at 4C on an end-to-end shaker.


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3. Extensively wash the beads three times with 1 ml interaction buffer. 4. Resolve the bead bound proteins on a 12% SDS-PAGE (Fig. 18.2A and B). In order to achieve better sensitivity of this assay, the bound histone could also be subjected to Western blotting analysis using highly specific antibodies against individual histones (see Note 4).

Fig. 18.2. Effect of acetylation on NPM1 and acetylated histone interaction. (A) In vitro interaction of NPM1 with HeLa core histones with increasing salt concentration: Ni-NTA pulldown assays with bead alone (lane 1); (lanes 2–4) 1 mg His-6 NPM1 incubated with 4 mg of HeLa core histones in presence of 300 mM, 500 mM and 850 mM KCl concentration in the interaction buffer, respectively. The top and bottom panel is the Coomassie blue-stained gel showing the amounts of the pulled down proteins. (B) Indicated amounts of hyperacetylated core histones purified from the HeLa nuclear pellet were tested for their interaction with 500 ng of either NPM1 (lanes 3, 4 and 5) or acetylated NPM1 (lanes 6, 7 and 8) using the Ni-NTA pulldown assay. Ni-NTA beads alone, without NPM1 or acetylated NPM1, was used as a negative control (lane 2). Two micrograms of the sample was loaded as input (lane 1). The top panel is the Coomassie blue-stained gel showing the amounts of the pulled down proteins, while the bottom panel shows the Western blot with antibodies against acetylated histone H3.(Adapted from ref. (3) with permission of American Society for Microbiology Journals, Washington, DC).

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Fig. 18.3. Acetylated NPM1 transfers histones more efficiently than NPM1. Histone transfer was tested with 500 ng of core histones and either 5 pmol of NPM1 (lane 7) or the acetylated NPM1 (lane 8). Also with 500 ng of hyperacetylated core histones with either NPM1 (lane 9) or acetylated NPM1 (lane 10). Lane 1, supercoiled DNA; lane 2, DNA relaxed by TopoI; lanes 3 and 4, relaxed DNA along with core histones and acetylated core histones, respectively. NPM1 (lane 5) or acetylated NPM1 alone (lane 6).

3.7. Histone Transfer Assay

The nucleosome assembly activity of NPM1 and acetylated NPM1 is assayed as depicted in (Fig. 18.3) (8). 1. Relax 200 ng of covalently closed circular plasmid pG5ML by pre-incubating with 40 ng dTopoI core catalytic domain in the 1 assembly buffer at 37C for 40 min. 2. At the same time, separately incubate 5 pmol of the NPM1 or acetylated NPM1 and 500 ng of core histones or acetylated core histones in 1 assembly buffer at 37C for 40 min. 3. Mix the two reactions together and further incubate at 37C for 40 min. 4. Stop the reaction by the addition of the proteinase K stop buffer. 5. At the end of the reaction, extract the plasmid by adding equal volume of phenol:chloroform:isoamyl alcohol (25:24:1) pH 8.0 followed by precipitation with 100% ethanol. 6. Resolve the topoisomers on a 1% agarose gel (40 Volts for 12 h) and visualize by staining with ethidium bromide (see Note 5).

3.8. Template Relaxation Assay

1. Incubate 6 mg of CsCl-purified supercoiled pG5ML array with 40 ng dTopo1 in relaxation buffer at 37C for 2 h. 2. Extract the relaxed DNA with equal volume of phenol: chloroform:isoamyl alcohol (25:24:1) pH 8.0.


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Fig. 18.4. Efficient assembly of chromatin. (A) Relaxation assay of pG5ML array template in presence of dTopo1 (lane 2) supercoiled DNA (Lane 1). (B) MNase digestion pattern of in vitro assembled chromatin (lane 1). (R ¼ Relaxed, S ¼ Supercoiled).

3. Precipitate the DNA with two volume of 100% ethanol, 1/10 volume 3 M Sodium acetate at –80C for 1 h. 4. Dissolve in 36 ml of double-distilled water. 5. Visualize the relaxed DNA on 0.8% agarose gel (run at 50 V for 12 h). Stain with ethidium bromide (Fig. 18.4A). 3.9. Chromatin Assembly

1. 1.8 mg mNAP1 and 520 ng HeLa core histones are incubated in 1 transcription assembly buffer at 37C for 15 min (4) (see Note 6) 2. Add 168 ng of relaxed pG5ML array template into the reaction mixture and incubate at 37C for 45 min. The efficiency of chromatin assembly is assessed by micrococcal nuclease digestion pattern.

3.10. Micrococcal Nuclease Digestion of Assembled Chromatin

1. 336 ng of chromatin (two assembly reactions) is taken in a final volume of 50 ml. Add equal volume of R buffer (4). 2. Add CaCl2 to a final concentration of 3 mM and digest with 1.0 U MNase at room temperature for 10 min. 3. The reaction is stopped by addition of 5 ml of MNase stop solution. Incubate at room temperature for 15 min. 4. Add 150 ml proteinase K stop buffer, mix by a brief vortex and incubate at 37C for 15 min. 5. Add equal volume (250 ml) of phenol:chloroform:isoamyl alcohol (25:24:1) pH 8.0 to extract the DNA in the upper phase at room temperature. 6. Add 0.25 M ammonium acetate and three volumes of 100% ethanol to precipitate the DNA.

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7. Dissolve the pellet in 5 ml TE and 1 sample buffer. 8. Electrophorese on a 1.25 % agarose gel in 1 TBE buffer (run at 200 V at 4C). Stain with ethidium bromide (Fig. 18.4B). 3.11. Histone Acetylation Assay on Chromatin Template

1. Assemble 168 ng of chromatin as mentioned in (see Subheading 3.9). 2. Incubate the chromatin in 1 HAT buffer and 180 ng Gal4-VP16 at 30C for 20 min. 3. Add 120 ng p300 and 1 ml of 2.5 Ci/mmol 3H-acetyl CoA. Incubate at 30C for 15 min. 4. Precipitate the reaction products with 25% trichloro acetic acid by keeping on ice for 30 min. Centrifuge at 13,000 rpm (15,700 g) at 4C for 30 min 5. Wash once with ice-cold acetone and centrifuge at 13,000 rpm (15,700 g) at 4C for 30 min 6. Pipette out the supernatant and dry the pellet in speed vac. 7. Dissolve the pellet in 10 ml of 1 SDS loading buffer and heat at 90C for 10 min. Resolve on 12% SDS polyacrylamide gel. Visualize the bands by staining with Coomassie brilliant blue. 8. Equilibrate the gel in Me2SO for 30 min twice. 9. Perform fluorography using a solution containing 22.5% 2, 5-diphenyoxazole in Me2SO for 45 min. rinse the gel with double-distilled water for 2 h. 10. Vacuum dry the gel at 60C for 90 min, perform autoradiography at –70C for 3 days.

3.12. In vitro Transcription Assay (9)

1. 28 ng of DNA or equivalent amount of chromatinized template is incubated with 30 ng of Gal4-VP16 in 0.4 HAT buffer at 30C for 20 min. (see Note 7). 2. Add 20 ng of p300 and 1.5 mM acetyl CoA and incubate at 30C for 15 min. 3. Further acetylation reaction can be stopped by using specific small molecule inhibitor against p300 like (300 mM) curcumin (10), (10 nM) lysyl-CoA (3, 11) or (50 mM) LTK-14 (12) (Fig. 18.5A and B). 4. Add indicated amount of histone chaperone NPM1 to the reaction mix and incubate at 30C for 30 min for allowing any remodeling event to occur. (Fig. 18.6A and B). 5. Add 5 ml HeLa nuclear extract (8.0 mg/ml protein in BC100). Add 0.25 mM DTT in the reaction and incubate at room temperature for 20 min (see Note 7). 6. Add 0.5 ml RNaseOUT (Invitrogen), 2.5 ml of 20 NTP mix and 15mCi 32pUTP. Incubate at 30C for 45 min.


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Fig. 18.5. Acetylation of nucleophosmin is critical for transcriptional activation from a chromatin template. (A) Schematic representation of in vitro transcription protocol modified to include the interrupted acetylation step (addition of the p300specific inhibitor lysyl-CoA). (B) About 28 ng of freshly assembled chromatin was subjected to the protocol described for panel A without or with Gal4-VP16, full-length p300 or 1.5 mM acetyl-CoA and without or with the p300-specific HAT inhibitor lysyl-CoA, along with 30 pmol (each) of full-length NPM1 or acetylated NPM1 as indicated. The relative transcription per lane (in several fold activation over the respective control lanes marked with the number 1) was determined by phosphorimage analysis (Fuji). (Adapted from ref. (3) with permission of American Society for Microbiology Journals, Washington, DC).

7. Add 1 ml of (20 U/ml) RNaseT1 (Calbiochem) and incubate at 30C for 20 min. 8. Add 200 ml transcription stop buffer and immediately extract the radiolabeled RNA transcript with 250 ml phenol: chloroform (acid equilibrated pH 5.7) (see Note 8). 9. Centrifuge at 13,000 rpm (15,700 g) at room temperature for 15 min. 10. Carefully recover the upper phase (230 ml) and add 23 ml 3 M sodium acetate (pH 5.2) and add 650 ml 100 % ethanol (v/v).

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Fig. 18.6. Nucleophosmin stimulates acetylation-dependent chromatin transcription in a dose-dependent manner. (A) Schematic representation of the in vitro transcription protocol. (B) 28 ng of freshly assembled chromatin was subjected to the protocol described for panel A. Lane 1 without activator; (lanes 2–8) with 30 ng Gal4-VP16; lanes 3–8 with 20 ng p300; lanes 4–8 with 1.5 mM acetyl-CoA; lanes 5–8 in presence of 5, 10, 30 and 50 pmol of NPM1, respectively.

11. Mix briefly by vortexing and incubate at –80C for 1 h. 12. Centrifuge at 13,000 rpm (15,700 g) 4C for 30 min. Immediately dry the RNA pellet in speed vac. Dissolve the pellet in 10 ml transcription loading dye. 13. Run the radiolabeled transcripts on a 7 M urea–5% polyacrylamide gel in 1 TBE for 2 h at 300 V (see Note 9). 14. Rinse the gel in distilled water for 15 min to remove urea and free 32pUTP. Vacuum dry the gel at 80C for 40 min. 15. Autoradiograph the gel with an intensifying screen (expose for 12 h) and analyze on Fuji BAS phosphor imager system.

4. Notes 1. The optimal salt concentration for the assay has to be empirically determined. 2. For histone transfer assay any other plasmid apart from pG5ML can be used, and hence the presence of a strong nucleosomal


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positioning sequences can be avoided. However, in order to assemble the chromatin for acetylation dependent in vitro chromatin transcription, a strong nucleosome positioning is important, especially when salt dependent, NAP1-mediated (7) chromatin assembly will be used to reconstitute the chromatin. Nucleosome positioning is brought about by 5S rDNA sequences flanking the G-less cassette in pG5ML plasmid. 3. During the washing step, when almost 150 ml of buffer A has been passed through the column, it is observed that there is a drastic reduction in the flow rate. In order to maintain the flow rate of the wash step a peristaltic pump may be set up at the rate of 25 ml/h. However, it is necessary to keep the peristaltic pump connected thereafter, until all the washes and elution step of the protein is completed. 4. Western blot analysis in order to detect the core histones interaction with NPM1, highly specific polyclonal antibodies against each of the four core histone H3, H4, H2A and H2B were used which are raised in the laboratory (PCT/IN2005/ 000377). 5. For the histone transfer assay involving the acetylated NPM1, reduce the post-relaxation interaction time by 10 min in order to perceive the difference in the histone transfer by acetylated NPM1. 6. Ratio of mNAP1 and HeLa core histones should be standardized for proper assembly into chromatin. This method of assembly does not involve ATP-dependent remodeling. This template is amenable to further remodeling events by exogenously added chaperone. The chromatin template should be close to complete repression in absence of acetylation. Only from this kind of template several fold activation may be brought about by the histone chaperone. This step may need titration with different histone : DNA ratio for chromatin assembly. Chromatin assembly can also be done with reconstituted recombinant core octamer to rule out the effect of in vivo modifications present in HeLa purified core histones. Additionally chromatin assembled with reconstituted core octamer can be further modified by acetylation and methylation for in vitro chromatin transcription assay. 7. It is important to gently tap and give a brief spin after each step of transcription protocol. However, never spin down the transcription reaction mixture after addition of HeLa nuclear extract, mix by tapping gently 5–6 times. 8. Vigorously vortex the reaction mix for phenol–chloroform extraction of labeled RNA. 9. To prepare 7 M urea–5% polyacrylamide gel, slowly dissolve 21 gm of urea in 12.5 ml 2 TBE, 6.25 ml of 40% acrylamide and 6.25 ml double-distilled water. Finally make up the volume up to 50 ml with double-distilled water.

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Acknowledgments We thank R. G. Roeder and J. Kadonaga for some of the valuable reagents used in this work.

References 1. Shandilya, J., Gadad, S., Swaminathan, V., and Kundu, T. K. (2007) Histone chaperones in chromatin dynamics: Implication in disease manifestation. Chromatin and Disease, Subcell. Biochem. 41, Chapter 6, 111–124, Springer Press. 2. Nelson, T., Hsieh, T. S., and Brutlag, D. (1979) Extracts of Drosophila embryos mediate chromatin assembly in vitro. Proc. Natl. Acad. Sci. USA. 76, 5510–5514. 3. Swaminathan, V., Kishore, A. H., Febitha, K. K., and Kundu, T. K. (2005) Human histone chaperone nucleophosmin enhances acetylationdependent chromatin transcription. Mol. Cell. Biol. 25, 7534–45. 4. Kundu, T. K., Palhan, V. B., Wang, Z., An, W., Cole, P. A., and Roeder, R. G. (2000) Activator-dependent transcription from chromatin in vitro involving targeted histone acetylation by p300. Mol. Cell. 6, 551–561. 5. Sambrook, J., and Russell, D. W. (2001) Molecular Cloning : A Laboratory Manual. 3rd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. 6. Fyodorov, D. V., and Kadonaga, J. T., (2003) Chromatin assembly in vitro with purified recombinant ACF and NAP-1. Meth. Enzymol. 371, 499–515. 7. Kundu, T. K., Wang, Z., and Roeder, R. G. (1999) Human TFIIIC relieves chromatinmediated repression of RNA polymerase III transcription and contains an intrinsic

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histone acetyltransferase activity. Mol. Cell. Biol. 19, 1605–1615. Munakata, T., Adachi, N., Yokoyama, N., Kuzuhara, T., and Horikoshi, M., (2000) A human homologue of yeast anti-silencing factor has histone chaperone activity. Genes Cells. 5, 221–233. An. W., and. Roeder, R. G. (2004) Reconstitution and transcriptional analysis of chromatin in vitro. Meth. Enzymol. 377, 460–474. Balasubramanyam, K., Varier, R. A., Altaf, M., Swaminathan, V., Siddappa, N. B., Ranga, U., and Kundu, T.K. (2004) Curcumin, a novel p300/CREB-binding protein-specific inhibitor of acetyltransferase, represses the acetylation of histone/nonhistone proteins and histone acetyltransferase-dependent chromatin transcription. J. Biol. Chem. 279, 51163–51171. Lau, O.D., Kundu, T.K., Soccio, R.E., AitSi-Ali, S., Khalil, E.M., Vassilev, A., Wolffe, A.P., Nakatani, Y., Roeder, R.G., and Cole, P.A. (2000) HATs off: selective synthetic inhibitors of the histone acetyltransferases p300 and PCAF. Mol. Cell. 5, 589–595. Mantelingu, K., Reddy, B.A., Swaminathan, V., Kishore, A.H., Siddappa, N. B., Kumar, G.V., Nagashankar, G., Natesh, N., Roy, S., Sadhale, P.P., Ranga, U., Narayana, C., and Kundu, T.K. (2007) Specific inhibition of p300-HAT alters global gene expression and represses HIV replication. Chem. Biol. 14, 645–657.

Chapter 19 Histone Deacetylase Activity Assay Zhigang Yuan, Natalie Rezai-Zadeh, Xiaohong Zhang, and Edward Seto Abstract Histone deacetylases (HDACs) are enzymes that catalyze the removal of acetyl groups from the "-amino groups of conserved lysine residues in the amino terminal tail of histones. In humans, there are 18 potential deacetylase enzymes that are responsible for the removal of acetyl groups and maintenance of the equilibrium of lysine acetylation on histones. Like most histone modification enzymes, accumulating evidence suggests that many, if not all, HDACs can also modify non-histone proteins. The focus of this article is to provide up-to-date, easy to follow, approaches and techniques specifically for the assay of HDAC enzymatic activities. Key words: HDAC assay, HDAC2, HDAC7, sirtuin, SIRT1, protein deacetylation, core histone, non-histone substrate.

1. Introduction Reversible lysine acetylation is an important regulatory mechanism for a wide variety of biological processes. In cells, the lysine acetylation status of proteins is regulated dynamically through the opposing action of histone acetyltransferases (HATs) and histone deacetylases (HDACs). HDAC enzymes catalyze the removal of acetyl groups from lysine residues in histone and non-histone proteins. Deacetylation may lead to a change in the conformation and/or activity of the substrates. Through deacetylation, dynamic modification of lysine residues near the N-termini of histones plays an important role in the regulation of chromatin structure and gene expression. In general, hypoacetylation of histones is linked to transcriptional repression. In addition to their crucial roles in the control of gene expression, HDACs also participate in Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_19 Springerprotocols.com

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the regulation of a number of other cellular processes, including DNA repair, cell cycle control, cell differentiation, and apoptosis. Based on protein size, sequence similarity, and organization of the protein domains, eleven mammalian HDACs, which are zinc-dependent hydrolases, have been divided into two classes (reviewed in ref. (1, 2)). Class I HDACs, which include HDAC1, HDAC2, HDAC3, and HDAC8, are highly related to the yeast RPD3 protein. Class II HDACs contain HDAC4, HDAC5, HDAC6, HDAC7, HDAC9, and HDAC10 are homologues of the yeast HDA1 protein. The Class II enzymes have been subdivided further into two subclasses, IIa (HDAC4, 5, 7, and 9) and IIb (HDAC6 and 10). HDAC11 uniquely shares sequence homology within the catalytic domains of both class I and II HDACs. Class III HDACs, also called sirtuins (SIR2 like proteins), include SIRT1, SIRT2, SIRT3, SIRT4, SIRT5, SIRT6, and SIRT7. Sirtuins are structurally and evolutionarily unrelated to the Class I and II HDACs and require NAD+ as a cofactor for catalytic activity. With increased interest in HDAC enzymes over the last decade, it is very important to develop reliable and efficient approaches to prepare enzymatically active HDACs and assays for the detection of their activity. In this chapter, we present practical step-by-step protocols for scientists outside of the HDAC field who are interested in incorporating HDAC studies into their research. 1.1. Purification of Enzymatically Active Class I & II HDACs

The biological activity of the Class I and II HDACs is tightly modulated by the binding of cofactors and post-translational modifications, particularly phosphorylation (3). Although all Class I and II HDACs can be expressed easily as recombinant proteins in bacteria, we have found that, with the exception of HDAC8, HDAC proteins produced in Escherichia coli display very little enzymatic activities. In contrast, enzymatically active HDACs that are useful for most functional assays can be obtained from immunoaffinity purified proteins transiently or stably expressed in mammalian cells or insect cells (4). The methods described here allow for direct and efficient purification of enzymatically active Class I and II HDACs from mammalian and insect cells. Unlike bacterially expressed HDACs, HDAC proteins isolated from mammalian or insect cells retain their native conformation, modification, and interaction with cofactors. We present protocols for the expression and purification of HDAC2 and HDAC7 as examples. Identical approaches can be taken to obtain other active Class I and Class II enzymes.

1.2. Purification of Enzymatically Active Class III HDACs

The Class III HDACs (sirtuins, SIRT1-7) are broadly conserved among species. The importance of the sirtuins in biology is underscored by the fact that they show quite broad non-histone

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substrate specificity (1–4). Unlike Class I and II HDACs, sirtuins use NAD+ as a cofactor to remove the acetyl group from lysine residues through two-step reactions. In the first reaction, an acetylated protein and NAD+ bind to the active site of the enzyme. The N-pyridinium glycosidic bond of NAD is cleaved due to enzyme conformational changes, and a high-energy ADP-ribose intermediate is generated. In the second reaction, the high-energy ADP-ribose intermediate reacts with the acetyl group on the -amino group of the lysine. The acetyl group is transferred to the reactive ADP-ribose high-energy intermediate, and acetylADP-ribose is created. The products (a deacetylated protein and acetyl-ADP-ribose) then are released from the enzyme active site (5). The development of assays for sirtuins gains benefit from the two-step reactions, which use two reactants and create three products. In theory, the activity of sirtuins can be detected by monitoring the consumption of the reactant(s) or the generation of the product(s) (6).

2. Materials 2.1. Materials for Class I and II HDACs Assay

1. pME18S-Flag-HDAC2: HDAC2 cDNA subcloned into the expression vector, pME18S, as a flag-tagged fusion protein 2. HEK293T cells 3. Lipofectamine 2000 (Invitrogen cat. #11668019)

2.1.1. To Purify HDAC2 from HEK293T Cells

4. Phosphate-buffered saline (PBS) (140 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4 pH 7.4) 5. Cell lysis buffer (50 mM Tris–HCl pH 7.5, 150 mM NaCl, 0.5% NP40, 10% glycerol) 6. EDTA-free protease #11617900)

inhibitor

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7. Anti-Flag M2 agarose (Sigma cat. #A2220) 8. 1 HDAC assay buffer (10 mM Tris–HCl pH 8.0, 150 mM NaCl, 10% glycerol) 9. Flag peptide (Sigma cat. #F3290) 2.1.2. To Purify HDAC2 from Sf 9 Insect Cells

1. HDAC2 cDNA subcloned into the baculovirus transfer vector pFastBac-HTa (Invitrogen), which introduces a 6 -His tag at the N-terminus of HDAC2. Recombinant baculovirus encoding HDAC2 generated according to the manufacturer’s protocol. 2. Sf 9 cells.

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3. Cell lysis buffer (50 mM Tris–HCl pH 7.5, 100 mM NaCl, 1 mM MgCl2, 5 mM -mercaptoethanol, 10% glycerol, 0.5% Triton X-100). 4. Ni-NTA (nitrilo-triacetic acid) resin (Qiagen). 5. Buffer A (20 mM Tris–HCl [pH 8.5], 1 M KCl, 20 mM imidazole, 10% glycerol, 5 mM -mercaptoethanol). 6. Buffer B (20 mM Tris–HCl [pH 8.5], 1 M KCl, 10% glycerol, 5 mM -mercaptoethanol) 7. Buffer C (20 mM Tris–HCl [pH 8.5], 100 mM KCl, 200 mM imidazole, 10% glycerol, 5 mM -mercaptoethanol). 2.1.3. To Purify an Enzymatically Active HDAC7 Complex from HeLa Cells

1. Recombinant adenoviruses encoding HA- and flag-tagged HDAC7 (Ad-HA-HDAC7-Flag) 2. Cell lysis buffer (1 PBS, 0.1% NP40, 10% glycerol) 3. EDTA-free protease inhibitor cocktail (Roche cat. #11617900) 4. Anti-Flag M2 agarose (Sigma cat. #A2220) 5. Anti-HA agarose (Sigma cat. #A-2095 ) 6. Flag peptide (Sigma cat. #F3290) 7. HA peptide (Sigma cat. #I-2149 ) 8. Poly-prep chromatography column (Bio-Rad, cat. # 731-1550) 9. Buffer A (20 mM Tris pH 7.5, 1.0 mM EDTA, 10% glycerol, 1.0 mM dithiothreitol, and a protease inhibitor mixture)

2.1.4. To Purify Radioactive Substrates for Class I and II HDAC Enzymatic Assay

To assay Class I and Class II HDAC activity, acetylated core histones or acetylated synthetic histone H4 peptides are commonly used as substrates (7, 8). These substrates can be radioor fluorescently labeled. Following are the materials needed for small-scale labeling and isolation of core histones from HeLa cells with [3H] acetic acid. 1. HeLa cells. 2. 10 mg/ml cyclohexamide. 3. 2 M sodium butyrate (NaB). 4. 1.0 mCi/ml [3H]acetic acid (Perkin–Elmer cat. #3499232). 5. IB buffer (10 mM Tris–HCl pH 7.4, 2 mM MgCl2, 3 mM CaCl2, 10 mM sodium butyrate, 1 mM phenylmethylsulfonyl fluoride (PMSF) (prepare immediately prior to use.) 6. NIB buffer (1% nonidet P-40 in IB buffer) (prepare immediately prior to use.) 7. Storage buffer (10 mM HEPES pH 7.5, 1 mM EDTA, 10 mM KCl, 0.2 mM PMSF, 10% glycerol). 8. 5 M NaCl. 9. SnakeSkin1 Pleated Dialysis Tubing (Pierce cat. #68053).


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The shortcomings of using radioactively labeled core histones include batch-to-batch variation of in vivo labeling, exposure to radioactivity and the need for radioactive waste management (10). As an alternative to radiolabeled histones or peptides, a number of nonradioactive substrates recently have been developed based on colorimetric or fluorescently labeled peptides. HDAC activity can be detected readily by measuring the OD or quantifying fluorescence. 1. HDAC substrate [Boc-Lys(Ac)-pNA, 10 mM] (BioVision cat. #K331-100-1) 2. 10 HDAC assay buffer (BioVision cat. #K331-100-2) 3. Lysine developer (BioVision cat. #K331-100-3)

2.1.6. Materials for Class I and II HDAC Enzymatic Assay

We will describe a detailed protocol for small-scale labeling and isolation of core histones from HeLa cells with [3H] acetic acid, which is performed routinely in our laboratory and is adapted from Carmen et al. (7). Here we describe the use of radiolabeled acetylated core histone substrates. However, histones can be replaced with radiolabeled acetylated H4 peptide substrate in this assay. 1. 5 HDAC buffer (50 mM Tris–HCl pH 8.0, 750 mM NaCl, 50% glycerol) 2. 3H-labeled core histones 3. Stop solution (1 M HCl and 0.4 M acetic acid) 4. Ethyl acetate

2.2. Materials for Class III HDACs Assay

1. SIRT1 cDNA subcloned into pGEX-5X-1 vector (Amersham). The construct generates an N-terminal GST-tagged SIRT1 recombinant protein. 2. BL21 bacteria.

2.2.1. Materials for Expression and Purification of Class III HDACs from E. coli

3. IPTG. 4. STE buffer (10 mM Tris–HCl pH 8.0, 150 mM NaCl, 1 mM EDTA). 5. 10% N-lauryl sarcosine. 6. 1 M DTT. 7. Sonicator (Sonic Dismembrator 60, Fisher Scientific). 8. 10% Triton X-100. 9. Glutathione-agarose beads (Sigma cat. #G4510). 10. Elution buffer (10 mM glutathione in 50 mM Tris–HCl pH 8.0, 1 mM DTT).

2.2.2. Materials for Preparation of Substrates for Class III HDACs

Although radiolabeled core histones can be used for many in vitro Class III HDAC assays, it has been shown that a variety of non-histone proteins are the natural substrates for sirtuins in vivo (11, 12). Different assays using different substrates have been

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developed to test the deacetylase activity of sirtuins (13). We would present a detailed protocol for sirtuins enzymatic assay using a peptide corresponding to a portion of p53 as a substrate. 1. [3H]-acetic acid (NEN cat. #NET-003H, 2–5 Ci/mmol in ethanol) 2. [3H]-acetylated peptides containing residues 364–389 of p53 (5’-AHSSHLKSKKGQSTSRHKKLMFKTEG-30 ) 3. BOP solution (0.24 M BOP [Aldrich cat# 22,608-4] and 0.2 M triethylamine [Aldrich cat. #13,206-3] in acetonitrile [Aldrich cat. #27,071-7]) 4. Microcon-SCX spin column (Millipore cat. #42460) 2.2.3. Materials for Class III HDACs Enzyme Reaction

1. 3H-labeled p53 peptide 2. 5 HDAC buffer (50 mM Tris–HCl pH 8.0, 750 mM NaCl, 50% glycerol) 3. NAD+ cofactor 50 mM stock in ddH2O 4. Stop solution (100 mM HCl and 160 mM acetic acid) 5. Ethyl acetate

3. Methods 3.1. Protocols for Class I and II HDACs Assay

3.1.1. Purification of HDAC2 from HEK293T Cells

1. Transfect one 10 cm plate of HEK293T cells with 10 mg of pME18S-Flag-HDAC2 plasmid using Lipofectamine 2000 according to the manufacturer’s manual. In a separate transfection, use pME18S vector as a negative control. 2. Thirty-six hours post-transfection, wash cells twice with 1 PBS. Harvest cells in 1 ml fresh 1 PBS in a 1.5 ml tube. After centrifugation at 1,500 g for 5 min, lyse cell pellet in five packed-cell-pellet volumes of cell lysis buffer in the presence of an EDTA-free protease inhibitor cocktail for 30 min at 4C. (It is preferable not to include EDTA in the preparation of cell lysate.). Remove cell debris by centrifugation at 10,000 g for 15 min at 4C and transfer the supernatant to a prechilled 1.5 ml tube. (See Notes 1 and 2.). 3. Subject the supernatant to immunoprecipitation reactions with anti-Flag M2 agarose. Add 30 ml anti-Flag M2 agarose (prewashed with cell lysis buffer) per ml of cell lysate and agitate for 2 h at 4C. 4. Two hours later, pellet immune complexes by centrifugation at 10,000 g for 2 min at 4C. Carefully remove supernatant and wash immune complexes four times using cell lysis buffer. Subsequently, wash the immune complexes three times in


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1 HDAC assay buffer. Beads containing the immunoprecipitated HDAC2 are now ready to be used in the enzymatic assay. 5. (Optional) Flag-HDAC2 can be eluted with 50 ml of 1 mg/ ml of flag peptide in 25 mM Tris (pH 7.1), 120 mM NaCl and 10% glycerol. Dialyze protein against 25 mM Tris (pH 7.1), 120 mM NaCl and 10% glycerol and distribute into 1.5 ml centrifuge tubes. Store materials at –80C. The HDAC sample should be re-frozen immediately at –20 or –70C after each use to reduce loss of activity. 3.1.2. Purification of HDAC2 from Sf9 Insect Cells

1. To produce a high titer stock baculovirus encoding HDAC2, infect Sf 9 cells with recombinant virus at a Multiplicity of Infection (MOI) of 0.01–0.1. Once 15–20% cell survival is achieved, harvest the virus-containing cell supernatant by centrifugation at 2,000 g for 3 min at room temperature. 2. Infect Sf 9 cells with viral particles (virus/cell ratio of 10:1) and harvest cells 48 h post-infection by centrifugation at 500 g for 2 min at room temperature. 3. Lyse cells in ice-cold cell lysis buffer containing 1 EDTAfree protease inhibitor. Centrifuge at 10,000 g for 10 min. 4. Incubate the supernatant with Ni-NTA resin on a rotator for 1 h at 4C. 5. Wash resin successively with 10 volumes of buffer A and 2 volumes of buffer B. 6. Elute protein with buffer C. 7. Determine protein concentration by the Bradford method using the Bio-Rad dye reagent with BSA as a standard. Analyze purified HDAC2 protein by SDS-PAGE and Coomassie blue staining.

3.1.3. Purification of an Enzymatically Active HDAC7 Complex from HeLa Cells

1. Infect two 150 mm plates of HeLa cells grown to 90% confluency with Ad-HA-HDAC7-Flag or control GFP adenoviruses. (See Note 3.) 2. Thirty-six hours later, wash cells with PBS and harvest into a 50 ml conical tube. Centrifuge for 3 min at 2,000 rpm. 3. Lyse cells by adding 4 ml of cell lysis buffer. 4. Incubate on ice for 20 min and mix gently every 5 min. Sonicate samples for 15 s using a sonic dismembrator. Centrifuge for 15 min at 10,000 g at 4C. Transfer supernatant into an ice-cold tube. 5. Equilibrate a Poly-Prep chromatography column with 1 ml cell lysis buffer. It is important that steps 5–12 are performed at 4C (e.g., in a cold room).

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6. Add 200 ml of anti-Flag agarose and wash agarose in the column with 5 ml of cell lysis buffer. 7. Cap bottom of the column. Add the cell lysate to the column and cap the top. Rock for 3 h at 4C. 8. Uncap column and collect flow-through. 9. Wash column extensively with lysis buffer using 10 ml lysis buffer. 10. Elute bound protein complex with 1 ml of 0.2 mg/ml flag peptide and collect five fractions of eluate (200 ml each fraction). Analyze 10 ml of each fraction by silver staining. 11. Pool eluted fractions and reload onto a 1 ml spin column containing 50–100 ml of anti-HA agarose. Repeat steps 7–9. 12. Elute bound protein with 100 ml of 0.5 mg/ml HA peptide five times. Collect fractions and analyze by silver staining. 13. Analyze eluate fractions by Western blotting with anti-Flag antibodies. 14. Remove a small fraction of the purified HDAC7 protein complex for enzymatic activity assay as described in Section 2.4. 15. Fractionate the HDAC7 protein complex further using a Superose 6 fast flow (Amersham Biosciences) column with 0.4 ml/min with buffer A containing 150 mM KCl and collection of eluates in 200 ml fractions. Analyze Aliquots from alternate fractions by silver staining and immunoblotting with appropriate antibodies (Fig. 19.1). 3.1.4. Preparation of Substrates for Class I and II HDACs Enzymatic Assay

1. Prepare one confluent 150 mm plate of HeLa cells. 2. Wash cells once with 1 PBS. 3. Incubate cells with mix A (20 ml PBS and 200 ml of 10 mg/ml cyclohexamide) for 10 min at 37C. 4. Aspirate mix A and incubate cells with mix B (20 ml PBS, 200 ml of 10 mg/ml cyclohexamide, 100 ml NaB and 400 ml 1.0 mCi/ml [3H]acetic acid) at 37C for 1 h. 5. Collect cells and chill on ice for 10 min. The following steps are done at 4C or on ice. 6. Centrifuge at 1,500 g for 5 min at 4C to pellet cells. 7. Wash cells with 5 ml PBS containing 25 ml of 2 M NaB three times. 8. Wash cells twice with 1 ml NIB buffer. (See Note 5.) 9. Wash cells once with 1 ml NIB buffer containing 100 mM NaCl. 10. Wash cells once with 1 ml IB buffer containing 100 mM NaCl. 11. Wash cells twice with 500 ml of IB buffer containing 400 mM NaCl.

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Fig. 19.1. A representative experiment to show purification of an enzymatically active HDAC7 protein complex. (A) Silverstained SDS-PAGE of the immunopurified Flag- and HA-tagged HDAC7-containing complexes (top panel ). Results of deacetylase activity assays from control and purified HDAC7 materials (bottom panel ). (B) Silver stained of SDS-PAGE (top panel ) and Western blots (bottom panels) of Flag- and HA-tagged HDAC7 after purification in immunoaffinity and size exclusion columns.

12. From steps 7–11, pellet cells by centrifugation at 500 g for 5 min at 4C. 13. Extract the nuclei pellet with 0.25 ml of 0.2 N H2SO4 for 90 min on ice. 14. Centrifuge at 30,000 g for 90 min at 4C. 15. Dialyze supernatant against 500 ml of storage buffer overnight at 4C. Aliquot purified core histones and store at –80C. (See Notes 4, 5 and 6.)

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3.1.5. Class I and II HDACs Enzymatic Assay Using Radiolabeled Core Histone

1. For each reaction, the following reagents are mixed in a 1.5 ml tube in a final volume of 200 ml, containing 40 ml of 5 HDAC buffer, 3H-labeled core histones (about 10,000 cpm), the HDAC sample and ddH2O. 2. Incubate reactions at 30C for 2 h. 3. Stop reaction by the addition of 50 ml of stop solution. 4. Extract released 3H-acetate by adding 0.4 ml ethyl acetate with mixing. Centrifuge at 10,000 g for 3 min at room temperature to separate phases. 5. Transfer 200 ml of the upper (organic) phase to a scintillation vial with 3 ml scintillation liquid and measure CPM in a scintillation counter. (See Note 7.) 6. To calculate the enzymatic activity, the equation in Fig. 19.2 is used. 7. HDAC activity ¼ (counts from test sample [cpm] – counts from control [cpm])/volume of sample (ml). For the negative control, HDAC reaction can be heated to 95C for 5 min to destroy any enzymatic activity. 1. Dilute the HDAC sample with ddH2O to a final volume of 85 ml in one well within a 96-well plates with U-shape bottom. (See Note 8).

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Fig. 19.2. A representative experiment to determine HDAC activity using radiolabeled core histones. Cells were transfected with either Flag-HDAC2 or empty vector for 48 h, lysed and immunoprecipitated with anti-Flag M2 agarose beads. Immunoprecipitated samples then were subjected to HDAC assays as described using approximately 8,000 cpm core histone as substrate. The expression of Flag-HDAC2 was monitored by Western blotting using anti-Flag antibodies.


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2. For the background reading, add 85 ml ddH2O only. For the negative control, dilute the heated-HDAC sample with ddH2O to a final volume of 85 ml. 3. Add 10 ml of the 10 HDAC assay buffer to each well. 4. Add 5 ml of the HDAC colorimetric substrate to each well. Mix thoroughly. 5. Incubate the reaction at 37C for 2 h or at room temperature overnight. 6. Stop the reaction by adding 10 ml of lysine developer and mix well. Incubate the plate at 37C for 30 min. (See Note 9.) 7. Read sample in an ELISA plate reader at 400 or 405 nm; the signal is stable for several hours at room temperature. HDAC activity can be expressed as the relative O.D. value per mg protein sample (See Fig. 19.3) (See Note 10.)

3.2.1. Expression and Purification of Class III HDACs from E. coli

1. Transform BL21 bacteria with plasmid expressing GSTSIRT1. Grow a 15–20 ml overnight culture in the presence of antibiotics. 2. Inoculate bacteria into 250 ml LB broth with antibiotics and incubate at 37C until the OD600 reaches 0.6 (about 2 h). 3. Add IPTG to the culture to a final concentration of 0.2 mM and incubate bacteria for 4 h. 4. Pellet cells by centrifugation at 6,500 g for 10 min at 4C.

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Fig. 19.3. A representative experiment using non-radiolabeled substrates from BioVision for measuring HDAC activity. Reactions were carried out either at 37C for 2 h or overnight at room temperature. HDAC activity was measured by O.D. at 405 nm.

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5. Resuspend pellet in 5 ml of ice-cold STE buffer. Addition of 100 mg/ml of lysozyme and 1 protease inhibitor to the STE buffer is optional. 6. Add DTT to final concentration of 5 mM. 7. Add N-lauryl sarcosine to a final concentration of 1.5% from a 10% stock. 8. Mix cells thoroughly and sonicate at 4C until the lysate becomes clear. (See Note 11.) 9. Add Triton X-100 to a final concentration of 3% from a 10% stock. 10. Centrifuge insoluble fractions at 10,000 g at 4C for 10 min. 11. Aliquot supernatants into microfuge tubes (100–200 ml each), snap freeze in liquid nitrogen (or dry ice with ethanol) and store at –80C. (optional). 12. Pre-swell 30 ml glutathione–agarose beads slurry and mix with the supernatant. Rotate to mix at 4C for 2 h or overnight. 13. Wash beads with PBS five times to remove unbound and non-specifically bound proteins. (See Note 3.) 14. Elute fusion protein by adding 100 ml elution buffer. Repeat 10 times. 15. Analyze individual fractions by SDS-PAGE and Coomassie blue staining; pool fractions containing the highest amounts of purified recombinant protein, aliquot and store at –80C. 3.2.2. Preparation of non-Histone Substrate for Class III HDACs Assay

1. Radiolabel the substrate by incubating 0.4 mg of p53 peptide with 5 mCi of [3H]acetic acid. Use synthetic p53 peptide that was purified by HPLC to greater than 90% purity. Add 10 ml of BOP solution to the peptide/label mix and rock gently on a rocker at room temperature overnight. BOP solution should always be prepared fresh. 2. Prewash a column with 500 ml of 10 mM HCl in methanol once and then with 500 ml of 10 mM HCl in 10% methanol, centrifuging columns at 1,200 g for 1 min and discarding eluates. 3. Purify the labeled peptide using the Microcon-SCX spin column in a chemical hood. Load 250 ml of the labeling mixture onto the prewashed column and spin down at 1,200 g for 1 min. 4. Wash the column twice with 500 ml of 10 mM HCl in 10% methanol. Invert and place the column in a new collection tube, then centrifuge at 1,200 g for 1 min. 5. To elute the labeled peptide, apply 50 ml of 3 N HCl in 50% isopropanol to the column, followed by centrifugation at 14,000 g for 15 s. The elution step can be repeated to ensure complete elution of the labeled peptide.


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6. Dry eluate in a fume hood with the cap of the collection tube open or in a Speedvac. Then, dissolve the lyophilized peptide in 500 ml of ddH2O. Aliquot the dissolved radioactive peptide and store at –80C. A successful acetylation reaction should yield a value of not less than 1,000 cpm. (See note 12.) 3.2.3. Class III HDACs Enzyme Reaction

1. Mix 40 ml of 5 HDAC buffer, 3H-labeled p53 peptide (about 20,000 cpm) and 1 mg of GST-SIRT1 in a 1.5 ml tube in a final volume of 200 ml. (See Notes 13 and 14.) 2. Start the enzymatic reaction by adding 1 mM NAD+ cofactor and incubate for 2 h at 30C. 3. Stop the reaction by the addition of 25 ml of stop solution. 4. Extract released 3H-acetate by adding 0.4 ml ethyl acetate and vortexing for 15 s. Centrifuge at 10,000 g for 5 min at room temperature. 5. Transfer 200 ml of the upper phase (the ethyl acetate fraction) to a scintillation vial with 3 ml scintillation liquid and measure CPM in a scintillation counter. The product of the reaction catalyzed by sirtuins is acetylADP-ribose, not acetate. It is not understood why the acetate release assay also functions with the Class III HDACs. Free acetate might be released from acetyl-ADP-ribose under the acidic experimental conditions (11). Several studies suggest that different sirtuins may utilize different substrates. For example, while SIRT1 targets p53, tubulin might be a more suitable substrate for SIRT2. Acetylated targets can be detected either by pulse labeling the cells with radiolabeled acetate or by Western blotting with target-specific acetylation antibody or pan-acetylated lysine antibodies. Pan-acetylated lysine antibodies are available commercially from many different companies including Millipore and Cell Signaling and are the most convenient reagent to study acetylation/deacetylation. It is wise to test several different pan-acetylated lysine antibodies against your protein of interest since each antibody targets different acetylated lysines with different efficiencies. Since the basal acetylation level of many proteins may be too low to be detectable by standard procedures, a common alternative is to treat cells overnight with nicotinamide (20 mM) or TSA (400 ng/ml), or both before analysis.

4. Notes 1. HEK293T cells can be detached easily from culture plates by scraping and transferring into a conical tube followed by two washes with PBS.

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2. To maximize the release of HDACs from nuclei, cells can be sonicated three times for 15 s each at the output setting of 4 (Sonic Dismembrator 60, Fisher Scientific). 3. HeLa cells grown in suspension are preferred for large-scale preparations. 4. To ensure that the isolated core histones are labeled correctly, an aliquot of core histones can be measured for specific activity using a scintillation counter. Using this protocol, the counts for 10 ml of core histones are approximately 10,000 cpm. 5. Since PMSF is readily degraded in water-based solutions, it is important that IB buffer, NIB buffer and storage buffer are freshly made. Alternatively, complete EDTA-free protease inhibitor cocktail tablets (Roche cat. #11836170001) can be used in place of PMSF. 6. For many HDACs, core histones or histone H4 peptide works well as substrate. However, some HDACs may require mononucleosome instead of free histones as a substrate. The preparation of mononucleosome substrate has been described elsewhere (7–9). 7. The upper phase should be very carefully transferred to the scintillation vial using a pipet tip to avoid background count carry over from interphase contamination. 8. 96-well plates with U-shape bottoms are preferred. Flat bottom plates may give unreliable low-value readings. 9. Refreeze the lysine developer immediately at –20 or –70C after each use, or aliquot for future use. 10. A variety of other nonradioactive colorimetric or fluorescencebased HDAC assay kits are commercially available from Millipore, BIOMOL and Calbiochem. 11. Typically, sonicating the bacteria five times for 15 s each at the output setting of 4 is sufficient. 12. To estimate labeling efficiency, 1 ml of the purified peptide is diluted into 100 ml of H2O and extracted with 400 ml of ethyl acetate for scintillation counting. The counts then should be adjusted to 20,000 cpm for each deacetylase assay. 13. GST-SIRT1 used in the assay can be replaced with an immunoprecipitated SIRT1 protein from transfected mammalian cells as described for HDAC2. 14. In addition to the radiolabeled p53 peptide, fluorogenic acetylated peptides of residues 379–382 or 317–320 of p53 can be used. These peptides substrates are commercially available from BIOMOL.


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References 1. Yang, X.J., and Seto, E. (2008) The Rpd3/ Hda1 family of lysine deacetylases: from bacteria and yeast to mice and men. Nat Rev Mol Cell Biol. 9(3):206–18. 2. Glozak, M.A., Sengupta, N., Zhang, X., and Seto, E. (2005) Acetylation and deacetylation of non-histone proteins. Gene. 363:15–23. 3. Heltweg, B., Trapp, J., and Jung, M. (2005) In vitro assays for the determination of histone deacetylase activity. Methods. 36(4):332–7. 4. Rezai-Zadeh, N., Tsai, S.C., Wen, Y.D., Yao, Y.L., Yang, W.M., and Seto, E. (2004) Histone deacetylases: purification of the enzymes, substrates, and assay conditions. Methods Enzymol. 377:167–79. 5. Sauve, A.A., Celic, I., Avalos, J., Deng, H., Boeke, J.D., and Schramm, V.L. (2001) Chemistry of gene silencing: the mechanism of NAD–dependent deacetylation reactions. Biochemistry. 40(51):15456–63. 6. Landry, J., and Sternglanz, R. (2003) Enzymatic assays for NAD-dependent deacetylase activities. Methods. 31(1):33–9. 7. Carmen, A.A., Rundlett, S.E., and Grunstein, M. (1996) HDA1 and HDA3 are components of a yeast histone deacetylase (HDA) complex. J Biol Chem. 271(26):15837–44. 8. Sun, J.M., Spencer, V.A., Chen, H.Y., Li, L., and Davie, J.R. (2003) Measurement of

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histone acetyltransferase and histone deacetylase activities and kinetics of histone acetylation. Methods. 31(1):12–23. Utley, R. T., Owen-Hughes, T. A., Juan, L. J., Cote, J., Adams, C. C., and Workman, J. L. (1996) In vitro analysis of transcription factor binding to nucleosomes and nucleosome disruption/displacement. Methods Enzymol. 274:276–91. Kolle, D., Brosch, G., Lechner, T., Lusser, A., and Loidl, P. (1998) Biochemical methods for analysis of histone deacetylases. Methods. 15(4):323–31. North, B.J., Schwer, B., Ahuja, N., Marshall, B., and Verdin, E. (2005) Preparation of enzymatically active recombinant class III protein deacetylases. Methods. 36(4):338–45. Imai, S., Armstrong, C.M., Kaeberlein, M., and Guarente, L. (2000) Transcriptional silencing and longevity protein Sir2 is an NAD-dependent histone deacetylase. Nature. 403(6771):795–800. Marcotte, P.A., Richardson, P.L., Guo, J., Barrett, L.W., Xu, N., Gunasekera, A., and Glaser, K.B. (2004) Fluorescence assay of SIRT protein deacetylases using an acetylated peptide substrate and a secondary trypsin reaction. Anal Biochem. 332(1):90–9.

Chapter 20 In Vitro and In Vivo Assays for Studying Histone Ubiquitination and Deubiquitination Ling Zhai, Heui-Yun Joo, and Hengbin Wang Abstract Posttranslational histone modifications play important roles in regulating chromatin structure and function (Martin and Zhang, Nat Rev Mol Cell Biol 6:838–849, 2005; Jenuwein and Allis, Science 293:1074–1080, 2001). One example of such modifications is histone ubiquitination, which occurs predominately on H2A and H2B. Recent studies have highlighted important regulatory roles of H2A ubiquitination in Polycomb group proteins-mediated gene silencing (Wang et al., Nature 431:873–878, 2004; Joo et al., Nature 449:1068–1072, 2007) and H2B ubiquitination in transcription, H3 methylation, and DNA methylation (Zhang, Genes Dev 17:2733–2740, 2003; Sun and Allis, Nature 418:104–108, 2002; Sridhar et al., Nature 447:735–738, 2007). Here we describe methods for in vitro histone ubiquitination and deubiquitination assays. We also describe approaches to investigate the in vivo function of a putative histone ubiquitin ligase and deubiquitinase. These experimental procedures are largely based on our studies in mammalian cells. These methods should provide useful tools for studying this bulky histone modification. Key words: Chromatin, histone ubiquitination, histone deubiquitination, in vitro assay, and in vivo assay.

1. Introduction The organization and function of chromatin, the physiological template for all nuclear processes involving genetic information, are largely dictated by posttranslational histone modifications (1–5). These modifications, which include acetylation, methylation, phosphorylation, ubiquitination, and sumoylation, are believed to mark an individual nucleosome or a short stretch of nucleosome arrays, with a specific ‘‘code’’, which is recognized by downstream regulatory proteins and elicits specific cellular Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_20 Springerprotocols.com

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responses (2–4). The extensive efforts recently made to identify and characterize the enzymes involved in histone acetylation, methylation, and phosphorylation, together with the development of antibodies which can specifically recognize acetylated, methylated, and phosphorylated histones, have brought significant insight into the subcellular distribution, dynamics of global and locus-specific levels, and biological functions of these modifications (2–4). Compared with other histone modifications, ubiquitination is unique in that the bulky ubiquitin protein is two-thirds the mass of an individual histone (6, 7). This is likely the reason why antibodies specifically recognizing ubiquitinated histones are difficult to generate and only the anti-ubiquitinated H2A (uH2A) antibody has been described (8). Despite this drawback, experimental procedures which employ a combination of genetic and immunoblot approaches have been developed for studying histone ubiquitination both in vitro and in vivo (9–11). These experimental procedures, together with the recently identified enzymes responsible for histone ubiquitination and deubiquitination, have revealed important functions of histone ubiquitination in regulating transcription, cell cycle progression, recombination, and DNA damage responses (6, 9, 10, 12, 13). Of the four core histones, H2A and H2B are ubiquitinated in a wide range of organisms and cell types (6, 7). Ubiquitination of H2B is highly conserved and has been reported in budding yeast, Arabidopsis, Drosophila, and mammals (6, 7). The abundance of ubiquitinated H2B (uH2B) varies from organism to organism, which can account for up to 10% of total cellular H2B (14). Rad6, a multifunctional ubiquitin conjugating E2 enzyme, was identified as the E2 enzyme for histone H2B and the specific activity of Rad6 for histone H2B is conferred by a Ring-finger containing protein Bre1 (14–16). Research interest in H2B ubiquitination was boosted by the finding that H2B ubiquitination regulates K4 and K79 methylation of histone H3 in an unidirectional, transhistone fashion (17, 18–20). Recently, H2B deubiquitination in Arabidopsis thaniliana was found to regulate DNA methylation and heterochromatin gene silencing (21). Unlike H2B, ubiquitination of H2A has not been detected in budding yeast, Caenorhabditis eleganslegans, and Arabidopsis (14, 21, 22). The ubiquitin ligase for histone H2A was recently identified as the Polycomb group protein Ring2, suggesting that H2A ubiquitination may be an epigenetic mechanism that is acquired later during evolution (9, 22). In addition to H2A and H2B, histones H3, H4, H2A.Z, and H1 have also been reported as substrates for ubiquitin modification (7). The ubiquitin ligase for histone H3 and H4 was identified as the Cullin 4-associated ubiquitin ligase complex and ubiquitination of these histones may facilitate the DNA damage repair process (10). The enzyme responsible for H1 ubiquitination was identified as TAFII 250 and ubiquitination of

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Fig. 20.1. Ubiquitination and deubiquitination reactions. For protein ubiquitination, ubiquitin is first activated by E1 ubiquitin activating enzyme in a ATP-dependent formation of ubiquitin-E1 thiol ester. Ubiquitin is then transferred to E2 ubiquitin conjugating enzymes, again through the formation of adenylate intermediate. Finally, the ubiquitin E3 ligase brings together the E2-ubiquitin complex and the substrates and mediates the isopeptide bond formation. Ubiquitin conjugated to target proteins can be cleaved by specific deubiquitinase and results in the release of intact ubiquitin.

histone H1 has been linked to gene repression in the dorsal pathway in Drosophila (23). Similar to ubiquitination of other proteins, histone ubiquitination occurs through a series of enzymatic reactions (Fig. 20.1). First, ubiquitin is activated by the E1 ubiquitin activating enzyme through an ATP-dependent formation of ubiquitin-E1 thiol ester via a ubiquitin adenylate intermediate. Second, E1 transfers activated ubiquitin to E2 ubiquitin conjugating enzymes through the formation of a new ubiquitin adenylate intermediate. Finally, the ubiquitin E3 ligase conjugates ubiquitin to the -N of lysine residues in the target protein through an isopeptide bond. Since the substrate specificity is primarily determined by the ubiquitin E3 ligase, a great deal of effort has been made to identify the ubiquitin ligases for different substrates (24, 25). Like other histone modifications, histone ubiquitination is a reversible process (26, 27). Ubiquitin conjugated to target proteins can be cleaved by thiol-specific proteases known as deubiquitinating enzymes or deubiquitinases (26, 27). The identification of H2B deubiquitinases in budding yeast and Arabidopsis revealed that histone deubiquitination also plays an important role in regulating chromatin functions (21, 28–31). In the following sections, we

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describe experimental procedures for in vitro histone ubiquitination and deubiquitination assays. We also describe experimental procedures for studying the in vivo function of a putative histone ubiquitin ligase and deubiquitinase. These procedures are mainly based on our studies in mammalian cells. Detailed experimental protocols for studying histone ubiquitination in budding yeast can be found in a recent review (11).

2. Materials 2.1. In Vitro Histone Ubiquitination Assay

1. Substrates. Histone octamers, mononucleosomes, and oligonucleosomes prepared from HeLa S3 cells (32). All materials should be dialyzed against histone storage buffer (10 mM HEPES–KOH [pH 7.5), 1 mM EDTA, 10 mM KCl, 10% glycerol, 0.2 mM PMSF) and stored at –80C (see Note 1). 2. Reaction buffer. Prepare reaction buffer from stock solutions immediately before use. 1 M Tris–HCl (pH 7.5), 1 M MgCl2,100 mM NaF, 100 mM ATP, 10 mM Okadaic acid, 0.05 M DTT. 3. Other reagents. E1 ubiquitin activating enzyme (Calbiochem), E2 ubiquitin activating enzyme (ubc5c), Flag-ubiquitin (Sigma), anti-Flag M2 antibody (Sigma). 4. Equipment. SDS–PAGE apparatus, semi-dry transfer unit, power supply.

2.2. In Vitro Histone Deubiquitination Assay

1. Substrate preparation. Cells: HeLa cells and yeast T85 strain (31). Constructs: pcDNA3-Flag-H2A (neomycin resistant), pcDNA3-Flag-H2B (neomycin resistant), and pcDNA3HA-ubiquitin (hygromycin resistant). Buffers: buffer A: 0.25 M sucrose, 60 mM KCl, 15 mM NaCl, 10 mM MES–KOH (pH 6.5), 5 mM MgCl2, 1 mM CaCl2, 0.5% Triton X-100, 1 mM DTT, and 0.1 mM PMSF; buffer B: 10 mM Tris–HCl (pH 8.0), 100 mM NaCl, 5 mM CaCl2,2 mM MgCl2, 0.3 M sucrose, 1 mM DTT, and 0.1 mM PMSF; nucleosome extraction buffer: 20 mM Tris–HCl (pH 7.9), 10 mM EDTA, 0.5 M KCl, 1 mM DTT, and 0.1 mM PMSF; denaturing buffer: 20 mM Tris–HCl (pH 7.4), 50 mM NaCl, 0.5% nonidet P-40 (NP-40), 0.5% sodium deoxycholate, 0.5% SDS, 1 mM EDTA, 1 mM DTT, and 0.1 mM PMSF;


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elution buffer: 50 mM Tris–HCl (pH 7.4), 150 mM NaCl, 0.1 mM PMSF; TAE buffer: 40 mM Tris–acetate and 1 mM EDTA; buffer P: 5 mM HEPES–KOH (pH 7.5), 0.04 M KCl, 10 mM potassium phosphate (ph 7.5) 0.01 mM CaCl2, 10 % glycerol, 1 mM DTT, and 0.1 mM PMSF; histone unfolding buffer: freshly prepared. 6 M guanidinium chloride, 20 mM Tris–HCl (pH 7.5), and 5 mM DTT; histone refolding buffer: 2 M NaCl, 10 mM Tris–HCl (pH 7.5), 1 mM EDTA, and 5 mM 2-ME; 2 and 5 M NaCl; 10 mM Tris–HCl (pH 7.5); 50% sucrose; 5% native PAGE. Other reagents: recombinant histones (33), neomycin, hygromycin, micrococcal nuclease (200 U/ml, 40% protein, balance primarily sodium acetate, Sigma, prepared in 5 mM NaH2PO4 [pH 7.0], 2.5 mM CaCl2, see Note 2), anti-Flag M2 affinity gel (Sigma), flag peptide (custom synthesized), anti-Flag M2 antibody (Sigma), anti-HA antibody, HA peptide (custom synthesized), YPD medium, Effectene (Qiagen). 2. 5 reaction buffer. 500 mM Tris–HCl (pH 8.0), 5 mM EDTA, 0.5 mM PMSF, 5 mM DTT, 5 mg/ml aprotinin, 5 mg/ml leupeptin, and 5 mg/ml pepstatin A. 3. Equipment. HPLC (GE Healthcare Life Sciences), 1 ml hydroxyapatite column (Bio-Rad), Seperdex-200 gel filtration column (GE Healthcare Life Sciences), Amicon ultra centrifugal filter (Millipore), SDS–PAGE apparatus, semi-dry transfer unit, power supply, and benchtop microcentrifuge. 2.3. In Vivo Histone Ubiquitination and Deubiquitination Assay

1. HeLa cells 2. Constructs: pcDNA3-Flag-H2A (neomycin resistant), pcDNA3-Flag-H2B (neomycin resistant), and pcDNA3-HAubiquitin (hygromycin resistant). 3. Other reagents: neomycin, hygromycin, anti-Flag M2 affinity gel (Sigma), flag peptide (custom synthesized), anti-HA antibody, Effectene (Qiagen), Lipofectamine 2000 (Invitrogen), and denaturing buffer (see Section 2.2.1) 4. Equipment: SDS-PAGE apparatus, semi-dry transfer unit, and power supply.

3. Methods 3.1. In Vitro Histone Ubiquitination Assay

1. Mix histone octamers (5 mg), mononucleosomes (5 mg), and oligonucleosomes (5 mg) with protein fractions or a putative histone ubiquitin ligase in a 32.5 ml reaction system containing

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50 mM Tris (pH7.5), 5 mM MgCl2, 2 mM NaF, 2 mM ATP, 10 mM Okadaic acid, 0.6 mM DTT, 0.1 mg E1, 0.2 mg E2 (ubc5c), 1 mg Flag-ubiquitin (Sigma). 2. Incubate the reaction mixture at 37C for 1 h. Stop the reaction by mixing with SDS sample loading buffer. 3. Resolve the reaction product in a 15% or 8–15% SDS-PAGE and transfer proteins to a nitrocellulose membrane. 4. Perform a standard Western blotting assay with anti-Flag antibody. Ubiquitinated histones migrate around the size of individual histone plus Flag-ubiquitin (8 kDa, Fig. 20.2). 5. Negative control assays: omit E1, E2, substrate, and E3 (protein fraction or the putative H2A ubiquitin ligase) individually from the ubiquitin ligase reaction and perform parallel experiments. The signal for ubiquitinated histones should be abolished.

Fig. 20.2. In vitro histone ubiquitination assay. a. Ubiquitin ligase assay using HeLa nuclear proteins fractionated on DE52 and P11 columns. Numbers on top of the panels indicate the salt concentration (M) for step elution. NE and NP represent nuclear extracts and nuclear pellet, respectively. Left and right panels use histone octamer and oligonucleosome substrates, respectively. b. The ligase activity depends on the presence of ATP, E1, E2, ubiquitin, nucleosomal histones, and proteins present in the 0.5 M P11 nuclear pellet fraction. Reproduced with permission from Wang et al. (9).


3.2. In Vitro Histone Deubiquitination Assay

3.2.1. Purification of Native uH2A-Containing Mononucleosomes

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1. Linearize pcDNA3-Flag-H2A and pcDNA3-HA-ubiquitin with Mfe1 and recover the linearized plasmid with the Wizard SV Gel and PCR Clean-up System (Promega). Cotransfect these two plasmids into HeLa cells with Effectene (Qiagen) following the manufacturer’s instruction. Forty-eight hours after transfection, split the cells (1:3) and add neomycin and hygromycin to a final concentration of 0.5 and 0.3 mg/ml, respectively. Continue culture until individual clones form. Test the expression of Flag-H2A and HA-ubiquitin in each clone by Western blot assay with anti-Flag and HA antibodies. Stable cells with high expression of both epitope-tagged proteins will be used for the following experiment (see Note 3). 2. Collect stable cells from two to three 15 cm plates (yield 7–10 mg of mononucleosomes) and wash twice with cold PBS. Resuspend cells in 1 ml buffer A and incubate on ice for 15 min. Collect nuclei by centrifugation at 1,200 x g, 4C for 10 min (benchtop microcentrifuge) and wash once more with buffer A. 3. Wash nuclei with buffer B once and resuspend the nuclei in 0.4 ml buffer B. Add 2 ml micrococcal nuclease (200 U/ml) and incubate at 37C for 30 min (see Note 4). Mix well every 10 min. After digestion, wash nuclei once with buffer B. 4. Resuspend digested nuclei in 0.4 ml nucleosome extraction buffer and incubate on ice for 15 min. Centrifuge at 16,100 g for 10 min (benchtop microcentrifuge) and save the supernatant as nucleosomes. Repeat the extraction until no protein comes out. 5. Combine the extracted nucleosomes and measure protein concentration by Bradford assay (Bio-Rad). Dilute nucleosomes to a final concentration of 0.25 mg/ml by adding drops of histone storage buffer while vortexing vigorously. To avoid precipitation, 0.05% NP-40 should be added to histone storage buffer prior to dilution. After dilution, dialyze samples against histone storage buffer for at least 4 h (see Note 5). 6. Wash 100 ml anti-Flag M2 affinity gel with histone storage buffer three times and incubate with nucleosome preparation at 4C for 2 h. After incubation, wash resin with histone storage buffer three times. Elute proteins bound to the resin with 0.2 mg/ml Flag peptide (in histone storage buffer) at 4C for 30 min. Repeat the elution step 2–3 times until no proteins are eluted out. Dialyze the eluate against histone storage buffer at 4C for 2 h. 7. Incubate the dialyzed anti-Flag eluate with anti-HA antibody that has been cross-linked to protein A at 4C for 2 h. Wash resin with histone storage buffer three times. Elute proteins bound to the anti-HA resin with 0.8 mg/ml HA peptide (in

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histone storage buffer containing 0.5 M NaCl). Dialyze the anti-HA eluate against histone storage buffer and store at 4C. Purified mononucleosomes should be used within 2 weeks. 3.2.2. Reconstitution of Ubiquitinated HistoneContaining Mononucleosomes

1. DNA fragments used for nucleosome reconstitution are amplified in a PCR reaction with the PXP-10 plasmid as template and 50 -Bio-TGTTACTCAGAATGGCAA-30 and 50 -TGATTACGAATTCGAGCT-30 as primers (see Note 6). Resolve the PCR products in a 1% agarose gel and purify with the Wizard SV Gel and PCR Clean-up System (Promega). 2. Recombinant H3, H4, H2A, and H2B are purified from E. Coli by chromatography and stored at –80C after lyophilization (33). 3. To purify histones containing uH2A, stable cells expressing Flag-H2A (2–3 15 cm plates, see Section 3.2.1) are dissolved in 0.5 ml denaturing buffer with the aid of sonication. After centrifugation at 16,100 x g, 4C (benchtop microcentrifuge) for 10 min, the supernatant is incubated with 100 ml anti-Flag M2 affinity gel at 4C for 2 h with rotation. After washing with denaturing buffer and elution buffer three times each, proteins bound to resin are eluted with 0.2 mg/ml flag peptide (in elution buffer) at room temperature for 20 min. Repeat the elution steps until no proteins are eluted out. Dialyze samples against water overnight and lyophilize. Store at –80C. 4. Histones containing uH2B are purified from yeast strain T85 under denaturing conditions and proteins are eluted in elution buffer (see Note 7). Dialyze samples against water overnight and lyophilize. Store at –80C. 5. To further purify uH2A and uH2B, resolve the anti-Flag eluate on a 15% SDS-PAGE. In order to obtain enough samples, multiple lanes can be loaded. uH2A and uH2B migrate around 25 kDa, while uH2B migrates a little higher than uH2A. Excise gel strips containing uH2A and uH2B, based on their relative positions with protein size markers. As controls, we also excise gel strips containing H2A and H2B. Place gel strips in 15 ml Falcon tubes and wash extensively with water. Transfer gel strips to a dialysis tube (with molecular weight cutoff of 6–8,000 Da) and immerse in elution buffer. Place the dialysis tube in TAE buffer. Apply 60 V to elute proteins from gel slices at 4C (See Note 8). Load sample on a 1 ml hydroxyapatite column and elute proteins with buffer P containing 1 M KCl. Dialyze fractions containing uH2A and uH2B against water at 4C for overnight. Lyophilize and store at –80C. 6. Histone octomer refolding. Dissolve recombinant histones, uH2A, uH2B, H2A, and H2B into histone unfolding buffer at a concentration of 1 mg/ml and incubate at 4C for 1 h. Mix the histones at equal molar ratios as judged by Coomassie blue


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staining. Dialyze the mixture against histone refolding buffer for 24 h with at least three changes of dialysis buffer. Load the samples onto a Superdex-200 gel filtration column (GE Healthcare Life Sciences). Concentrate fractions containing histone octamers (Amicon Ultra, Millipore). Samples can be stored at 4C upto 2 months. 7. Nucleosome reconstitution. Dilute DNA with 5 M NaCl to a final concentration of 2 M NaCl. Add appropriate amount of octamers (See Note 9). Incubate for 30 min at room temperature, then add an equal volume of 10 mM Tris–HCl (pH 7.5) and incubate for 1 h at room temperature. Sequentially dilute the mixture to reduce the concentration of NaCl from 1.0, 0.8, 0.67, 0.2 M, 0.1 M by adding 10 mM Tris–HCl (pH 7.5). Incubate for 1 h at room temperature after each dilution. 8. Resolve a small aliquot of the reconstituted nucleosomes in a 5% native PAGE. Stain the gel with ethidium bromide and visualize under UV light. Successfully reconstituted mononucleosomes migrate around 700 bp, while naked DNA migrates around 200 bp (Fig. 20.3, see Note 10). 3.2.3. Histone Deubiquitination Reaction

1. Mix 1.5 mg uH2A-containing mononucleosomes or 0.8 mg of reconstituted uH2A or uH2B-containing mononucleosomes with protein fractions or a putative histone deubiquitinase in deubiquitination reaction buffer. Set up reactions for five time points (0, 10, 20, 40, and 60 min). 2. Incubate at 37C for the appropriate time. Stop each reaction by mixing with SDS sample loading buffer. Resolve reaction mixture in a 15% SDS-PAGE and transfer proteins to a nitrocellulose membrane.

Fig. 20.3. Nucleosome reconstitution with uH2A and uH2B.a. A Coomassie blue stained gel containing different histone octamers as indicated on the top of the panel. The identity of each protein is indicated on the right side of the panel. b. An ethidium bromide-stained 5% native polyacrylamide gel containing different reconstituted nucleosomes, and naked DNA that was used to reconstitute nucleosomes. Reproduced with permission from Joo et al. (34).

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Fig. 20.4. Histone deubiquitination assay of HeLa nuclear proteins fractionated on DE52 and P11 columns. Numbers on top of the panels indicate the salt concentration (M) for step elution. NE and NP represent nuclear extracts and nuclear pellet, respectively. Left and right panels show western blot analysis of the reaction mixture after 0 min and 45 min incubation at 37C. Adapted from Joo et al. (34).

3. Perform a standard Western blot assay with the anti-HA antibody. The histone deubiquitinase activity is reflected by decreased levels of ubiquitinated histones and increased release of intact ubiquitin (Fig. 20.4, see Note 11). 3.2.4. In Vivo Assays for Histone Ubiquitination and Deubiquitination

1. Linearize pcDNA3-Flag-H2A and pcDNA3-Flag-H2B with Mfe1 and recover the linearized plasmid with the Wizard SV Gel and PCR Clean-up System (Promega). Transfect these plasmids into HeLa cells with Effectene (Qiagen) following the manufacturer’s instruction. Forty-eight hours after transfection, split cells (1:3) and add neomycin at a final concentration of 0.5 mg/ml. Continue culture until individual clones form. Test the expression of Flag-H2A and Flag-H2B in individual clones by Western blot assay with the anti-Flag antibody. Choose cell lines with high expression of Flag-H2B and low expression of H2A for further experiments.


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2. Seed stable cells in a 6 well plate at a density of 1 105 cells/ well and incubate at 5% CO2, 37C overnight. 3. Transfect siRNA against the putative histone ligase or deubiquitinase with Lipofectamine 2000 and incubate for an additional 24 h at 37C. Transfect HA-ubiquitin into cells with Effectene and incubate for another 24 h at 37C. 4. Harvest cells and wash once with PBS. Dissolve cell pellet in denaturing buffer (add protease inhibitors prior to use) with the aid of sonication. Centrifuge at 16,100 x g at 4C for 10 min (benchtop microcentrifuge), take one-tenth of the supernatant as input. Incubate the remaining supernatant with anti-Flag M2 affinity gel at 4C for 2 h. 5. Wash resin three times with denaturing buffer and elute the bound proteins with 0.2 mg/ml Flag peptide (in denaturing buffer). Resolve the eluate in SDS-PAGE and transfer proteins to a nitrocellulose membrane. 6. After blocking, cut the nitrocellulose membrane into two parts around the size of the molecular weight marker of 20 kDa. Blot the top half of the membrane with anti-HA antibody and the bottom half with anti-Flag antibody. Perform a standard Western blot assay (see Note 12) (Fig. 20.5).

Fig. 20.5. In vivo histone deubiquitination assay of Ubp-M. HeLa stable cell lines expressing F-H2A (lanes 1–3) and F-H2B (lanes 4–6) were transfected with the indicated combinations of siRNA and plasmid. Western blot analysis was performed with total cellular proteins (top three panels) and anti-Flag immunoprecipitate under denaturing conditions (bottom two panels). Reproduced with permission from Joo et al. (34).

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4. Notes 1. Oligonucleosomes can be stored at 4C for at least 1 month without significant change in their ability to serve as substrates for histone ubiquitin ligase reactions. Mononuclesomes are less stable and precipitation can occur even after overnight storage at 4C. Therefore, it is recommended to store nucleosomes, especially mononucleosomes, in small aliquots at –80C to avoid multiple rounds of freeze and thaw. We also found that the presence of histone H1 slightly reduces the ability of nucleosomes to serve as substrates for some histone ubiquitin ligases. Therefore, it is recommended to use oligonucleosomes without H1 as substrates for histone ubiquitination reactions. In addition, we found that the ability of nucleosomes to serve as substrates for histone ubiquitination varies from batch to batch. 2. Micrococcal nuclease from Sigma is less precisely controlled digestion. Digestion nuclease from Worthington Biochemical harder to control and therefore it is not use in this experiment.

active but allows with micrococcal (Freehold, NJ) is recommended to

3. We routinely obtain 5–6 clones that can be used for further experiments from cotransfection of two 6-well plates. Clones that only express one epitope-tagged protein can be used for the experiments in Sections 3.2.2 and 3.3. Sequential transfection with one plasmid and then using the stable clones for the second transfection is time-consuming and less efficient. 4. Digestion of nuclei (from two to three 15 cm plates) for 30 min yields primarily mononucleosomes. To obtain oligonucleosomes, we first optimize the conditions of digestion by comparing different time points. Bulk nuclei are digested to give nucleosomes ranging from 1 to 15 nucleosomes. We separate mono- and oligonucleosomes by a 5 ml 5–30% sucrose gradient, which is prepared by loading layers of increasing concentrations of sucrose (mixing the 5% and 30% sucrose solution at different ratios). Histone octamers containing uH2A are prepared by a hydroxyapatite column (32). 5. To observe the release of intact ubiquitin, uH2A-containg mononucleosomes, which are purified from anti-Flag and anti-HA tandem immunoprecipitation, are required. However, as a readout of the histone deubiquitination activity, the reduction of signal from ubiquitinated histones can be observed in bulk nucleosome preparations. Therefore, it is recommended to first confirm that the reduction of uH2A signal is indeed due to the presence of a specific deubiquitinase


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using double immunoprecipitation-prepared nucleosomes and once confirmed, the reduction of uH2A signal can be used as an indicator for the histone deubiquitination activity. 6. PCR program: 94C, 4 min, 1 cycle; 94C, 30 sec 50C, 30 sec 40 cycles; 72C, 30 sec 72C, 7 min, 1 cycle. Expected size of PCR product: 232 bp. 7. The level of uH2B in budding yeast depends on cell density and culture conditions. We routinely culture T85 yeast strains in YPD medium and use OD600 of 0.3–0.5 to prepare uH2B. 8. We wash gel strips three times in a 15-ml tube for 30 min with rotation. The gel strips are then transferred to a dialysis tubing with molecular weight cutoff of 6–8,000 Da. To further reduce the contamination with SDS, we use elution buffer to elute proteins and TAE buffer for electrophoresis. 9. For every freshly prepared DNA and/or histone octamers, we first calculate the molar ratio of DNA and histone octamers. We then use equal molar ratio and ratios slight higher or lower to reconstitute nucleosomes in a small scale. The quality of reconstituted nucleosomes is checked in a 5% native PAGE to determine the best ratio of DNA and histone octamers for large-scale reconstitution. 10. Pre-run the 5% native PAGE (59:1, acrylamide:bisacrylamide) for 1 h at 4C at 150 V. Mix reconstituted nucleosomes with 50% sucrose at ratio of 10:1 (volume) and load the sample. 11. In some cases, it is difficult to see the increase of intact ubiquitin with the anti-HA antibody. In order to see the released intact ubiquitin, we use the anti-ubiquitin (FK2) antibody. 12. It is recommended to cut the membrane to probe each half separately. If the entire membrane is probed with the antiFlag antibody, the ubiquitinated histones cannot be seen due to the large amount of non-ubiquitinated histones. Additionally, probing the top half of the membrane with anti-HA antibody gives less background. References 1. Martin, C., and Zhang, Y. 2005. The diverse functions of histone lysine methylation. Nat Rev Mol Cell Biol 6:838–849. 2. Jenuwein, T., and Allis, C.D. 2001. Translating the histone code. Science 293:1074–1080.

3. Strahl, B., and Allis, C. 2000. The language of covalent histone modifications. Nature 403:41–45. 4. Turner, B. 2000. Histone acetylation and an epigenetic code. Bioessays 22: 836–845.

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5. Peterson, C.L., and Laniel, M.-A. 2004. Histones and histone modifications. Curr Biol 14:R546–R551. 6. Osley, M.A. 2004. H2B ubiquitylation: the end is in sight. BBA-Gene Struct Expr 1677:74–78. 7. Jason, L., Moore, S., Lewis, J., Lindsey, G., and Ausio, J. 2002. Histone ubiquitination: a tagging tail unfolds? Bioessays 24:166–174. 8. Vassilev, A., Rasmussen, H., Christensen, E., Nielsen, S., and Celis, J. 1995. The levels of ubiquitinated histone H2A are highly upregulated in transformed human cells: partial colocalization of uH2A clusters and PCNA/ cyclin foci in a fraction of cells in S-phase. J Cell Sci 108:1205–1215. 9. Wang, H., Wang, L., Erdjument-Bromage, H., Vidal, M., Tempst, P., Jones, R.S., and Zhang, Y. 2004. Role of histone H2A ubiquitination in Polycomb silencing. Nature 431:873–878. 10. Wang, H., Zhai, L., Xu, J., Joo, H.-Y., Jackson, S., Erdjument-Bromage, H., Tempst, P., Xiong, Y., and Zhang, Y. 2006. Histone H3 and H4 Ubiquitylation by the CUL4-DDB-ROC1 Ubiquitin Ligase Facilitates Cellular Response to DNA Damage. Mol Cell 22:383–394. 11. Kao, C.-F., and Osley, M.A. 2003. In vivo assays to study histone ubiquitylation. Methods 31:59–66. 12. Shilatifard, A. 2006. Chromatin modifications by methylations and ubiquitinations: Implications in the Regulation of Gene Expression. Annu Rev Biochem 75:243–269. 13. Fleming, A., and Osley, M.A. 2004. Silence of the rings. Cell 119:449–451. 14. Robzyk, K., Recht, J., and Osley, M.A. 2000. Rad6-dependent ubiquitination of histone H2B in yeast. Science 287:501–504. 15. Hwang, W.W., Venkatasubrahmanyam, S., Ianculescu, A.G., Tong, A., Boone, C., and Madhani, H.D. 2003. A conserved RING finger protein required for histone H2B monoubiquitination and cell size control. Mol Cell 11:261–266. 16. Wood, A., Krogan, N.J., Dover, J., Schneider, J., Heidt, J., Boateng, M.A., Dean, K., Golshani, A., Zhang, Y., and Greenblatt, J.F. 2003. Bre1, an E3 ubiquitin ligase required for recruitment and substrate selection of Rad6 at a promoter. Mol Cell 11:267–274.

17. Sun, Z.-W., and Allis, C.D. 2002. Ubiquitination of histone H2B regulates H3 methylation and gene silencing in yeast. Nature 418:104–108. 18. Briggs, S.D., Xiao, T., Sun, Z.-W., Caldwell, J.A., Shabanowitz, J., Hunt, D.F., Allis, C.D., and Strahl, B.D. 2002. Gene silencing: Trans-histone regulatory pathway in chromatin. Nature 418:498. 19. Dover, J., Schneider, J., Tawiah-Boateng, M.A., Wood, A., Dean, K., Johnston, M., and Shilatifard, A. 2002. Methylation of histone H3 by COMPASS requires ubiquitination of histone H2B by Rad6. J Biol Chem 277:28368–28371. 20. Ng, H.H., Xu, R.-M., Zhang, Y., and Struhl, K. 2002. Ubiquitination of histone H2B by Rad6 is required for efficient Dot1-mediated methylation of histone H3 lysine 79. J Biol Chem 277:34655–34657. 21. Sridhar, V.V., Kapoor, A., Zhang, K., Zhu, J., Zhou, T., Hasegawa, P.M., Bressan, R.A., and Zhu, J.-K. 2007. Control of DNA methylation and heterochromatic silencing by histone H2B deubiquitination. Nature 447:735–738. 22. de Napoles, M., Mermoud, J.E., Wakao, R., Tang, Y.A., Endoh, M., Appanah, R., Nesterova, T.B., Silva, J., Otte, A.P., and Vidal, M. 2004. Polycomb group proteins Ring1A/B link ubiquitylation of histone H2A to heritable gene silencing and X inactivation. Dev Cell 7:663–676. 23. Pham, A.-D., and Sauer, F. 2000. Ubiquitinactivating/conjugating activity of TAFII250, a mediator of activation of gene expression in drosophila. Science 289:2357–2360. 24. Pickart, C.M. 2001. Mechanisms underlying ubiquitination. Annu Rev Biochem 70:503–533. 25. Pickart, C.M. 2004. Back to the future with ubiquitin. Cell 116:181–190. 26. Wilkinson, K. 1997. Regulation of ubiquitin-dependent processes by deubiquitinating enzymes. FASEB J 11:1245–1256. 27. Wilkinson, K.D. 2000. Ubiquitination and deubiquitination: Targeting of proteins for degradation by the proteasome. Semin Cell Dev Biol 11:141–148. 28. Daniel, J.A., Torok, M.S., Sun, Z.-W., Schieltz, D., Allis, C.D., Yates, J.R., III, and Grant, P.A. 2004. Deubiquitination of histone H2B by a yeast acetyltransferase complex regulates transcription. J Biol Chem 279:1867–1871.


29. Henry, K.W., Wyce, A., Lo, W.-S., Duggan, L.J., Emre, N.C.T., Kao, C.-F., Pillus, L., Shilatifard, A., Osley, M.A., and Berger, S.L. 2003. Transcriptional activation via sequential histone H2B ubiquitylation and deubiquitylation, mediated by SAGA-associated Ubp8. Genes Dev 17:2648–2663. 30. Gardner, R.G., Nelson, Z.W., and Gottschling, D.E. 2005. Ubp10/Dot4p regulates the persistence of ubiquitinated histone H2B: distinct roles in telomeric silencing and general chromatin. Mol Cell Biol 25:6123–6139. 31. Emre, N.C.T., Ingvarsdottir, K., Wyce, A., Wood, A., Krogan, N.J., Henry, K.W., Li, K., Marmorstein, R., Greenblatt, J.F., Shilatifard, A., et al. 2005. Maintenance of low histone ubiquitylation by Ubp10 correlates with telomere-proximal sir2

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Chapter 21 Detection of Histone H3 Phosphorylation in Cultured Cells and Tissue Sections by Immunostaining Jaya Padmanabhan Abstract Growth factor stimulation results in phosphorylation of histone H3 at ser 10 and this correlated with expression of immediate early genes suggesting that this phosphorylation is associated with transcriptional activation. Although Western immunoblot analysis allows the detection of protein modifications in histones, in order to determine the localization of histones during different phases of cell cycle or during treatment of cells with different drugs we have to use immunohistochemistry. The protocol described here allows the detection of phosphorylated histones in tissue-cultured cells and tissue sections by fluorescent or bright-field immunostaining analysis. Here we used a serine 10 specific P-histone H3 antibody to determine the localization of this phosphoprotein in an asynchronously growing H4 glioma cell line and brain sections. It has been shown that long-term potentiation (LTP) is associated with gene transcription, and histone acetylation plays a major role in LTP formation (Wood et al., Learn Mem 13:241–244, 2006; Wood et al., Hippocampus 15:610–621, 2005; Alarcon et al., Neuron 42:947–959, 2004; Korzus et al., Neuron 42:961–972, 2004). Stimulus-induced phosphorylation of histone H3 at serine 10 has also been implicated in hippocampal neurons and striatal neurons (Li et al., J Neurochem 90:1117–1131, 2004; Crosio et al., J Cell Sci 116:4905–4914, 2003). Co-staining with a cell-specific antibody will allow us to determine the type of cells that show activation of histone phosphorylation in the brain. Key words: Histone, phosphorylation, immunofluorescence, immunohistochemistry.

1. Introduction Protein phosphorylation is one of the several posttranslational modifications that affect the activity and function of a protein and this affects the signaling cascades that is associated with that specific protein (1). Many of these signal transduction pathways are initiated at the cell surface and transported to the nucleus where the transcription is under tight control. The chromatin fiber in Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_21 Springerprotocols.com

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the nucleus is composed of repetitive units called nucleosomes that are comprised of core histones. These include H2A, H2B, H3 and H4. Chromatin organization is altered by modifications of the histones by acetylation, phosphorylation, methylation, ubiquitination, etc. (2, 3). Histone acetylation has been extensively correlated with transcriptional activation. H3 phosphorylation at serine 10 has been shown to be associated with transcriptional activation and chromosome condensation during mitosis and meiosis (1, 4–15). During cell division, phosphorylation is first detected in pericentromeric heterochromatin and is associated with the whole chromosome in mitosis. This phosphorylation is also important for passage of cells from metaphase to anaphase. During interphase histone H3 ser10 phosphorylation is associated with the activation of several genes associated with signal transduction. Dephosphorylation of H3 begins in anaphase and ends in early telophase. Association of phosphorylated histones with different mitotic machinery can be detected using immunostaining analysis of cells (6, 8, 16, 17). In addition to its role in cell proliferation, histone modifications have additional functions in quiescent cells such as neurons (18–23). Dissociated cultures or sections of brains can be used to determine histone phosphorylation. The cells or tissue sections are fixed. Permeabilization and non-specific blocking are achieved in a single step. After incubation of cells with the antibody of interest, the staining is detected using fluorescent secondary antibodies or bright-field techniques.

2. Materials 1. Chamber slides for tissue culture (Fisher Scientific) 2. Fisherfinest cover glasses (Fisher Scientific) 3. Colorfrost plus microscope slides (Fisher Scientific) 4. Biomedia mounting media (Gel/Mount) (Fisher Scientific) 5. Anti-phospho-histone H3 (serine 10) rabbit polyclonal antibody (Cell Signaling Technology, Inc., Danvers, MA) 6. Anti-alpha-tubulin mouse monoclonal antibody (SigmaAldrich, St. Louis, MO) 7. Anti-neuron-specific nuclear (NeuN) antibody (Millipore, Bedford, MA) 8. Alexa fluor 488 goat anti-mouse IgG (H+L) (Invitrogen Corporation, Carlsbad, CA) 9. Alexa fluor 568 goat anti-rabbit IgG (H+L) (Invitrogen Corporation, Carlsbad,CA) 10. Biotinylated anti-rabbit IgG (H+L) (Vector Laboratories, Inc., Burlingame, CA)

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11. GIBCO goat serum (Invitrogen Corporation, Carlsbad, CA) 12. Poly L-lysine (Sigma-Aldrich, St. Louis, MO) 13. Bovine serum albumin (BSA) (fraction V) (Sigma-Aldrich, St. Louis, MO) 14. bisBenzimide H 33342 trihydrochloride (Hoechst 33342) (Sigma-Aldrich, St. Louis, MO) 15. VECTASTAIN Elite ABC kit (Vector Laboratories, Inc., Burlingame) 16. DAB peroxidase substrate kit (Vector Laboratories, Inc., Burlingame) 17. ImmEdge Pen, a hydrophobic barrier pen (Vector Laboratories, Inc., Burllingame) 18. Ethanol (EtOH), 100 and 95% 19. Xylene 20. Nembutal for anesthetizing the mice is obtained from the animal facility at the University 21. Cytoseal XYL: xylene-based mounting media (Fisher Scientific) 2.1. Solutions

1. Phosphate-buffered saline (PBS): 1 PBS: dissolve 8 g NaCl, 0.2 g KCl, 1.44 g Na2HPO4, 0.24 g KH2PO4 in 800 ml of distilled H2O and adjust the pH to 7.4 10 PBS: dissolve 80 g NaCl, 2 g KCl, 14.4 g Na2HPO4 and 2.4 g KH2PO4 in 800 ml water. Adjust the pH to 7.4 and bring up the volume to 1 L with distilled water. 2. Tris-buffered saline (TBS): 1 TBS: dissolve 3.03 g Tris base (25 mM) and 8.766 g NaCl (150 mM) in distilled water. Adjust pH to 7.4 and bring up the volume to 1 L with distilled water. 10 TBS: dissolve 30.3 g Tris base and 87.66 g NaCl in 800 ml distilled water. Adjust pH to 7.4 and bring up the volume to 1 L with distilled water. 3. 10% Triton X-100: mix 5 ml Triton X-100 with 45 ml distilled water by end-over-end mixing. Triton X-100 is really viscous and will take time to get a complete homogenous solution. Vortexing and vigorous shaking will result in foaming of the solution. The prepared 10% solution can be diluted further with required buffers to make solutions that contain 0.2% Triton X-100. 4. Tris-buffered saline containing 0.2% Triton X-100 (TBST): mix 1 ml of 10% Triton X-100 with 49 ml 1 TBS. 5. 10% sodium azide: dissolve 1 g sodium azide in 10 ml of water and store at room temperature (see Note 1).

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6. Blocking solution (1% BSA/TBST): dissolve 0.5 g bovine serum albumin (BSA) in 50 ml of 1 Tris-buffered saline (TBS) containing 0.2% Triton X-100 (TBST). Add 0.1 ml of 10% sodium azide to prevent bacterial growth, filter sterilize and keep at 4C. 7. Alternative blocking solution: 10% normal goat serum in TBST: dilute 10 ml normal goat serum with 90 ml of TBS containing 0.2% Triton X-100 to make 100 ml solution. 8. bisBenzimide H 33342 trihydrochloride (Hoechst 33342): Hoechst is dissolved in distilled water at a concentration of 10 mg/ml and aliquoted and stored at –20C. 9. 4% paraformaldehyde (PFA): add 4 g paraformaldehyde to 80 ml dH2O at 60C, add 1 drop of 2 N NaOH to the solution and stir until the PFA is completely dissolved. Do not let the temperature of the solution shoot above 60C. Once dissolved, add 10 ml of 10 PBS and bring up the volume to 100 ml. Filter the solution through a 0.45 mm filter and store at 4C (see Note 1). 10. Saline (0.9% NaCl): dissolve 0.9 g NaCl in 100 ml distilled water or 9 g NaCl per liter. 11. Citrate buffer, pH 6.0: A: Prepare 0.2 M solution of acetic acid by diluting 11.55 ml acetic acid in 1 L dH2O B: Prepare 0.2 M solution of sodium acetate by dissolving 16.4 g of C2H3O2Na or 27.2 g of C2H3O2Na.3H2O in 1 L dH2O. 10 mM citrate buffer: mix 4.8 ml of A þ 45.2 ml of B and dilute to 900 ml with dH2O, adjust pH to 6 and bring up to 1 L. 2.2. Equipment

1. Leica Model SM2000R freezing stage microtome 2. Zeiss AxioImager upright microscope with Axio Vision Rel 4.6 or any fluorescent microscope with appropriate software program for fluorescent and bright-field analysis of cells and tissue sections.

3. Methods 3.1. Immunostaining Analysis Using P-Histone H3 Serine 10 Antibody

1. Culturing cells for immunostaining analysis: H4 glioma cells are adherent and are cultured in chamber slides for immunocytochemical analysis. The chamber slides are coated with 100 mg/ml poly L-lysine from Sigma for 30 min in the tissue culture hood at room temperature. After the 30 min


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incubation, the slides are washed once with distilled water and cells are plated (see Note 2). 2. Fixation of cells: after treatment of the cells, aspirate the tissue culture medium with great care and fix the cells with 4% paraformaldehyde (PFA). Add enough PFA to cover the cells (250 ml/well for eight-chamber slides) and incubate for 10 min at room temperature. Remove PFA and carefully wash the cells with 1 PBS three times. 3. Inhibition of non-specific binding: incubate the cells with blocking solution (1% BSA/TBST) for 1 h at room temperature, on a platform rocker at very low speed. Alternatively, blocking can be achieved by incubating the cells with 10% normal goat serum diluted in TBS/0.2% Triton-X-100 for 1 h at room temperature. 4. Primary antibody incubation: anti-P-histone H3 (ser10) antibody is used at a concentration of 5 mg/ml diluted in 1% BSA/TBST (blocking buffer) and cells are incubated overnight at 4C, on a platform rocker at low speed. After the primary antibody incubation, the slides are washed thoroughly with PBS (4 times, 5 min each). At this point cells can be stained for analysis by bright-field or fluorescent microscopy. 3.2. For Fluorescent Microscopy

1. Cells are incubated with fluorescent-labeled secondary antibody. The secondary antibody is chosen depending on the host of the primary antibody. In this particular case the P-His H3 antibody used is a rabbit polyclonal antibody and the secondary antibody used is Alexa fluor 568 anti-rabbit. Alexa fluor 568 is diluted 1–4000 with blocking buffer and cells are incubated for 2 h at room temperature on a platform rocker. The fluorescent antibodies are light sensitive and this step has to be done in dark. The slides can be placed in a dark box or covered with aluminum foil before placing on the rocker. 2. Nuclear stain: at the end of the incubation cells are washed with PBS twice and incubated with Hoechst. Hoechst is diluted in PBS to get a concentration of 1 mg/ml and the cells are incubated for 2–5 min with Hoechst at room temperature. Cells are washed twice more with PBS. 3. Removal of wells and gasket from slides: the wells from the chamber slides are removed by prying it apart, and the rubber gasket is removed using a forceps. Rinse the slide once more with PBS and mount using Gel/Mount. Mounting media from Vector laboratories also work very well. 4. Storage: the slides are placed flat in slide trays and kept at 4C protected from light overnight and analyzed for P-HisH3 positive cells using a Zeiss Axioimager upright microscope using Axiovision Rel 4.6 software.

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3.2.1. Double Labeling of Cells Using Two Different Antibodies

If the cells need to be double labeled with a cell-specific marker or another antibody specific for some other antigen, then the primary antibody to that specific antigen has to be chosen in such a way that it is raised in a different host. For example, an alpha-tubulin antibody raised in mouse can be used along with anti-P-HisH3 antibody raised in rabbit to determine the phosphorylation of histones and microtubule organization simultaneously in cells. 1. Cells are fixed and blocked as mentioned under Section 5.1 2. The primary antibodies are diluted in 1% BSA/TBST as follows: 5 mg/ml anti-Ser10-P-His H3 rabbit polyclonal and 1 in 3000 dilution of anti-alpha tubulin mouse monoclonal antibodies. Cells are incubated with this mixture overnight at 4C. 3. Secondary antibody incubation: after washing thoroughly, the cells are incubated in the dark with secondary antibodies Alexa 568 anti-rabbit (1–4000) and Alexa 488 anti-mouse (1–1000) fluorophores diluted in 1% BSA/TBST for 1–2 h at room temperature. Cells are washed twice with PBS by rocking for 5 min each. 4. Nuclear staining using Hoechst: cells are incubated with Hoechst (1 mg/ml in PBS) for 2–5 min at room temperature. After two more washes the wells and gasket are removed from the slide. 5. Mounting of slides: the slides are mounted using Gel/Mount for analysis with fluorescent microscope. A few drops of the mounting media is added to the slides and coverglass is laid carefully. Any air bubbles captured between the slide and the coverglass is squeezed out by applying gentle pressure on the coverglass. The cells shown in the figure are double labeled with antitubulin and anti-P-His H3 (Ser10) antibodies, and Hoechst is used for nuclei staining (Fig. 21.1 and Color Plate 1, following p. 322). Do not allow the slides to dry in between the staining procedure (see Note 3).

3.2.2. For Bright-Field Analysis

This is done using the VECTASTAIN ABC kit from Vector laboratories with slight modifications from the manufacturer’s protocol. (For bright-field staining we always use only one primary antibody specific for a particular antigen, in this case P-His H3, although coimmunostaining can be done using two specific primary antibodies). The steps up to the secondary antibody incubation are done as in Section 5.1. 1. After primary antibody incubation the cells are washed thoroughly with PBS and incubated for 1 h at room temperature with biotinylated anti-rabbit IgG (H+L) from Vector laboratories diluted 1–200 in blocking buffer. 2. VECTASTAIN ABC reagent: while the cells are being incubated, the VECTASTAIN ABC reagent is prepared as follows. To 5 ml of blocking buffer add 2 drop of reagent A, mix


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Fig. 21.1. Coimmunostaining analysis of H4 cells using P-histone H3 (serine 10) and alpha-tubulin antibodies: The cells are immunostained using (A) P-histone H3 (serine 10) antibody and (B) alpha-tubulin antibody. Staining was visualized using Alexa 594 goat anti-rabbit or Alexa 488 goat anti-mouse fluorophores, respectively. (C) Represents staining of the same cells using Hoechst to analyze nuclei, and D is the composite image showing all the three stainings. The arrows indicate P-histone H3 positive cells in different phases of the cell cycle. (see Color Plate 1, following p. 322)

and add two drop of reagent B and mix again. Incubate this mix at room temperature for 30 min before use. 3. Once the incubation with biotinylated antibody is finished, cells are washed and incubated with the VECTASTAIN ABC reagent for 1 h at room temperature. 4. After this incubation cells are washed again thoroughly and incubated with peroxidase substrate using the DAB kit from Vector Laboratories following the manufacturer’s protocol. Develop until the desired intensity is reached, and the slides are washed in water to stop further development of the color. 5. The wells and gaskets are removed from the slide and mounted using Gel/Mount as mentioned under fluorescent microscopy or using xylene-based mounting media as follows. 6. Prior to mounting with xylene-based mounting media cells are dehydrated: Incubate for 2 min in 70% EtOH Incubate for 2 min in 95% EtOH

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Incubate for 2 min in 100% EtOH twice Incubate for 2 min in 100% xylene twice 7. Mount the slides using xylene-based mounting media, Cytoseal, for analysis with bright-field microscopy using a Zeiss Axioimager upright microscope using Axiovision Rel 4.6 software or Nikon E1000 microscope and Image Pro-Plus software program. 3.2.3. Staining of Cells Grown in Suspension

When the cells to be analyzed are grown in suspension, centrifuge the cells at 3000 rpm for 4 min and fix by resuspending in 4% paraformaldehyde. After 10 min fixation, cells are washed three times with PBS by centrifuging and aspirating the PBS and replacing with fresh PBS. After the final wash cells are incubated with blocking buffer and primary and secondary antibodies in suspension as mentioned above. Once the staining is finished cells are washed and collected on to a slide by cytocentrifugation (cytospin) and analyzed by fluorescent or bright-field microscopy depending on the secondary antibody used.

3.3. Double Immunohistochemical Analysis of Brain Tissue Sections with Ser10-P-Histone H3 and Neuron-Specific NeuN Antibodies

Preparation of tissue for immunostaining: 1. Perfusion: mice are sedated by nembutal (10 mg/kg body weight) and perfused with saline (50 ml) until the liver becomes pale. The perfusion is continued using 4% paraformaldehyde (30 ml), and brains are removed carefully. Immersion-fix the brain overnight in 4% paraformaldehyde at 4C. 2. Cryoprotection of tissue: after the fixation is complete, the brain is cryoprotected by passing through three different sucrose gradients; 10, 20 and 30%. Let the brain sink through the sucrose solution before replacing with the higher concentration of sucrose solution. Once the brain sinks through all the sucrose solutions it is ready for sectioning. 3. Sectioning of brain: the brains are frozen onto the freezing stage and 25 mm thick sections are made using the sliding microtome (Leica model SM2000R). The sections are placed in PBS containing 0.02% sodium azide and stored at 4C. 4. Mounting of section on slides: the sections are mounted on to frosted slides. To do so, a drop of 30% ethanol was placed on the slide and sections are placed into this using a thin paintbrush. Allow the section to slowly spread with the help of the brush. Let the sections dry onto the slide. It takes few hours to overnight depending on the amount of liquid on the slide. Once dry, using rubber cement make an edge around the sections (see Note 4). This will help to retain the solutions on the slide during staining protocol. Once the rubber cement is dry, swell the tissue by immersing in PBS in a coplin jar for 5 min. Wash once with PBS.


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1. To inhibit non-specific binding the sections are blocked for 2–3 h with 1% BSA/TBST containing 0.2% Triton X100 (same as in staining for cells mentioned above, except the tissue is incubated for longer time period). Blocking can also be achieved by incubating the sections with 10% normal goat serum in TBST. All the incubations are done in a humid atmosphere to avoid drying of the sections on slides. 2. Incubate with primary antibodies Ser10-P-histone H3 rabbit (5 mg/ml) and anti-NeuN mouse (1 to 200 dilution) antibodies diluted in blocking buffer (1% BSA/TBST) overnight at 4C on the platform rocker at low speed. 3. Sections are washed with PBS thoroughly and incubated with Alexa 568 anti-rabbit (1 in 4000) and Alexa 488 anti-mouse (1 in 1000) secondary antibodies for 2–3 h at room temperature, protected from light. 4. At the end of the incubation the slides are washed twice with PBS and incubated with the DNA stain, Hoechst diluted in PBS to a concentration of 1 mg/ml for 5 min in dark. Remove the rubber cement and wash the slides two more times before mounting using Gel/Mount. After overnight at 4C in dark, sections are analyzed under a Zeiss Axioimager upright microscope using Axiovision Rel 4.6 software or Nikon E1000 fluorescent microscope using Genus 2.8 software program (Fig. 21.2 and Color Plate 2, following p. 322).

3.5. Immunohistochemical Analysis of ParaffinEmbedded Sections

Removal of paraffin and rehydration of sections: With paraffinembedded sections, the paraffin is first removed as follows: 1. The slides containing the sections are heated at 60C for 1 h. 2. Following this the sections are immersed in xylene for 3 min twice. 3. The sections are rehydrated by passing sequentially through 100% EtOH 2 min twice, 95% EtOH 2 min once and 70% EtOH 2 min once. The sections are then rinsed in distilled water. 4. Antigen retrieval: this is achieved by heating the sections for 10 min at 95C in citrate buffer pH 6.0. The sections are then cooled slowly to room temperature and rinsed twice with dH2O. 5. Inhibition of endogenous peroxidase activity: quenching of peroxidase activity is achieved by incubating the sections for 20 min in methanol containing 0.5% H2O2 at room temperature. 6. After rinsing the sections twice with PBS, they are incubated with blocking buffer and staining continued as mentioned

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Fig. 21.2. Immunohistochemical analysis of brain sections using P-histone H3 (serine 10) and NeuN antibodies: Brain sections are coimmunostained using (A) P-histone H3 (serine 10) and (B) neuron-specific NeuN antibody. Staining was visualized using Alexa 594 goat anti-rabbit or Alexa 488 goat anti-mouse fluorophores, respectively. (C) Represents the nuclear staining (in pyramidal layer of hippocampus) in sections using Hoechst and (D) is a composite image of all the three stainings. The arrows indicate P-histone H3 positive cells in the brain section. (see Color Plate 2, following p. 322)

above under Section 3.1.1.1.1 for bright-field analysis. If preferred, double staining can be achieved in bright-field analysis as well, but this is not described here.

4. Notes 1. Sodium azide is toxic and care should be taken to avoid inhalation of the powder by wearing a mask. Paraformaldehyde is a carcinogen and caution should be taken when making the solution. Wear mask and gloves and prepare the solution in a fume hood. Do not overheat as heating the solution above 60C ruins the solution. The prepared solution is good for a week at 4C.


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2. Depending on the experiments (the number of parameters to be tested), cells can be grown in chamber slides with different number of wells 1, 2, 4, 8 or 16. This allows analysis of cells without dissociating them from the dishes, and comparative studies using multiple drugs can be achieved. 3. Do not allow the slides (containing cells or tissue sections) to dry during the whole-staining procedure. Drying will result in non-specific binding of antibodies and the results will not be reliable. Sealing the cover-glassed slides with nail polish after 24 h will prevent further drying and formation of air pockets in between the coverglass and slide. The given concentration of primary and secondary antibodies is the one that worked in our hand and they are working very well on different cell types and brain sections. If problems occur, each user may have to titer the antibody concentration that work well under their analysis conditions. 4. Not all type of slides work well when plating sections and it is very important to use the colorfrost or equivalent ones otherwise sections will not attach very well. If the sections are not dried well on the slides, during washes they can come off and therefore it is important to let them dry completely. Also care should be taken when spreading the sections onto the slide. Allow the sections to open up in the 30% EtOH solution on slides and using a thin paintbrush spread the sections. If the sections are going to be used for bright-field staining with antigen retrieval and dehydration steps, instead of rubber cement it is better to use ImmEdge Pen to create the barrier around the sections. Paraffin-embedded sections are not recommended for use with fluorescent methods as the sections may autofluoresce and hence make it difficult for actual detection of specific staining.

References 1. Hans, F., and Dimitrov, S. (2001). Histone H3 phosphorylation and cell division. Oncogene 20:3021–3027. 2. Peterson, C.L., and Laniel, M.A. (2004). Histones and histone modifications. Curr Biol 14:R546–551. 3. Johansen, K.M., and Johansen, J. (2006). Regulation of chromatin structure by histone H3S10 phosphorylation. Chromosome Res 14:393–404. 4. Koshland, D., and Strunnikov, A. (1996). Mitotic chromosome condensation. Annu Rev Cell Dev Biol 12:305–333.

5. Bradbury, E.M. (1992). Reversible histone modifications and the chromosome cell cycle. Bioessays 14:9–16. 6. Wei, Y., Mizzen, C.A., Cook, R.G., Gorovsky, M.A., and Allis, C.D. (1998). Phosphorylation of histone H3 at serine 10 is correlated with chromosome condensation during mitosis and meiosis in Tetrahymena. Proc Natl Acad Sci U S A 95:7480–7484. 7. Wei, Y., Yu, L., Bowen, J., Gorovsky, M.A., and Allis, C.D. (1999). Phosphorylation of histone H3 is required for proper

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chromosome condensation and segregation. Cell 97:99–109. Gurley, L.R., D’Anna, J.A., Barham, S.S., Deaven, L.L., and Tobey, R.A. (1978). Histone phosphorylation and chromatin structure during mitosis in Chinese hamster cells. Eur J Biochem 84:1–15. Mahadevan, L.C., Willis, A.C., and Barratt, M.J. (1991). Rapid histone H3 phosphorylation in response to growth factors, phorbol esters, okadaic acid, and protein synthesis inhibitors. Cell 65:775–783. Thomson, S., Mahadevan, L.C., and Clayton, A.L. (1999). MAP kinase-mediated signalling to nucleosomes and immediate-early gene induction. Semin Cell Dev Biol 10:205–214. Van Hooser, A., Goodrich, D.W., Allis, C.D., Brinkley, B.R., and Mancini, M.A. (1998). Histone H3 phosphorylation is required for the initiation, but not maintenance, of mammalian chromosome condensation. J Cell Sci 111 (Pt 23):3497–3506. Hendzel, M.J., Wei, Y., Mancini, M.A., Van Hooser, A., Ranalli, T., Brinkley, B.R., Bazett-Jones, D.P., and Allis, C.D. (1997). Mitosis-specific phosphorylation of histone H3 initiates primarily within pericentromeric heterochromatin during G2 and spreads in an ordered fashion coincident with mitotic chromosome condensation. Chromosoma 106:348–360. Nowak, S.J., and Corces, V.G. (2004). Phosphorylation of histone H3: a balancing act between chromosome condensation and transcriptional activation. Trends Genet 20:214–220. Prigent, C., and Dimitrov, S. (2003). Phosphorylation of serine 10 in histone H3, what for? J Cell Sci 116:3677–3685. Allis, C.D., and Gorovsky, M.A. (1981). Histone phosphorylation in macro- and micronuclei of Tetrahymena thermophila. Biochemistry 20:3828–3833.

16. Hendzel, M.J., and Bazett-Jones, D.P. (1997). Fixation-dependent organization of core histones following DNA fluorescent in situ hybridization. Chromosoma 106:114–123. 17. Crosio, C., Fimia, G.M., Loury, R., Kimura, M., Okano, Y., Zhou, H., Sen, S., Allis, C.D., and Sassone-Corsi, P. (2002). Mitotic phosphorylation of histone H3: spatiotemporal regulation by mammalian Aurora kinases. Mol Cell Biol 22:874–885. 18. Wood, M.A., Hawk, J.D., and Abel, T. (2006). Combinatorial chromatin modifications and memory storage: a code for memory? Learn Mem 13:241–244. 19. Wood, M.A., Kaplan, M.P., Brensinger, C.M., Guo, W., and Abel, T. (2005). Ubiquitin C-terminal hydrolase L3 (Uchl3) is involved in working memory. Hippocampus 15:610–621. 20. Alarcon, J.M., Malleret, G., Touzani, K., Vronskaya, S., Ishii, S., Kandel, E.R., and Barco, A. (2004). Chromatin acetylation, memory, and LTP are impaired in CBP+/– mice: a model for the cognitive deficit in Rubinstein-Taybi syndrome and its amelioration. Neuron 42:947–959. 21. Korzus, E., Rosenfeld, M.G., and Mayford, M. (2004). CBP histone acetyltransferase activity is a critical component of memory consolidation. Neuron 42:961–972. 22. Li, J., Guo, Y., Schroeder, F.A., Youngs, R.M., Schmidt, T.W., Ferris, C., Konradi, C., and Akbarian, S. (2004). Dopamine D2-like antagonists induce chromatin remodeling in striatal neurons through cyclic AMP-protein kinase A and NMDA receptor signaling. J Neurochem 90:1117–1131. 23. Crosio, C., Heitz, E., Allis, C.D., Borrelli, E., and Sassone-Corsi, P. (2003). Chromatin remodeling and neuronal response: multiple signaling pathways induce specific histone H3 modifications and early gene expression in hippocampal neurons. J Cell Sci 116:4905–4914.

Chapter 22 Chromatin Immunoprecipitation Assays: Analyzing Transcription Factor Binding and Histone Modifications In Vivo Smitha Pillai, Piyali Dasgupta, and Srikumar P. Chellappan Abstract Studies in the past decade have shown that differential gene expression depends not only on the binding of specific transcription factors to discrete promoter elements but also on the epigenetic modification of the DNA as well as histones associated with the promoter. While techniques like electrophoretic mobility shift assays could detect and characterize the binding of specific transcription factors present in cell lysates to DNA sequences in in vitro binding conditions, they were not effective in assessing the binding in intact cells. Development of chromatin immunoprecipitation technique in the past decade enabled the analysis of the association of regulatory molecules with specific promoters or changes in histone modifications in vivo, without overexpressing any component. ChIP assays can provide a snapshot of how a regulatory transcription factor affects the expression of a single gene or a variety of genes at the same time. Availability of high-quality antibodies that recognizes histones modified in a specific fashion further expanded the use of ChIP assays to analyze even minute changes in histone modification and nucleosomes structure. This chapter outlines the general strategies and protocols used to carry out ChIP assays to study the differential recruitment of transcription factors as well as histone modifications. Key words: Chromatin immunoprecipitation, histone modification, methylation, acetylation.

1. Introduction Chromatin immunoprecipitation is a powerful tool for identifying proteins, including transcription factors and histone proteins, associated with specific regions of genome, using specific antibodies that recognize the protein of interest. Chromatin immunoprecipitation assays provide an unbiased glimpse into the changes occurring on chromatin in vivo, in Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_22 Springerprotocols.com

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response to extracellular signals or during differentiation and development (1–3). It has long been established that differential gene expression correlates with the recruitment of specific and general transcription factors on relevant promoters; in addition, it has also become clear that covalent modification of histones in the promoter region also contributes significantly to the process (1, 4). Indeed, the pattern of acetylation and methylation of histones in the promoter region is so vital to modulating gene expression, it is thought that these processes code for the gene expression profiles per se (5–7). While modifications of histones and recruitment of transcription factors are dynamic events, it is possible to freeze the process chemically and examine the changes (8–10). ChIP assays provide an efficient, cost-effective means to achieve this. Studies from the laboratories of Doug Dean, Peggy Farnham and others in the 1990s established many of the protocols currently in use. Our lab has utilized ChIP assays to study the recruitment of E2F1, Rb, Raf-1, prohibitin or transcriptional co-repressors like HDAC1, HP1g, etc. to promoters, in response to specific signaling events. The protocol described here is an updated and modified version of an earlier protocol (11). Generally, ChIP assays can be done on tissue culture cells or from tissue samples. The basic assay protocol used to study histone modifications is shown in Fig. 22.1; this has been modified in certain situations. The initial step in the ChIP assay is to cross-link DNA to the associated proteins by treating the cells with formaldehyde (12–14); in certain situations, investigators have by-passed this step to prevent chemical modifications of the chromatin (4, 15). The second step is to isolate the DNA–protein complexes; the isolated DNA is then fragmented into 1–2 kb fragments, either by sonication (16–18) or by treatment with micrococcal nuclease (4, 19, 20), in the third step. The DNA fragments are then immunoprecipitated using specific antibodies of interest – antibodies to transcription factors, modified histones, co-activators or repressors, etc., which is the fourth step in the ChIP assay. The immunoprecipitated DNA/protein complexes are decross-linked in the fifth step; lastly, this is followed by PCR using specific primers to the promoters of interest in the sixth step. Alternately, the DNA can be used for probing microarrays of promoter regions to get an idea of global changes in factor recruitment and histone modifications (1, 13, 21–27) (See Note 1). Protocols for conducting such a ChIP on chip experiment and analyzing the data are described in the next two chapters.

Chromatin Immunoprecipitation Assays

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Fig. 22.1. The six basic steps in the ChIP assay protocol.

2. Materials 1. 37% formaldehyde (Molecular Biology Reagent Grade): formaldehyde is supplied as a 37% solution in water and can be purchased from Sigma-Aldrich (catalog # F-8775) or


other sources. Formaldehyde is a biohazard and needs to be handled inside a fume hood. 2. 1 M glycine: dissolve 3.75 g of tissue culture grade glycine (Fischer Biotech, catalog # BP381-1) in 50 ml of water to obtain a 1 M solution. 3. 20 PBS: dissolve 160 g NaCl, 4 g KCl, 2.88 g Na2HPO4, 4.8 g KH2PO4 in 1 l MilliQ water. Adjust the pH of the buffer to 6.4 to obtain 20 PBS stock solution. Take 50 ml of this buffer and make it up to 1 l with MilliQ water to obtain 1 PBS. 4. BSA (10 mg/ml): dissolve 100 mg of BSA (Sigma-Aldrich, catalog # A3059-50G) in 10 ml of MilliQ water. Aliquot in 100 ml and store at –20C 5. Salmon sperm DNA: Salmon sperm DNA is obtained from Sigma-Aldrich (catalog # D7696) as a 10 mg/ml solution. Use 10 ml for pre-clearing steps. 6. Herring sperm DNA: some labs use herring sperm DNA as a substitute for salmon sperm DNA. Herring sperm DNA is obtained from Sigma-Aldrich (catalog # D7290) as a 10 mg/ ml solution. Use 10 ml for pre-clearing. 7. Lysis buffer: 44 mM Tris–HCl (pH 8.1), 1% SDS, 10 mM EDTA, 10 mM sodium butyrate (for histone acetylation ChIP assays). Add the following protease inhibitors fresh to the lysis buffer just before use: 1 mM PMSF, 10 mg/ml aprotinin, 10 mg/ml leupeptin and 10 mg/ml pepstatin. 8. 100 mM PMSF: dissolve 0.174 g of PMSF (Sigma-Aldrich) in 10 ml of absolute ethanol. Aliquot into 1 ml in microcentrifuge tubes and store at –20C. Remember that the half-life of 1 mM PMSF (after it has been added to the lysis buffer) is 30 min, so add it just before use. 9. Dilution buffer: 16.7 mM Tris pH 8.1, 250 mM NaCl, 0.01% SDS, 1% Triton-X-100. 10. Wash buffer: 16.7 mM Tris–HCl pH 8.1, 375 mM NaCl, 0.01% SDS, 1% Triton-X-100. 11. Elution buffer: 0.1 M NaHCO3, 1% SDS, 5 mM NaCl. 12. Resuspension buffer: 50 mM Tris–HCl pH 8.0, 2 mM EDTA, 0.2% Sarkosyl. 13. IP Elution buffer: 50 mM NaHCO3, 1% SDS. 14. Proteinase K: dissolve 50 mg of proteinase K (Fischer Biotech, cat# BP-1700-100) in 2 ml of MilliQ water. Aliquot into 100 ml and store at –20C. 15. 5 PK buffer: 50 mM Tris–HCl pH 7.5, 25 mM EDTA, 1.25% SDS. 16. 3 M sodium acetate, pH 5.2: dissolve 20.4 g of sodium acetate in 50 ml of water. Adjust the pH to 5.2.


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17. Phenol–chloform–isoamyl alcohol: phenol–chloroform– isoamylalcohol is obtained from Fischer Biotech (catalog # BP1752-100). Phenol is a highly corrosive chemical and should be handled with extreme care. 18. 1 TE: 10 mM Tris pH 8.1, 1 mM EDTA. 19. Glycogen: glycogen is supplied as a 20 mg/ml solution from Roche (catalog # 901393). 20. RNase A: dissolve 0.1 g of RNase A (Fischer Biotech, catalog # BP2539-250) in 10 ml of water. Store at 4C. 21. 5 M NaCl: dissolve 58.4 g NaCl in 200 ml of MilliQ water. 22. MNase enzyme: MNase is obtained from Amersham Biosciences (catalog # E70196Y). Dissolve in MilliQ water containing 20–50% glycerol at a concentration of 1 mg/ml. Aliquots (10–20 ml) are stored at –20C and used only once after thawing on ice. 23. MNase digestion buffer: 0.32 M sucrose, 50 mM Tris–HCl pH 7.5, 1 mM CaCl2, 4 mM MgCl2, 0.1 mM PMSF. 24. Protein A (Amersham Biosciences, catalog # 17-0872-05) Protein G (Amersham Biosciences, catalog # 17-0618-01), Protein A/G beads (Santa Cruz Biotechnology, catalog # sc2003) or Staph A (American Type Culture Collection) cells.

3. Methods 3.1. Chromatin Immunoprecipitation (ChIP) Assay Protocol

3.1.1. Preparation of ChIP Cell Lysate

1. ChIP assays to analyze multiple factors or modifications will require a substantial amount of cells as the starting material. It is recommended that at least 2 107 adherent or suspension cells be used per sample (28–31) (See Note 2 and 3). Add formaldehyde directly to the tissue culture media to a final concentration of 1%. This corresponds to 280 ml of the commercially available 37% formaldehyde per 10 ml of tissue culture media. Incubate the cells with the formaldehyde on an orbital shaker for 20 min at room temperature for efficient cross-linking of DNA to the bound proteins. For histone acetylation studies, addition of histone deacetylase inhibitor sodium butyrate (20 mM) to the culture medium prior to fixation has been recommended to preserve histone lysine acetylation (32). 2. Terminate the cross-linking reaction by adding glycine to a final concentration of 0.125 M, using the 1 M stock solution. Incubate the cells on the orbital shaker for additional 5 min. 3. If the assay is performed on adherent cells, scrape the cells in 5 ml of 1 ice-cold PBS and transfer to a fresh 15 ml


centrifuge tube. In case of non-adherent cells, collect the cells along with the media and transfer to a 15 ml tube. Centrifuge the cells at 800 g (1000 rpm) for 5 min in a tabletop centrifuge (ThermoForma, General purpose centrifuge) to collect the cells. Wash the cell pellet twice with 10 ml of ice-cold PBS. Resuspend the pellet carefully in 1 ml 1 cold PBS and transfer it to an eppendorf tube. Use a sheared 1 ml tip for resuspending the cells. Spin at 800 g (3000 rpm) for 5 min in a refrigerated tabletop microcentrifuge at 4C (Beckman-Coulter, Microfuge R or an equivalent model). 4. Aspirate the supernatant carefully and resuspend the cell pellet in ice-cold lysis buffer containing protease inhibitors (for 75 ml cell pellet, add 200 ml lysis buffer). Mix well using a pipetman to obtain a uniform homogenate; avoid clumping of the cells. Do not vortex the lysate. The efficiency of lysis can be checked by trypan blue staining; if cells are not lysed they can be homogenized on ice using a Dounce homogenizer with B type pestle. Certain studies have used a glass beads disruption method to prepare ChIP lysates (29), as described in Alternate Protocols, Section 3.3.1. 5. Sonicate the lysate to shear the chromatin to below an average length of 1 kb. The time and the number of pulses vary according to cell type, the sonicator used and extent of cross-linking (See Note 4). However, we have found for most cells, three cycles of sonication at power 4 for 30 s on a Fisher Sonic Dismembrator (Model 100), followed by incubation on ice for 30 s, are sufficient to shear the chromatin properly. As an alternative to sonication, digestion with micrococcal nuclease (4, 15, 19) can be also done to cleave the chromatin into oligonucleosomes (refer to Alternate Protocols, Section 3.3.2). This method is referred to as ‘‘native chromatin immunoprecipitation’’. However, this technique is incompatible with formaldehyde cross-linking since cross-linked chromatin is inefficiently cleaved by nucleases. The protocol for cell lysis and nuclei preparation in native ChIP assays is described in (4, 15, 19). 6. Spin at 15,000 rpm in a tabletop refrigerated microcentrifuge for 15 min. Carefully transfer the supernatant into a new microcentrifuge tube. At this point the chromatin can be snap frozen in liquid nitrogen and stored at –70C, if needed. 3.1.2. Setting up the Immunoprecipitation (IP) Reaction

7. If previously frozen, thaw out the ChIP lysate on ice. Save 20% of the lysate as the input control. Keep the input control aside and do not use it for the immunoprecipitation reaction. The rest of the lysate is equally divided into


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three microcentrifuge tubes. Set up the following immunoprecipitation reactions: (1) Irrelevant antibody (2) Positive control antibody (3) The antibody that is specific to the protein of interest. 8. Use 3 mg of antibody per immunoprecipitation reaction. Add 300 ml of dilution buffer and rotate the mixture overnight at 4C. Pre-clearing ChIP lysates is a good method to reduce non-specific backgrounds and eliminate false positives during the assay (for pre-clearing protocols see Support Protocols, Section 2.2). While it may not be necessary to pre-clear in all cases, it should be carried out if non-specific background bands are observed in PCR. See Note 5 9. ChIP assays can be performed with protein A, protein G, protein A/G beads or Staph A cells. The choice of beads depends on the isotype of the antibody to be used for the immunoprecipitation. Add the 120 ml of 1:1 protein G slurry or 400 ml of Protein A slurry (for preparation of beads or Staph A, refer to Support Protocols Section 3.2.1). Thereafter, make up the volume to 700 ml with dilution buffer and rotate for 2 h at 4C. If you are using Staph A beads, incubate each sample with 10 ml of pre-blocked Staph A beads (10, 30, 31, 33) (see Support Protocols, Section 3.2.2.1). Rotate on a nutator at 4C for 15 min. 3.1.3. Extraction of Immunoprecipitated DNA

10. Spin the tubes from Step 8, Section 3.1.2 at (600 g) 2500 rpm at room temperature for 30 s, in a tabletop microcentrifuge at room temperature. Carefully aspirate the supernatant and discard. 11. Efficient washing is critical to reduce background. Wash the beads three times with dilution buffer and twice with wash buffer. It has been found that transferring the beads to a new tube after the first wash helps reduce non-specific bands. Remove the supernatant and discard each time; care should be taken not to lose beads during the wash steps. Carefully remove all the supernatant using a gel-loading tip in the final wash. See Note 6. 12. Add 300 ml of elution buffer. Rotate for 15 min at room temperature. 13. Spin the tubes at 2000 g (5000 rpm) for 2 min at room temperature, in a tabletop microcentrifuge. 14. Transfer the supernatant to fresh tubes and incubate 65C overnight to reverse formaldehyde cross-linking. The tube containing the input DNA (Section 3.1.2, Step 7) should also be incubated at 65C overnight.


15. Add 0.1 volumes of 3 M sodium acetate pH 5.2, followed by two and a half volumes of ice-cold 100% ethanol to the tubes and incubate at –70C for 10 min to precipitate the DNA. 16. Centrifuge the samples at 17,000 g (14,000 rpm) for 15 min at 4C, in a refrigerated microcentrifuge. 17. Wash the pellet with 2.5 volumes of ice-cold 70% ethanol. Spin the tubes at 14,000 rpm for 15 min at 4C. Air dry the pellet. 18. Resuspend the DNA in 200 ml of MilliQ water. Add 50 mg of proteinase K (2 ml from a stock solution of 25 mg/ml proteinase K, to each sample) and incubate at 37C for 30 min. 19. Extract the DNA by adding equal volumes of phenol– chloroform–isoamyl alcohol. Vortex well to mix the layers. Spin the tubes at 17,000 g (14,000 rpm) for 15 min at 4C, in a refrigerated microcentrifuge to separate the layers. Collect the upper layer carefully using a pipetman, transfer to a fresh tube and repeat steps 14–16 to extract the DNA. Alternately, use a QiaQuick spin column (Qiagen Corporation) to extract DNA and elute the DNA in 50 ml of TE. 20. Resuspend the immunoprecipitated DNA samples in 50 ml of water. The input DNA is resuspended in 250 ml water. The DNA of interest is amplified by PCR using specific primers to the sequence of interest. Use 5 ml of immunoprecipitated DNA for PCR analysis; 1 ml of the input DNA would be sufficient for PCR. PCR conditions vary according to the primers used; generally 30 cycles of amplification will result in visible bands. Analyze the DNA fragments by agarose gel electrophoresis. See Note 7. 3.2. Support Protocols for ChIP Assay

3.2.1. Preparation of Protein A, G, A/G Beads or Staph A Cells

Appropriate beads are chosen depending on the isotype of the antibodies used and their affinity for protein A, protein G and protein A/G. Protocols from the Farnham lab use Staph A cells for immunoprecipitation reaction. We use protein G or protein A for most applications in our laboratory. 1. Protein A: weigh out 60 mg of protein A and resuspend it in 2 ml of dilution buffer. Incubate for 30 min on ice. Subsequently, use 400 ml of the above slurry for each ChIP reaction. 2. Protein G: we use protein G slurry containing 20% ethanol. The required volume of beads are taken and washed three times with dilution buffer to remove the ethanol preservative. Subsequently, the beads are resuspended in equal volume of dilution buffer and 120 ml of beads are used per sample. The same procedure can be also used for preparation of protein A/G beads.


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3. Staph A cells: resuspend 1 g of Staph A cells in 10 ml of 1 resuspension buffer (8, 33). Centrifuge at 10,000 rpm for 5 min at 4C. This step is repeated once again. Thereafter resuspend in 3 ml of 1 PBS containing 3% SDS and 10% ME. Boil the beads for 30 min and collect by centrifugation at 1000 rpm for 5 min. Wash twice in 1 resuspension buffer and repeat centrifugation at 10,000 rpm for 5 min. Resuspend in 4 ml of resuspension buffer, aliquot in 100 ml volumes in microcentrifuge tubes and snap freeze. 3.2.2. Pre-clearing of ChIP Lysates

Pre-clearing lysates is a good method to eliminate non-specific background in ChIP assays. ChIP assay protocols using Staph A cells require an initial preblocking step to reduce non-specific binding (30, 34).

3.2.2.1. Preblocking Staph A Beads

For every tube (100 ml) of Staph A cells add 10 ml herring sperm DNA (10 mg/ml) and 10 ml BSA (10 mg/ml). Incubate on a nutator at 4C for at least 3 h or more (overnight incubation can also be done). Centrifuge at 10,000 rpm for 3 min. Remove supernatant and wash twice with 1 resuspension buffer. Resuspend the cells in equal volume of resuspension buffer and use 10–15 ml for each pre-clearing step.

3.2.2.2. Pre-clearing Protocols with Protein A, Protein G or Protein A/G

Add 120 ml of 1:1 protein G slurry (or 400 ml of protein A) to the ChIP lysate (obtained from 2 107 cells as mentioned previously in Section 3.1.1) in a microcentrifuge tube. Add 15 ml of salmon sperm DNA (10 mg/ml) and 10 ml of BSA (10 mg/ml). Rotate on a nutator for 30 min at 4C. Thereafter spin the tubes at 14,000 rpm for 30 s in a tabletop centrifuge at 4C. Collect supernatant fraction and discard the beads. This supernatant fraction is used for the immunoprecipitation reaction described in Section 3.1.2.

3.2.2.3. Pre-clearing Protocols Using Staph A Cells

Use the pre-blocked Staph A cells as described in Section 3.2.1 to pre-clear the ChIP lysate. Ten to fifteen microliter of pre-blocked Staph A beads are added to the ChIP lysate and incubated on a nutator for 15 min at 4C. Centrifuge the tube at 14,000 rpm for 5 min and transfer the supernatant to a fresh tube. Use this precleared lysate for the subsequent immunoprecipitation reaction (Section 3.1.2).

3.2.3. DNA Extraction from Immunoprecipitates Using Staph A Beads

1. The immunoprecipitation reaction is set up as detailed in Section 3.1.2, Steps 7–8 2. Centrifuge the samples at 14,000 rpm for 3 min at room temperature. 3. Wash the pellets twice with 1.4 ml of 1 resuspension buffer followed by four washes with IP buffer. The washes are done


by first dissolving the Staph A pellet in 200 ml of the appropriate buffer and adding 1.2 ml of the same buffer. The tubes are incubated with the wash buffer on a nutator for 3 min at room temperature. Thereafter, the tubes are centrifuged at 14,000 rpm for 3 min. The wash buffer is carefully aspirated off. After the last wash, the entire buffer is removed using a gel-loading tip. 4. Elute the immunocomplex by adding 150 ml of IP elution buffer. Mix very well for about 15 min. Microcentrifuge the tubes at 14,000 rpm for 3 min. Transfer the supernatants to fresh tubes. 5. Repeat the elution step. Both the elution fractions are pooled. Spin the tubes at 14,000 rpm for 5 min at room temperature. Transfer the supernatants to fresh tubes. Add 1 ml of 10 mg/ ml RNaseA and NaCl to a final concentration of 0.3 M. Incubate samples at 67C for 4–5 h (or overnight) to reverse cross-linking between protein and DNA. Take the input DNA sample and incubate at 65C for 4–5 h (or overnight) to reverse cross-linking. 6. Precipitate the DNA by adding 2.5 volumes of ice-cold ethanol to each of the tubes and incubate at –20C overnight. 7. Centrifuge the DNA at 14,000 rpm for 15 min at 4C. Remove the residual ethanol and air dry the DNA pellet. 8. Dissolve the pellet in 100 ml TE. The sample may be viscous and may have to be dissolved in a larger volume. Add 25 ml of 5 PK buffer and 1.5 ml of proteinase K to each sample. Incubate for 1–2 h at 45C. 9. Add 175 ml of TE to each sample. Extract with 300 ml of phenol–chloroform–isoamyl alcohol. 10. Add 30 ml of 5 M NaCl, 5 mg of tRNA and 5 mg of glycogen to each sample. Mix well and then add 750 ml of ice-cold ethanol. Incubate at –20C overnight. 11. Next day, centrifuge the tubes at 14,000 rpm for 20 min at 4C. Aspirate the supernatant carefully and air dry the DNA pellet. Resuspend the DNA pellet in 30 ml of TE. Use 2–3 ml for PCR analysis. For the total input sample, dilute the sample 300-fold and then perform the PCR reaction (30, 31). 3.3. Alternate Protocols for ChIP Assay

1. Perform steps 1–3 as detailed in Section 3.1.1 2. Resuspend the pellet in equal volume 200 ml of ChIP lysis buffer 3. Add an equal volume of glass beads.

3.3.1. Preparation of ChIP Lysate

4. Vortex at the maximum setting six times for 30 s at 4C each with 3 min pauses between each vortex.


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5. Pierce the bottom of the microcentrifuge tube with a needle and place the tube carefully in another microcentrifuge tube. Spin at 700 g for 2 min. The liquid should move through the first microcentrifuge tube into the bottom tube. 6. Add 200 ml of ChIP lysis buffer and sonicate the samples as described in Section 3.1.1, steps 5–6.

3.3.2. Fragmentation of Chromatin Using Micrococcal Nuclease (MNase)

1. Prepare nuclear pellet from cells as detailed in Refs. 9 and 36. 2. Resuspend the nuclear pellet in 1 ml MNase digestion buffer and place on ice. 3. Aliquot two 1.5 ml eppendorf tubes with 500 ml of resuspended nuclei. 4. Add 1 ml of micrococcal nuclease (MNase) enzyme to each tube and mix gently. 5. Incubate the tubes in a 37C water bath for 10 min. The incubation times as well as MNase concentrations have to be optimized to generate mainly tri-, di- and mononucleosomes. 6. Add 20 ml of stop solution. Chill on ice. 7. The suspension was centrifuged at 13,000 g for 20 s. The supernatant comprised of the second soluble fraction S2 (containing larger fragments of chromatin), whereas the pellet contained the smaller fragment S1 fraction. The pellet was resolubilized in lysis buffer and pooled with the S2 fraction in a fresh clean tube. 8. Continue the ChIP assay as described in Section 3.1 from Step 6 onwards.

3.4. ChIP Assay on Tissues

1. Chop the tissues with a razor blade or scalpel into small pieces and resuspend them in serum-free media. Add 1% formaldehyde (some studies have used 0.4% formaldehyde) and rotate the tubes for 10 min at room temperature. 2. Stop the cross-linking reaction by adding glycine to a final concentration of 0.125 M. Rotate at room temperature for 5 min. 3. Centrifuge the samples at 250 g for 5 min and discard the supernatant. Wash the sample twice in PBS. Resuspend the chopped tissue in 1–2 ml PBS. Homogenize the tissue on ice using a Dounce homogenizer or Polytron Homogenizer. Centrifuge the sample at 250 g to collect all the cells; discard the supernatant. 4. Follow the standard ChIP protocol as detailed in Section 3.1.1, Step 5 onwards (10, 35, 36).


4. Notes 1. Chromatin immunoprecipitation is a powerful approach that allows the elucidation of interactions between endogenous proteins and their native chromatin sites, thereby providing insights into physiological transcription (37–42) and transactivation mechanisms (3, 43, 44). ChIP assays have been extensively used to study histone acetylation and methylation patterns at a variety of genomic loci (2, 5, 9, 34, 43, 45). In addition, ChIP methodology has been coupled to microarrays (ChIP-chip) containing genomic regions to facilitate genomewide binding patterns of transcription factors to specific promoters (1, 2, 25). Procedures by Ren et al. (2004) involve immunoprecipitation of the DNA by ChIP assays is blunt ended by T4 DNA polymerase to allow ligation of a universal linker (LM-PCR) (13, 46). The ChIP-enriched DNA and input DNA are then labeled with different fluorescent dyes and hybridized to a single DNA microarray containing promoter sequences (for detailed description of the method refer to (9, 13, 22, 23, 25, 46)). Despite the many advantages of the ChIP assay, it can be fraught with many obstacles. The first important thing to keep in mind is to make the lysates using large number of cells (at least 2 107) to achieve a good signal-to-noise ratio. Efficient lysis of cells is another important parameter for a successful ChIP assay. It may be a good idea to check the efficacy of cell lysis using trypan blue exclusion method. In case cells are not being efficiently lysed, use a different lysis buffer. A list of commonly used lysis buffers are given below. Other lysis buffers which have been reported in literature: 1. http://genomecenter.ucdavies.edu/farnham/farnham/ protocols/chips.html (30). 5 mM PIPES pH 8.0, 85 mM KCl, 0.5% NP-40, protease inhibitors. 2. 50 mM Tris pH 8.1, 150 mM NaCl, 5 mM EDTA, 0.1% sodium deoxycholate, 1% Triton-X-100, protease inhibitors (29). 3. http://www.biochem.northwestern.edu/ibis/morimoto/ Protocols/ 50 mM Tris–HCl pH 8.0, 150 mM NaCl, 1% SDS, 2% Triton-X-100, 1 mM EDTA, protease inhibitors (47). 4. 250 mM sucrose, 10 mM Tris–HCl pH 7.4, 10 mM Na– butyrate, 4 mM magnesium chloride, 0.1 M PMSF, 0.1 M benzamidine, 0.1% w/v Triton-X-100, protease inhibitors (15). Moreover, cells can be homogenized on ice after addition of cell lysis buffer to facilitate the release of nuclei. In case of heterogeneous samples, it is essential that the cells be sorted


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to enrich the cell population of interest before performing the ChIP assay. To analyze histone acetylation, add 10 mM sodium butyrate to the lysis buffer, dilution and elution buffers. Sodium butyrate prevents the loss of histone acetylation during sample preparation by inhibition of histone deacetylase activity. 2. It is essential to perform all steps of cell lysis and immunoprecipitation on ice or at 4C, wherever indicated to prevent chromatin degradation by endogenous nucleases. Similarly, protease inhibitors should be added wherever mentioned to preserve the integrity of proteins bound to the chromatin. 3. Another crucial step in the ChIP assay is the sonication of DNA. It is essential that sonication should cause efficient shearing of the genomic DNA to about 1 kb in size. The efficiency of sonication should be checked by agarose gel elecrophoresis when the ChIP assays are being standardized with new cell lines or sonicators. It must be remembered that sonication of cross-linked chromatin gives rise to randomly sheared DNA fragments, which may not produce small enough chromatin fragments at the region of interest. Furthermore, crosslinking the DNA to the protein by formaldehyde may mask critical protein epitopes resulting in poor efficiency of immunoprecipitation. On the other hand, a disadvantage of native ChIP is that certain protein–DNA interactions are transient or intrinsically unstable, precluding detection unless the chromatin–DNA complexes are trapped by cross-linking. The selection of conventional ChIP assay versus native ChIP assay depends upon the nature of the experimental system. Whereas the native ChIP is particularly suited for proteins like core histones that are tightly bound to chromosomes, the conventional ChIP assay works better for studying linker histones, transcription factors and proteins which may relocalize or dissociate during sample preparation. 4. Although, the use of PCR ensures a high degree of signal amplification, a low efficiency in the immunoprecipitation reaction can give rise to non-specific signals or no signal. Hence, it is essential to use 2–4 mg of a high-affinity, high specificity antibody for the immunoprecipitation reaction. Pre-clearing the lysates before immunoprecipitation is a good way to improve specificity and minimize non-specific background in the ChIP assay (Methods, Section 3.2.2). However, we have found that even in the absence of pre-clearing, the ChIP assay works provided all other safeguards are carefully followed. Furthermore, one may have to try different antibodies to optimize the ChIP assay because certain antibodies may efficiently immunoprecipitate the native protein but may be not be effective in immunoprecipitating proteins cross-linked


to chromatin. The exact conditions of the immunoprecipitation may have to be further optimized to ensure a high recovery of target DNA during the IP. It is crucial to incorporate a positive control as well an irrelevant antibody in the ChIP assay to facilitate critical interpretation of data. 5. After the immunoprecipitation reaction, efficient and stringent washing is critical to reduce non-specific background. The number of washes can be increased if necessary. The wash buffer can be varied according to the nature of antibody or conditions of immunoprecipitation. Some laboratories use 2 PBS as the wash buffer, others wash the immunocomplexes sequentially with 1 ml of the following buffers: low salt wash buffer, high salt wash buffer, LiCl wash buffer and 1 TE. A list of wash buffers used by different laboratories is given below: 1. http://genomecenter.ucdavies.edu/farnham/farnham/protocols/chips.html (30). 100 mM Tris–HCl pH 9.0 (8.0 for monoclonal antibodies), 500 mM LiCl, 1% NP-40, 1% sodium deoxycholate. 2. 250 mM sucrose, 10 mM sodium butyrate, 4 mM magnesium chloride, 0.1 mM PMSF (15). 3. Low salt wash buffer: 20 mM Tris–HCl pH 8.1, 150 mM NaCl, 0.1% SDS, 1% Triton-X-100, 2 mM EDTA (30, 48). 4. High salt wash buffer: 20 mM Tris–HCl pH 8.1, 500 mM NaCl, 0.1% SDS, 1% Triton-X-100, 2 mM EDTA. 5. LiCl wash buffer: 10 mM Tris–HCl pH 8.0, 0.25 mM LiCl, 1% NP-40, 1% deoxycholate, 1 mM EDTA. 6. It is important to be extremely careful and meticulous during this assay. The amplification by PCR is highly sensitive; it will amplify even minimal non-specific signals. We recommend all possible precautions to prevent contamination from other DNA sources; amplification in a dedicated space, use of fresh gloves, sterile filter tips and clean microfuge tubes for the assay. 7. Recent studies involving ChIP assays have used real-time PCR for detecting as little as 2-fold changes in protein–DNA interactions. Real-time PCR allows data to be collected during cycles when amplification is occurring exponentially (49). It increases the precision of ChIP measurements allowing accurate detection of small variations in protein–DNA interactions (32). Another possibility is to perform duplex amplification PCR (50, 51), which involves co-amplification of a fragment from the region of interest along with a control fragment. Applications, in particular for allelic studies, or on genomic imprinting in mammals, one can apply SSCP (52, 53) or


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hot-stop PCR (54), especially to differentiate silent alleles from the active alleles. The different PCR-based approaches have been reviewed in (3, 4, 55).

Acknowledgments Work in the Chellappan lab is supported by the grants CA63136, CA77301 and CA127725 from the NCI.

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Chapter 23 ChIP on Chip Assays: Genome-Wide Analysis of Transcription Factor Binding and Histone Modifications Smitha Pillai and Srikumar P. Chellappan Abstract Deregulation of transcriptional activity of many genes has been causatively linked to human diseases including cancer. Altered patterns of gene expression in normal and cancer cells are the result of inappropriate expression of transcription factors and chromatin-modifying proteins. Chromatin immunoprecipitation assay is a well-established tool for investigating the interactions between regulatory proteins and DNA at distinct stages of gene activation. ChIP coupled with DNA microarrays, known as ChIP on chip, allow us to determine the entire spectrum of in vivo DNA-binding sites for a given protein. This has been of immense value because ChIP on chip assays can provide a snapshot of the transcriptional regulatory mechanisms on a genome-wide scale. This article outlines the general strategies used to carry out ChIP-chip assays to study the differential recruitment of regulatory molecules based on the studies conducted in our lab as well as other published protocols Key words: Genome-wide chromatin immunoprecipitation, transcription factors, histone modifications, promoter arrays, whole genome amplification.

1. Introduction DNA-binding proteins perform a variety of important functions in cells, including transcriptional regulation, chromosome maintenance, replication and DNA repair. The interactions between transcription factors and their DNA binding sites are an integral part of transcriptional regulatory networks. These interactions control the coordinated expression of thousands of genes during normal growth and in response to external stimuli. Studies on the sequence-specific binding of transcription factors to promoter DNA were provided a major boost by the development of electrophoretic mobility shift assays (EMSAs or gel-shift assays) three Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_23 Springerprotocols.com

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decades ago (1, 2). While this technique provided the capability to analyze the binding of proteins to DNA in vitro, it could not provide a snapshot of the binding status in the intact cells. Development of chromatin immunoprecipitation assays (ChIP assays), where DNA bound proteins are cross-linked to DNA in the living cells and analyzed by PCR provided the ability to examine the binding of proteins to specific targets in vivo in response to specific signals or during normal development. Further, these assays also enabled the analysis of histone modifications and contributed significantly to the elucidation of the histone code and the epigenetic regulation of gene expression. Given the ability of chromatin immunoprecipitation assays to provide a picture of the promoter occupancy of protein in an in vivo situation, attempts were made to extend this technology to genome-wide analysis of DNA-binding and histone modifications (3, 4). Recent advances in DNA microarray technology as well as the availability of whole genome sequences make it possible to map the entire spectrum of interactions of a particular protein with specific DNA sequences across the whole genome. This is achieved by an immunoprecipitation of the protein of interest cross-linked to its target DNA sequences; the bound fragments are identified by a microarray-based analysis of enriched DNA fragments. This technique, referred to as ChIP on chip or genome-wide location analysis, is currently the most widely used method for identifying in vivo transcription factorbinding sites in a high-throughput manner (5, 6). In conventional ChIP assays, the cells are treated with formaldehyde to covalently cross-link protein and the bound DNA (7, 8); the cells are then lysed and the chromatin is sheared into 200–1000 base pair fragments by sonication (7) or treatment with micrococcal nucleases (9, 10). An antibody specific for a protein of interest is then used to immunoprecipitate protein bound DNA fragments; the associated DNA is de-cross-linked by incubating in a low pH buffer, protease treated and the associated DNA is examined for the presence of the promoter or DNA sequence of interest using appropriate PCR primers (11). This technique, as can be imagined, used to analyze the association of specific transcription factors, or modified histones with DNA. The ChIP-on-chip version of this protocol differs only in the postIP steps. In this case, the immunoprecipitated DNA is amplified using a random primed PCR or the immunoprecipitated DNA is blunt ended by T4 DNA polymerase to allow ligation of a universal linker (LM-PCR) or whole genome amplification method (WGA) and subsequently labeled and hybridized into DNA microarrays (12–15). The extent of ChIP-chip application depends, in part, on the development of microarray technology, especially the availability of arrayed slides for DNA from human, mouse or other model

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organisms. There are two different types of DNA microarrays available; the first type is created by spotting DNA onto polylysine-modified glass slides using pin-spotting robots (16). The density of spotted arrays is limited to 20,000–30,000 features since the feature sizes below 100 microns are difficult to achieve (17). A different type of oligonucleotide array is available which is created by synthesizing oligomers in situ on the array by photolithography. These chips marketed by Affymetrix can be of higher density containing up to several million features (18). Today, arrays representing promoter regions, CpG islands, entire chromosomes or entire genomes are used for ChIP-chip assays (19–22). High-density oligonucleotide arrays that tile the entire human non-repetitive genomic sequences are available from Affymetrix, NimbleGen Systems and Agilent Technologies (See Notes 1 and 2). The major applications of ChIP-chip assays are the identification of binding targets for transcription factors and identification of post-translational modifications on DNA-associated proteins including specific histone modifications. Initial studies using ChIP-chip technology were directed to detect transcription factor chromatin binding in the yeast Saccharomyces cerevisiae (23–26). ChIP-chip technology was later applied to mammalian systems to study the recruitment of transcription factors such as E2F, GATA1, Rb, etc., but these studies surveyed only a small fraction of the genome due to the limitations in the microarray technology and also the incomplete information on genome annotation (21, 27, 28). With the recent advances in DNA microarray synthesis technology and the completion of human genome project revealing more promoter regions, allowed extensive screening of the genome for the binding position of protein factors. In recent years ChIP-chip has been expanded to investigate post-translational modifications of DNA-binding proteins. Different strategies can be employed to study histone modifications using ChIP-chip technology. One is to detect the distribution of histone modifications using antibodies specially targeting the modifications (29). and the other is using ChIP-chip to locate the enzymes that catalyze the histone modification reactions (30). Another useful application of these kinds of studies would be to identify genes silenced by histone modification in carcinogenesis (20). It is also possible to combine the ChIP-chip data with expression profiling to establish the correlation of histone markers with transcriptional activity (29, 31). ChIP-on-chip assays can be done on tissue culture cells or from tissue samples. The assay involves multiple steps, each of which might require standardization based on the specific experimental conditions and antibody used (Fig. 23.1). We have utilized ChIP-chip assays to study the global analysis of binding of proteins of interest to promoter regions in response to specific

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Fig. 23.1. Schematic of ChIP on chip workflow. (A) Chromatin immunoprecipitation. Cells are grown under the desired experimental conditions and protein–DNA cross-links are created by treating the cells with formaldehyde. Cells are lysed, DNA is sheared either by sonication or by micrococcal nuclease and immunoprecipitated with antibody against protein of interest. Cross-links are reversed and DNA is purified. (B) ChIP-chip experiment. The immunoprecipitated DNA is amplified either by random primed PCR or by ligation-mediated PCR or whole genome amplification. The amplified DNA is fragmented, labeled with a fluorophore, and hybridized to a microarray and analyzed.

signaling events, using Affymetrix promoter arrays (GeneChip human promoter 1.0R array, Affymetrix, Cat. No. 900775). The protocol we have successfully used is as follows:

2. Materials 1. Formaldehyde (37%) (Molecular Biology Reagent Grade; Sigma-Aldrich). 2. Glycine, 1 M: 3.75 g of tissue grade glycine (Fischer Biotech) in 50 ml of water. 3. 20 phosphate-buffered saline (PBS): 160 g NaCl, 4 g KCl, 2.88 g Na2HPO4, 4.8 g KH2PO4 in 800 ml of water. Adjust the pH of the buffer to 6.4 and make up the volume to 1 l.

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4. IGEPAL CA-630 (Sigma-Aldrich). 5. 100 mM PMSF: 0. 174 g of PMSF (Sigma-Aldrich) in 10 ml of absolute ethanol. Aliquot into microcentrifuge tubes and store at –20C. Add PMSF to the lysis buffer just before use as the half-life of 1 mM PMSF in the buffer is 30 min. 6. Protease inhibitor stock: prepare a 25 stock by dissolving one protease inhibitor tablet (Roche, cat. no. 11873580001) in 2 ml of nuclease-free water. 7. Proteinase K (Fischer Biotech, cat. no. BP-1700-100). 8. LiCl (10 M): 42.39 g LiCl (Sigma-Aldrich cat. no. L-8895) in 100 ml of water. 9. Protein G Sepharose 4 Fast Flow (GE Healthcare, cat. no. 170618-01). 10. Sequenase Version 2.0 DNA polymerase (USB, cat. no. 70775Y). 11. Primer A: 200 mM GTTTCCCAGTCACGGTC(N)9 (HPLC Purified). 12. Primer B: 100 mM GTTTCCCAGTCACGGTC (HPLC Purified). 13. Taq Polymerase 5 u/ml (Invitrogen, cat. no. 10342-020). 14 100 mM dNTPs (Promega, dATP- cat no. U120-B, dCTP- cat no. U121B, dGTP- cat no. U122B, dTTP- cat no. U123B,) 15. 100 mMdUTP (Promega, cat. no. U119A). 16. BSA (20 mg/ml): 100 mg of BSA (Sigma-Aldrich cat. no. A3059-50G) in 5 ml of water. Aliquot 100 ml into 1.5 ml tubes and store at –20C. 17. 1 M DTT (dithiothreitol): 1.54 g DTT (Fisher Biotech cat. no. BP172-5) in 10 ml of water. Aliquot 100 ml into 1.5 ml tubes and store in –20C. 18. 5 M NaCl: 58.4 g NaCl in 200 ml of water. 19. 1 M MgCl2: 20.3 g of MgCl2.6 H2O in 100 ml of water. 20. 1 M CaCl2: 14.7 g of CaCl2. 2 H2O in 100 ml water. 21. Deoxycholate (sodium salt) (Sigma-Aldrich cat. no. 21115). 22. GeneChip WT Double-Stranded DNA Terminal Labeling Kit (Affymetrix, cat. no. 900812). 23. GeneChip sample clean up module (Affymetrix, cat. no. 900371). 24. Control oligonucleotide B2, 3 nM (Affymetrix, cat. no. 900301). 25. GeneChip hybridization, wash, and stain kit (Affymetrix, cat. no. 900301).

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2.1. Buffers

1. Lysis buffer: 10 mM Tris–HCl (made from stock 1 M Tris–HCl pH 7.5), 10 mM NaCl 3 mM MgCl2, 1 mM CaCl2, 4% IGEPAL, 1 mM PMSF (add fresh). Store at room temperature. 2. IP dilution buffer: 20 mM Tris–HCl (made from stock 1 M Tris–HCl pH 8), 2 mM EDTA 1% Triton X-100, 150 mM NaCl, protease inhibitor stock (add fresh). Store at room temperature. 3. Protease inhibitor stock: prepare a 25 stock by dissolving one protease inhibitor tablet (Roche, cat. no. 11873580001) in 2 ml of nuclease-free water. Store the aliquots at –20C. 4. ChIP wash 1: 20 mM Tris–HCl (made from stock 1 M Tris–HCl pH 8), 2 mM EDTA, 1% Triton X-100, 150 mM NaCl, and 1 mM PMSF (add fresh). Store at room temperature. 5. ChIP wash 2: 20 mM Tris–HCl (made from stock 1 M Tris–HCl pH 8), 2 mM EDTA 1% Triton X-100, 0.1% SDS, 500 mM NaCl, and 1 mM PMSF (add fresh). Store at room temperature. 6. ChIP wash 3: 10 mM Tris–HCl (made from stock 1 M Tris–HCl pH 8), 1 mM EDTA 0.25 M LiCl, 0.5% IGEPAL, and 0.5% deoxycholate (sodium salt). Store at room temperature. 7. TE (10 mM Tris–HCl pH 8, 1 mM EDTA). 8. Elution buffer: 25 mM Tris–HCl (made from stock 1 M Tris–HCl pH 7.5), 10 mM EDTA, and 0.5% SDS. Store at room temperature. 9. Buffers for hybridization: A. 12 MES stock buffer (1.22 M MES, 0.89 M Na+): ME Shydrate (64.61 g), MES sodium salt, (193.3 g), MilliQ water (800 ml). Adjust the pH between 6.5 and 6.7. Adjust the volume to 1000 ml. Do not autoclave. Filter sterilize by passing through 0.2 mm filter. Store at 2–8C away from light. B. 2 hybridization buffer (100 mM MES, 1 M [Na+], 20 mM EDTA, 0.01% Tween-20): 12 MES stock buffer (8.3 ml), 5 M NaCl (17.7 ml), 0.5 M EDTA (4 ml), 10% Tween 20 (100 ml). Make up the volume to 50 ml with water and store at 2C–8C away from light. C. Wash buffer A: non-stringent wash buffer (6 SSPE, 0.01% Tween 20): 20 SSPE (300 ml), 10% Tween 20 (1 ml). Make up the volume to 1 l and filter sterilize using 0.2 mm filter.

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D. Wash buffer B (100 mM MES, 0.1 M Na+, 0.01% Tween 20): 12 MES stock buffer (83.3 ml), 5 M NaCl (5.2 ml), 10% Tween 20 (1 ml). Make up the volume to 1 l, filter through 0.2 mm filter and store at 2–8C protected from light. E. 2 stain buffer (100 mM MES, 1 M Na+, 0.05% Tween 20): 12 MES stock buffer (41.7 ml), 5 M NaCl (92.5 ml), 10% Tween 20 (2.5 ml). Make up the volume to 250 ml, filter through 0.2 mm and store at 2–8C protected from light. 2.2. Instruments and Software Required

1. Genechip Scanner 3000 7G 2. GeneChip Operating Software (GCOS) v1.3 or higher 3. GeneChip Fluidics Station 400 or 450 4. Tiling array software for data analysis

3. Methods 3.1. Preparation of ChIP Lysate from Cells

1. ChIP assays require large amounts of cells as the starting material. Usually, for each IP reaction 50–200 million cells are recommended (32–34). For a ChIP on chip experiment at least two IP reactions should be done; an experimental sample where IP is done using antibody against the protein of interest and a negative control with a nonspecific antibody or a mock IP with no antibody. Add formaldehyde directly to the tissue culture plate to a final concentration of 1% (280 ml of 37% formaldehyde/10 ml of media) and incubate in an orbital shaker for 10 min for cross-linking DNA to the protein (See Note 3). 2. Add glycine to a final concentration of 0.125 M using a 1 M stock solution and inoculate for 5 min in the orbital shaker to terminate the cross-linking. 3. Aspirate the formaldehyde and media into a waste container and wash the cells twice with 10 ml of ice-cold 1 PBS. In case of adherent cells, scrape the cells in 3–5 ml of ice-cold 1 PBS and transfer into a 15 ml centrifuge tube. If the cells are nonadherent, collect them along with the media in a 15 ml centrifuge tube. Centrifuge the cells at 800 g (1000 rpm) for 5 min in a tabletop centrifuge with a swing bucket rotor (ThermoForma, General purpose centrifuge) at 4C to collect the cells. Wash the cell pellet twice with ice-cold 1 PBS and centrifuge at 800 g (1000 rpm) for 5 min in a refrigerated tabletop centrifuge at 4C. Discard the supernatant and proceed to the next step or snap freeze the pellet and store at –80C.

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4. Resuspend the pellet in 1.5 ml ChIP lysis buffer containing 60 ml PMSF. To the tube add 40 ml of 100 mM PMSF, 100 ml of 25 protease inhibitor stock, 460 ml of lysis buffer, 100 ml of 20 % SDS, 80 ml of 5 M NaCl, and 220 ml of nuclease-free water to bring the total volume to 2.5 ml. Mix the cells using a pipetman to obtain a uniform homogenate. Transfer the lysate equally (800 ml) into 1.5 ml centrifuge tubes. 5. Sonicate the samples to lyse the cells and shear the DNA to 100–1000 bp fragments. The time and number of pulses required to shear DNA depend on the cell type, extent of cross-linking, and the instrument used for sonication. We have observed that for most adherent cells three to four cycles of sonication at power four for 30 s on a Fisher Sonic Dismembrator (Model 100), followed by incubation on ice for 30 s is sufficient to shear the chromatin. However, number of pulses needed to shear DNA depends on cell density and cell type. Some cells are resistant to sonication treatment; micrococcal nuclease treatment may improve chromatin shearing in such cell lines (9, 11, 35–37). Refer alternate protocol for micrococcal nuclease treatment. 6. Transfer the sonicated samples into 1.5 ml tubes and centrifuge at 17,000 g for 15 min in a refrigerated microcentrifuge (Beckman-Coulter, Microfuge R or an equivalent model) to remove the cell debris. The samples can be snap frozen and stored at –80C or proceed directly to the immunoprecipitation step. 7. Sonication efficiency can be checked by de-cross-linking 100 ml of the sample and loading in a 1% agarose gel.

3.2. Setting up Immunoprecipitation Reaction

1. Thaw the frozen ChIP lysates on ice. Transfer lysate from three 1.5 tubes into one 15 ml tube and add five volumes of IP dilution buffer containing freshly added protease inhibitors (Roche). 2. Use 10–15 mg of antibody per immunoprecipitation reaction. The amount of antibody to be added depends on quality, affinity, and specificity of the antibody (See Note 4). Usually a negative control is performed using the same number of cells with a nonspecific IgG or no antibody control (See Note 5). Preclearing ChIP lysate is recommended to reduce the nonspecific backgrounds and eliminate false positives. For preclearing protocols see support protocol Section 3.2.1. 3. After adding the appropriate amount of antibody, rotate the reaction overnight at 4C.

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4. The antibody bound protein–DNA complexes can be recovered using protein A (Amersham Biosciences), protein G (Amersham Biosciences), protein A/G (Santa Cruz Biotechnology), or staph A (ATCC) cells. The choice of the beads depends on the isotype of the antibody to be used for the immunoprecipitation and their affinity for protein A, protein G, or protein A/G. Protein G is used for most of the applications in our laboratory. Add the required volume of protein G beads containing 20% ethanol (GE Healthcare) to a microcentrifuge tube and centrifuge at 3000 rpm for 1 min. Discard the supernatant, add 1 ml of wash buffer to the tube, mix well, and centrifuge. Discard the supernatant and repeat the washings for three times with wash buffer to remove the ethanol preservative. Add 200 ml of 1:1 protein G slurry or 400 ml of protein A slurry to the IP reaction. 5. Add PMSF to the IP reaction to a final concentration of 1 mM. Incubate IP reaction with the beads at room temperature for 2–4 h. 6. Centrifuge the tubes at 800 g (3000 rpm) at 4C (BeckmanCoulter, Microfuge R or an equivalent model) for 1 min. Discard the supernatant by aspirating carefully to avoid losing beads. 7. Add 700 ml of wash buffer 1 containing freshly added PMSF, mix well, and transfer to a new microcentrifuge tube. Rotate the tube in a nutator at room temperature for 1 min. Centrifuge at 800 g for 1 min and discard the supernatant. 8. Repeat the steps 7 and 8. 9. Add 700 ml of wash buffer 2 containing PMSF, rotate in a nutator for 5 min, and centrifuge at 800 g for 1 min. 10. Discard the supernatant, add 700 ml of wash buffer 3, rotate the tubes in nutator for 5 min, and centrifuge at 800 g for 1 min. 11. Discard the supernatant; wash the beads with 700 ml of TE. 12. Rotate the tubes in nutator for 1 min. Centrifuge at 800 g for 1 min and discard the supernatant. 13. Repeat washing with TE, centrifuge at 800 g for 1 min, and discard the supernatant. 14. Add 200 ml of elution buffer to the beads and incubate at 65C for 30 min. 15. Centrifuge at 2000 g (5000 rpm) for 2 min. Transfer the supernatant into a new tube. This is the immunoprecipitated sample. 3.3. Reverse CrossLinking

1. Add 10 ml of proteinase K (20 mg/ml) to 200 ml of the sample. 2. Incubate overnight in heat block at 65C.

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3. Clean up the de-cross-linked samples using Affymetrix cDNA cleanup columns. Elute twice with 25 ml elution buffer. 3.4. Amplification of Immunoprecipitated DNA

A single chromatin immunoprecipitation sample does not provide enough DNA for hybridization into a promoter array (See Note. 6–9). Commonly used techniques for amplifying the immunoprecipitated DNA are random primed PCR amplification, LM-PCR (See alternate protocol Section 3.3.1), and WGA (See alternate protocol Section 3.3.2). Random primed PCR amplification of immunoprecipitated DNA 1. Use sequenase enzyme for linear amplification of immunoprecipitated DNA. Dilute sequenase stock with sequenase dilution buffer to 1.3 U/ml. Four microliters of the 1.3 U/ ml working stock will be needed for each sample being amplified. For initial round of linear amplification use 10 ml of immunoprecipitated DNA. Set up eight reactions (four with immunoprecipitated sample and four with the control IP) with the following components (volume shown is for one reaction) in 200 ml PCR tubes.

Purified DNA

10 ml

5 sequenase reaction buffer

4 ml

Primer A (200 mM)

4 ml

Total volume

18 ml

2. Cycle conditions for random priming 1. 95C for 4 min 2. Snap cool samples on ice 3. 10C on ice 3. Prepare the cocktail using the following components (volume shown is for one reaction)

1. BSA (20 mg/ml)

0.1 ml

2. DTT (0.1 M)

1 ml

3. dNTPs (25 mM)

0.5 ml

4. Diluted sequenase (1/10 from 13 U/ml stock)

1 ml

5. Total volume

2.6 ml

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3a. Prepare cocktail for eight reactions. 1. Add 2.6 ml of the cocktail to each sample; mix well by pipetting, give a brief spin to bring down the components, and keep the samples back in thermocycler. 2. 10C for 5 min. 3. Ramp from 10 to 37C over 9 min. 4. 37C for 8 min. 5. 95C for 4 min. 6. Snap cool on ice for 5 min. 7. 10C for 5 min. 8. Add 1 ml of diluted sequenase (1.3 U/ml) to each sample. 9. 10C for 5 min. 10. Ramp from 10 to 37C over 9 min. 11. 37C for 8 min. 12. Repeat from steps 7–13 for two more cycles. 13. 4C hold. 4. For each IP purify with Microspin S-300 HR (GE Healthcare) columns. 5. Set up PCR for incorporating dUTP into the immunoprecipitated DNA, Prepare a dNTP mixture containing dUTP at concentrations indicated below: dCTP-10 mM dATP-10 mM dGTP-10 mM dTTP-10 mM dUTP-10 mM This mixture containing dNTP + dUTP can be stored at –20C. Set up PCR using the following components: Random primed DNA from the previous step

20 ml

10 PCR buffer

10 ml

50 mM mgCl2

3 ml

10 mM dNTPs+dUTP

3.75 ml

100 mM primer B

4 ml

Taq polymerase (5 U/ml)

2 ml

Nuclease-free water Total volume

58.75 ml 100 ml (continued )

352

Pillai and Chellappan (continued) Cycle conditions A. 95C for 30 s 45C for 30 s 55C for 30 s 72C for 60 s B. 95C for 30 s 45C for 30 s 55C for 30 s 72C for 60 s, add 5 s for every subsequent cycles. 4C hold.

7. After PCR amplification, check amplified DNA on 1% gel. 8. Purify PCR samples with Affymetrix cDNA cleanup columns, elute with 20 ml elution buffer. 9. Quantitate DNA using spectrophotometer or NanoDrop. Ideally, more than 5 mg of amplified DNA is obtained from each IP reaction. To check the differences between the IP and the negative control samples, real-time PCR should be performed using primer sets designed for DNA regions that are known to be specifically immunoprecipitated using the antibody used in the experiment. 3.5. Fragmentation of Amplified DNA

Components for fragmentation are available in GeneChip WT double-stranded DNA terminal labeling Kit (P/N 900812). The conventional method of fragmentation of DNA molecules utilizes DNase 1 to digest the DNA molecules, which produces fragments with 30 –OH termini compatible for terminal labeling by TdT. However, a serious issue in using DNase 1 in fragmenting DNA is that it may result in fragments with less than desired lengths. Recently, uracyl DNA glycosilase (UDG) in combination with apurinic or apyrimidinic endonuclease 1 (APE 1) has been successfully employed to generate DNA fragments. UDG specifically recognizes uracil and removes it by hydrolyzing the N–Cl0 glycosylic bond linking the uracil base to the deoxyribose sugar. The

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loss of the uracil creates an abasic site (also known as an AP site or apurinic/apyrimidinic site) in the DNA (38, 39). Treatment of the sample DNA molecule with APE 1 will cause controlled breaks in the DNA at the abasic site. An apurinic/apyrimidinic endonuclease (APE1) can cleave the DNA molecule at the site of the dU residue yielding fragments possessing a 30 -OH termini, thus allowing for subsequent terminal labeling. The amount of dU incorporation may be regulated to determine the average length of fragments after UDG/APE 1 treatment. The ratio of dUTP to dTTP may be, for example, about 1–4, or about 1–5, 1–6, 1–10, or 1–20. The amount of dUTP incorporated depends on the ratio of dUTP to dTTP which in turn will decide the average size of fragments. The higher the ratio of dUTP to dTTP the more uracil incorporated and the shorter the average size of the fragments. 1. Set up the fragmentation reaction as follows (volume shown is for one reaction): Double-stranded DNA from the previous step

5.5–7.5 mg

10 cDNA fragmentation buffer

4.8 ml

UDG (10 U/ul)

1.5 ml

APE (100 U/ul)

2.25 ml

Nuclease-free water

up to 48 ml

Mix the components and give brief spin. 2. Incubate the reactions at 37C for 1 h, followed by 93C for 2 minutes and 4C for at least 3 min. Flick mix the tubes and spin down briefly and transfer 45 ml to a new tube. 3. The remaining sample can be used for fragmentation analysis using a Bioanalyzer using RNA 6000 NanoLabChip Kit which will reveal the degree and uniformity of the fragmented products. The fragmented samples can be stores at –20C until proceeding to the next step. 3.6. Labeling Fragmented DoubleStranded DNA

1. Set up the reaction by adding the following components [available in GeneChip WT double-stranded DNA terminal labeling Kit (P/N 900812)]. 5 TdT buffer

12 ml

TdT

2 ml

DNA-labeling reagent (5 mM)

1 ml

Total volume

15 ml

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Add 15 ml of the labeling mix to the fragmented DNA samples. Flick mix the tube and spin down. 2. Incubate the labeling reaction at 37C for 1 h followed by 70C for 10 min and 4C for 3 min. 3. Use 2 ml of each sample for gel-shift analysis. 3.7. Hybridization and Array Processing

1. Prepare the hybridization mixture as follows Labeled DNA (5–7.5 ug)

60 ml

Control oligonucleotide B2 (50 pM)

3.3 ml

2 hybridization mix

100 ml

DMSO

14 ml

Nuclease-free water

to make up the total volume to 200 ml

The components for making the hybridization mixture are available in GeneChip hybridization, wash, and stain kit (Affymetrix). For preparing buffers see support protocol Section 3.2.2 Mix the components of the tube and give a brief spin. 2. Heat the mixture at 99C for 5 min, cool to 45C for 5 min, and centrifuge at 17,000 g for 1 min. 3. Inject 200 ml of the labeled sample into the array (GeneChip human promoter 1.0 R array, Affymetrix) through one of the septa. 4. Keep the array in 45C hybridization oven, at 60 rpm, and incubate for 16 h. 5. After hybridization, remove the hybridization mix and store at –20C for future use. 6. For washing and staining Affymetrix arrays, the GeneChip Fluidics Station 450/250 or 400 is used and it is operated through GCOS. Fill the probe array completely with the appropriate volume of non-stringent wash buffer (6 SSPE, 0.01% Tween-20). 7. Select the experiment name from the drop-down experiment list in the fluidics station. 8. In the protocol drop-down list, select FS450_0001 or FS450_0002 to control the washing and staining of the array. 9. To begin the washing and staining, choose RUN from the dialog box. Follow the instructions in the LCD window on the fluidics station. 10. Insert the appropriate probe array into the designated module of the fluidics station while the cartridge lever is in the

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eject position. The lever returns to the engaged position when finished. 11. Place a microcentrifuge tube (1.5 ml) containing 600 ml of SAPE solution mix/stain cocktail 1 in sample holder 1, one vial containing 600 ml of antibody solution mix/stain cocktail 2 in sample holder 2, and another vial containing 800 ml of array holding buffer in sample holder 3. 12. Press down on the needle lever to snap needles into position and to start the run. 13. When the run is completed, remove the tubes containing the solutions, and replace with fresh microcentrifuge vials. 14. Remove the array from the fluidics station by pressing down the lever to eject position. 15. Scan the array using GeneChip Scanner 3000 7 G. Probe arrays can be stored for a short while at 4C, in the dark. 16. Turn the scanner on 10 min prior to use for warming up the laser. 3.8. Protocol Check Points

The protocol was successfully used for hybridization with Affymetrix human promoter 1.0 array in our lab. Protocol conditions should be optimized by each user depending on array type, cell type, specificity of antibodies, assay conditions, etc. The following steps are recommended as checkpoints to assess the success of the immunoprecipitation protocol prior to performing hybridizations. 1. Visualization of sheared chromatin in a 1% agarose gel. Ideally, 100–1000 bp fragments should be obtained after successful sonication. Alternatively a bioanalyzer can be used to determine the accurate size of the sheared DNA. Care should be taken not to overshear or oversonicate DNA as it may result in damaged DNA–protein complex and unsuccessful immunoprecipitation. The optimum sonication should be determined for each cell type. 2. Gene-specific PCR of a known target to check IP enrichment. Validate the antibody with RNA pol II to see whether the immunoprecipitation is working. 3. Monitor DNA yield after DNA cleanup and amplification of immunoprecipitated DNA preferably using NanoDrop. Unfortunately it does not provide information on the quality of amplified DNA.

3.9. Support Protocols

3.9.1. Preclearing of ChIP lysate

Preclearing of ChIP lysates is an efficient method to eliminate nonspecific background in ChIP assay (22, 40). 1. Add 120 ml of 1:1 protein G slurry (or 400 ml of protein A) to the ChIP lysate (mentioned in Section 1.1) in a microcentrifuge tube. 2. Add 15 ml of 10 mg/ml salmon sperm DNA (Sigma-Aldrich, cat. no. D7696) and 10 ml of BSA (10 mg/ml).

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3. Rotate on a nutator for 30 min at 4 0C. Spin the tubes at 17,000 g for 30 min in a tabletop centrifuge at 4C. 4. Collect the supernatant and discard the beads. This supernatant fraction is used for the immunoprecipitation reaction described in Section 3.1.2 3.10. Alternate Protocols

3.10.1. Amplifying Immunoprecipitated DNA by LigationMediated PCR

A. Blunting of DNA ends using T4 polymerase 1. Take approximately 200 ng of immunoprecipitated DNA in a microcentrifuge tube. Set up separate reactions for sample and control DNA. Make up the volume to 55 ml using nuclease-free water. 2. Make the following master mix in a 0.5 ml tube.

Final conc.

Stock

1 Mix

1

10 T4 DNA polymerase buffer

11.0 ml

5 mg

10 mg/ml BSA (NEB)

0.5 ml

40 nM

10 mM each dNTP

1.0 ml

1.5 U

3 U/ml T4 DNA polymerase (NEB)

0.5 ml

ddH2O

42.0 ml 55.0 ml

3. Add 55 ml of the above master mix to all samples. Flick mix the tubes and briefly spin down the contents in the tube. 4. Incubate for 20 min at 12C in thermal cycler. 5. Transfer reaction to pre-chilled 1.5 ml tubes and extract the DNA by adding 120 ml phenol:chloroform:isoamyl alcohol (Fisher Biotech, cat. no. BP1752-100). Vortex well to mix the layers. Spinthe tubes at 14,000 rpm for 15min at 4C toseparate the layers. Transfer aqueous layer to new centrifuge tube containing 250 ml ethanol. Incubate for 30 min at –80C. Pellet the DNA by centrifuging at 15,000 rpm for 20 min at 4C. 6. Wash the pellet with 70% ethanol and centrifuge again at 15,000 rpm for 20 min at 4C. Dry the pellet and resuspend DNA in 25 ml of water. 7. Alternatively, use a QiaQuick spin column (Qiagen corporation) to extract DNA and elute the DNA in 25 ml of TE. B. Blunt-end ligation 1. Prepare 15 mM linker stock by mixing the following: 250 ml Tris–HCl (1 M) pH 7.9

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375 ml oligo A (50 -GCGGTGACCCGGGAGATCTGAATTC) (40 mM stock) 375 ml oligo B (50 -GAATTCAGATC) (40 mM stock) Aliquot 50 ml into PCR tubes, place the tubes in a thermocycler, and run the following program 95C for 5 min 70C for 1 min Ramp down to 4C (0.4C/min) 4C hold Store the linkers at –20C. 2. Make master mix for ligation on ice (25 ml per reaction): Final conc.

Stock

1 Mix

1

5 ligase buffer

10.0 ml

2 mM

15 mM linkers

6.7 ul

200 U

400 U/ml T4 DNA ligase (NEB)

0.5 ml

ddH2O

7.8 ml 55.0 ml

3. Add 25 ml of ligation mix to 25 ml of sample. 4. Incubate 16 h in 16C water bath. 5. Add 6 ml of 3 M sodium acetate and 130 ml ethanol. Incubate 30 min at –80C. 6. Spin at 15,000 rpm for 20 min at 4C to pellet DNA. Wash pellets with 500 ml of 80% ethanol. 7. Dry pellets and resuspend each in 25 ml H2O. C. Ligation-mediated PCR 1. Transfer 25 ml of IP and control sample in PCR tubes. 2. Make two buffer mixes per reaction:

Buffer mix A 10 thermopol buffer (NEB)

4.00 ml

dNTP mix (2.5 mM each)

5.00 ml

oligo A (40 mM)

1.25 ml

ddH2O

4.75 ml

Total

15.00 ml

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Buffer mix B 10 thermopol buffer (NEB)

1.0 ml

Taq polymerase (5 U/ml)

0.5 ml

ddH2O

8.5 ml

Total

10.0 ml

3. Add 15 ml of mix A to each sample and run the following program

Step 1:55C

4 min

Step 2:72C

3 min

Step 3:95C

2 min

Step 4:95C

30 s

Step 5:60C

30 s

Step 6:72C

1 min

Step 7:GOTO Step 4 twenty-four times Step 8:72C

5 min

Step 9:4C

HOLD

4. Midway through Step 1, add 10 ml mix B to each tube to hot start reactions. If necessary, pause program in Step 1 so tubes remain at 55C while adding mix B. 5. After PCR is completed, precipitate the DNA by adding 25 ml of 7.5 M ammonium acetate and 225 ml of 100% ethanol. 6. Pool samples where appropriate. Add 250 ml precipitation mix per 50 ml of PCR reaction. Incubate 30 min at –80C. 7. Spin at 20,000 g for 10 min at 4C to pellet DNA. Wash pellets with 500 ml of 80% ethanol. 8. Dry pellets and resuspend each in 50 ml H2O. 9. Measure DNA concentration with NanoDrop (use 10fold dilutions, if necessary) and normalize all samples to 500 ng/ml.

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3.10.2. Whole Genome Amplification PCR for Amplifying Immunoprecipitated DNA

359

This protocol is adapted from the protocol provided with Sigma GenomePlex kit (Product No. WGA1-50RXN) (41). GenomePlex Whole Genome Amplification (WGA) Kit utilizes a proprietary technology based on random fragmentation of genomic DNA and conversion of the resulting small fragments to PCR-amplifiable library molecules flanked by universal priming sites. WGA is achieved by PCR amplification of the library molecules using universal oligonucleotide primers. A. Library Preparation 1. Add 2 ml of 1 library preparation buffer to 10 ml of immunoprecipitated DNA and transfer to PCR tubes (0.2 ml). Usually, the concentration of immunoprecipitated DNA is too low to get an accurate quantitation. The entire volume of reverse cross-linked, purified DNA (either by QiaQick or by Affymetrix c DNA clean up kit) is lyophilized and resuspended in 10 ml of DNA. If the concentration of DNA from a single IP is very low, a couple of IPs can be individually performed and the DNA can be subsequently pooled and concentrated as mentioned above. 2. Add 1 ml library stabilization solution, mix by pipetting, and give a brief spin. Place the tubes in a thermocycler at 95C for 2 min. 3. Immediately cool on ice. 4. Add 1 ml of library preparation enzyme, mix by pipetting, and quick spin. 5. Incubate in thermocycler as follows 16C for 20 min 24C for 20 min 37C for 20 min 75C for 5 min 4C hold 6. Proceed to first round of amplification or freeze at –20C. B. Amplification 1. Prepare master mix for amplification for each sample as follows: 10 amplification master mix Nuclease-free water WGA DNA polymerase

7.5 ml 47.5 ml 5 ml

2. Add 60 ml master mix to each sample. Mix well and give a brief spin

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3. Incubate in thermocycler block as follows: 95C for 3 min, then 14 cycles of 94C for 15 s 65C for 5 min, then 4C hold Amplified material can be stored at –20C indefinitely, if needed. 4. Purify samples using QiaQuick PCR clean up columns and elute the samples in water. (Subsequent labeling reactions are efficient if eluted in water). 5. Quantitate DNA by NanoDrop or spectrophotometer. Usually, the DNA amount will be 1–4 mg which is sufficient for downstream labeling reactions. If the yield of DNA is low, reamplify the sample using Sigma GenomePlex WGA Reamplification Kit. C. Reamplification Transfer 15 mg purified amplification product in 10 ml volume to 0.2 ml PCR tubes. Use this as the starting material for second round amplification. Prepare master mix for each sample following the same method as in first round amplification. Follow the same thermocycler conditions as described earlier. Purify the samples using QiaQuick PCR clean-up columns. 3.10.3. Fragmentation of Chromatin Using Micrococcal Nuclease

Micrococcal nuclease treatment can be employed to cleave chromatin into oligonucleosomes (9, 35). This method is known as native chromatin immunoprecipitation. However, this technique is incompatible with formaldehyde cross-linking since crosslinked chromatin is inefficiently cleaved by nucleases. 1. Prepare nuclear pellet from cells as described in Refs. (9, 35). 2. Resuspend the nuclear pellet in 1 ml MNase digestion buffer and place on ice. 3. Aliquot into 1.5 ml microcentrifuge tubes with 500 ml of resuspended nuclei. 4. Add 1 ml of MNase enzyme (Amersham Bioscience, cat. no. E70196Y) to each tube and mix gently. 5. Incubate the tubes at 37C in water bath for 10 min. The incubation times as well as MNase concentrations have to be optimized to generate mainly tri-, di-, and mono-nucleosomes. 6. Add 20 ml of stop solution. Chill on ice. 7. Centrifuge the suspension at 13,000 g (12,000 rpm) for 20 s at 4C in a refrigerated microcentrifuge. The supernatant is made up of the second soluble fraction S2 containing larger fragments of chromatin, whereas, the pellet contained the smaller fragment S1 fraction. Resolubilize the pellet in lysis buffer and pool with the S2 fraction in a fresh tube. 8. Continue Chip assay as described in Section 3.1.1 from Step 6 onwards.

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3.10.4. Preparation of Magnetic Beads

361

The use of magnetic beads for immunoprecipitation has been widely used in recent years (42, 43). The beads are available in different forms, precoupled with protein G, protein A, or secondary antibodies (Dynabeads protein G (Invitrogen, cat. no. 100–03D), Dynabeads protein A (cat. no. 100–01D). For each IP 100 ml of Dynabeads is recommended. 1. Transfer 100 ml of Dynabeads to 1.5 ml microfuge tubes. 2. Add 1 ml blocking buffer (0.5% BSA in 1 PBS). 3. Collect the beads using a magnetic stand. Remove the supernatant. 4. Repeat washing with blocking buffer for two more times. 5. Resuspend beads in 250 ml blocking buffer and add antibody (10 g) to the tube. 6. Incubate overnight in a nutator at 4C. 7. Next day, wash the beads three times with blocking buffer and resuspend the beads in 100 ml of blocking buffer. 8. For immunoprecipitation, add 100 ml magnetic bead–antiboby mix to the cell lysate from Step 3.2.1 and incubate over night in a nutator at 4C. For detailed description of ChIP using Dynabeads refer the protocols from Young lab (43).

3.11. Conclusions

ChIP-chip or chromatin immunoprecipitation followed by DNA microarray analysis has proven to be an efficient means of mapping protein–genome interactions on a genome-wide scale increasing our understanding of diverse cellular processes. Much of the work so far has focused on binding of transcription factors and many computational methods have been developed to identify the bound regions. Recently this technology has been extended to the genomic mappings of the other features such as histone modifications, transcriptionally active regions, and binding sites for other protein complexes. This article outlines the general strategies used to carry out ChIP-chip assays to study the differential recruitment of regulatory molecules based on the studies conducted in our lab as well as other published protocols.

4. Notes 1. GeneChip human promoter 1.0R array (Affymetrix, Cat. No. 900775) is available as a single array comprising of over 4.6 million probes tiled through over 25,500 human promoter regions. Sequences used in the design of this array are based on the sequences available from NCBI genome assembly (Build

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34). Oligonucleotide probes are synthesized in situ complementary to each corresponding sequence. Probes are tiled at an average resolution of 35 bp, as measured from the central position of adjacent 25 mer oligos, leaving a gap of approximately 10 bp between probes. Each promoter region covers approximately 7.5 kb upstream through 2.45 kb downstream of 50 transcription start sites. For more than 1300 cancer associated genes additional promoter region is included in the array, covering 10 kb upstream of transcription start site. 2. In addition to Affymetrix arrays, ChIP-chip microarrays are available from Agilent and NimbleGen. Available formats include whole genome tiling arrays for model organisms like human and mouse as well as focused arrays for promoter regions and custom array designs. Each Agilent ChIP-onChip microarray features a total of 244,000 60-mer oligonucleotide probes. Probes are spaced every 100–300 bp across regions of interest in both coding and non-coding DNA sequence. Such focused microarrays include proximal promoter (–0.8 KB upstream to +0.2 KB downstream of identified transcriptional start sites) and expanded promoter (–8.0 KB upstream to +2.0 KB downstream) designs. NimbleGen’s current high-density microarrays with long oligonucleotide probes (50–85 mer) contain 385,000 probes on a single glass slide. The entire non-repetitive human and mouse genomes can be surveyed at 100 bp intervals, each with a set of 38 arrays. Agilent and NimbleGen microarray platforms utilize two color labeling as opposed to single color labeling of Affymetrix arrays. Two color labeling using Cy3/ Cy5 can be done using CGH kit (Invitrogen) which utilizes a random primed, Klenow-based extension protocol. 3. In conventional ChIP protocol, cells or tissues are treated with formaldehyde, a cell permeable small molecule that can mediate protein–protein and protein–DNA cross-linking through Schiff base formation. To increase the degree of protein–protein cross-linking, treatment with dimethyl adipimidate can be done first, followed by formaldehyde treatment (44). This is an efficient method to observe indirect protein–DNA interactions. 4. Antibodies used for ChIP-on-chip can be an important limiting factor. Not all antibodies can effectively immunoprecipitate protein–DNA complexes. An antibody that gives a specific signal in Western blot need not be ideal for binding to a DNA bound, cross-linked protein. ChIP-on-chip requires highly specific antibodies that must recognize its epitope in free solution and also under fixed conditions. If the antibody is demonstrated to successfully immunoprecipitate cross-linked chromatin, it is termed ‘‘ChIP-Grade’’.

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Validated ChIP-grade antibodies are available from Abcam Inc., Santa Cruz Biotechnology Inc., and Upstate (Millipore Corporation). Since the affinity and avidity of antibodies can vary, the amount of antibody in each ChIP-chip reaction needs to be empirically determined. To demonstrate that the antibody was successful in immunoprecipitating the protein–DNA complex, PCR amplification on a known bindingsite region for the protein of interest should be performed using either conventional PCR methods followed by agarose gel electrophoresis or by quantitative PCR. The antibody used for immunoprecipitation should be a high affinity, high specificity antibody to avoid nonspecific background in ChIP assay. It is important to incorporate a positive control as well as an irrelevant antibody in the ChIP assay to facilitate critical interpretation of data. 5. It is necessary to have appropriate experimental controls in ChIP-chip experiments. Listed below are common controls that researchers are using in their experiments: (a) Nonspecific IgG antibodies – The most common type of negative control involves adding antibodies that do not recognize a specific epitope, for example, pre-immune serum or IgG. A potential pitfall is that since the antibodies do not immunoprecipitate effectively, the nonspecific DNA yield is often extremely low. Hence, the hybridization tends to be much noisier and can result in many false positives due to amplification of trace amounts of nonspecific DNA. Alternatively, the primary antibody can be omitted (i.e., no antibody control). (b) Protein deficient cell line – Several upstream applications, such as target deletion or siRNA, can be used to perturb the expression of the protein of interest, hence decreasing the amount of immunoprecipitated material. Alternatively, a cell line that does not express the protein of interest could be used as a negative control. (c) Tag-specific antibodies – If your protein of interest has a tag, such as GST or GFP, you may want to consider using antibodies against them in cells that do and do not express this tagged protein of interest. Generally genomic DNA is used as an input control and samples from no antibody or immunoglobulin G groups are used as negative controls. Other control designs such as transformed cell lines versus empty vector cell lines, wild type target versus mutation target, and drug treated versus untreated can all be used as control options (45). 6. As mentioned previously, the amount of starting material required for successful microarray hybridization is very critical in

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a ChIP-chip experiment. The amount is highly variable depending on the quality of the antibody, binding frequency of protein to DNA, and abundance of the protein. It is recommended that pooling of immunoprecipitated DNA samples to acquire enough DNA for subsequent amplification is a feasible strategy. Unfortunately, pooling ChIP samples is not always possible, especially if specialized cell types or tumor tissues are used. The amplification of immunoprecipitated DNA using T7 polymerase is widely used to linearly amplify ChIP DNA. However, a PCR bias may occur, especially in mammalian systems where large amounts of repeat sequence may skew data. 7. The statistical analysis of the huge amount of data generated from arrays is a challenge and normalization procedures should aim to minimize artifacts and determine what is really biologically significant. So far, application to mammalian genomes has been a major limitation, for example, due to a significant percentage of the genome that is occupied by repeats. However, as ChIP-on-chip technology advances, high resolution whole mammalian genome maps are achievable. 8. Although ChIP-on-chip can be a powerful technique in the area of genomics, it is very expensive. Most published studies using ChIP-on-chip repeat their experiments at least three times in order to obtain biologically meaningful maps. The cost of the DNA microarrays is often a limiting factor to whether a laboratory should proceed with a ChIP-on-chip experiment. 9. Another limitation is the size of DNA fragments that can be achieved. Most ChIP-on-chip protocols utilize sonication as a method of breaking up DNA into small pieces. However, sonication is limited to a minimal fragment size of 200 bp. In order for higher resolution maps, this limitation should be overcome to achieve smaller fragments, preferably to single nucleosome resolution.

Acknowledgments Studies in the author’s laboratory are supported by the grants CA63136, CA77301, and CA127725 from the NIH.

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Chapter 24 ChIP-on-Chip Analysis Methods for Affymetrix Tiling Arrays Sean J. Yoder and Steven A. Enkemann Abstract ChIP-Chip microarray data analysis is a multi-step approach that requires several different applications to progress from the initial stages of raw data analysis to the identification and characterization of ChIPbinding sites. There are multiple approaches to data analysis and several applications available for each stage of the analysis pipeline. Each application must be evaluated for its suitability for the particular experiment as well as the researcher’s computer background. This chapter is a review of the commonly available applications for Affymetrix ChIP-Chip data analysis, as well as the general workflow of a ChIPChip analysis approach. The purpose of the chapter is to allow the researcher to better select the appropriate applications and provide them with the direction necessary to proceed with a ChIP-Chip analysis. Key words: ChIP-chip, Affymetrix, tiling, data, analysis, normalization, algorithm, application, chromatin, binding.

1. Introduction Chromatin immunoprecipitation followed by microarray hybridization (ChIP-Chip) allows for a genome-wide survey of in vivo transcription factor to DNA-binding events, as well as the identification of histone and DNA modifications. With a range of a 7–35 base pair resolution, Affymetrix highdensity oligonucleotide tiling arrays can interrogate both coding and non-coding regions, generating an enormous amount of data. This presents multiple challenges for data normalization and analysis. These challenges are compounded by the fact that because this particular application of microarray analysis is in its infancy, there are few welltested applications for the analysis of ChIP-Chip data. Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_24 Springerprotocols.com

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Multiple studies have been published describing novel algorithms that intend to address the statistical problems associated with ChIP-Chip data, but many of these may not be appropriate for every researcher’s experiment or level of computer expertise. The analysis of ChIP-Chip data is a process that requires the identification of positive probes and ChIP-enriched binding regions, the mapping of those hits to the genes or the genome, and possibly the sequence analysis and binding motif discovery of those hits. There are various approaches to each stage of ChIPChip data analysis, and each application will employ one or more of these techniques to arrive at a conclusion. Each of these approaches will be explained here, allowing the reader to better understand the implementation of each application as it is reviewed. Each of the stages of ChIP-Chip data analysis requires a specific treatment of the data and often different applications to produce the output for the next stage. It should be stressed that a complete treatment of ChIP-Chip data will most likely include the use of at least one of each of the applications available for each stage of data analysis. Fortunately, there are standard file formats that may be exported at one stage and subsequently imported into one or more of the applications at the next stage. As long as the user is cognizant of the file types used for input and output, as well as the genomic annotation version that was employed when generating the output, ChIPChip data can be processed through the analysis pipeline in a straightforward manner. This chapter provides a survey of the algorithms and associated software applications that are currently available for Affymetrix ChIP-Chip analysis. Many algorithms were developed to address specific concerns or shortcomings of previous analysis solutions, but each will include its own set of statistical assumptions and biases. Therefore, the researcher will ultimately need to decide which approach is the most appropriate for their experiment. The intent of this chapter is to provide enough information to allow the researcher to make an informed decision about ChIP-Chip data analysis, as well as downstream visualization and annotation. It will also address the hardware and software requirements of each application as well as the level of computer expertise that is necessary to install and use the software effectively. Finally, it establishes a workflow of the ChIP-Chip data analysis process from raw data to binding site detection to visualization and biological characterization of the binding sites.

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2. Methods 2.1. Array Normalization and Probe Analysis

2.1.1. Affymetrix Tiling Analysis Software (TAS)

Affymetrix tiling arrays are designed using the typical perfect match (PM) and mismatch (MM) probe pair approach where the PM is a 25-mer oligonucleotide probe that is identical to the genomic sequence data at a specific locus, and the MM is a 25-mer that is identical to the PM except for the central nucleotide. The purpose of the MM probe is to estimate the contribution of the signal intensity that is derived from non-specific hybridization. Typically, the algorithms in Affymetrix analysis packages are designed to subtract the MM from the PM intensity, but many applications allow the user to ignore the MM data and estimate signal from the PM values alone. Currently, Affymetrix offers a variety of tiling arrays that includes a set that interrogates all the non-repetitive sequences of the human genome, a set that focuses on chromosomes 21 and 22, as well as an array that focuses only on the promoter regions of each gene. There are also arrays available for several other species that are commonly used in research. The analysis of any of these arrays may be carried out in the same manner. The first stage of ChIP-Chip data analysis ultimately results in the initial identification of ChIP-enriched segments of the genome. This stage of data analysis is crucial as all downstream binding site analysis is based on these results. Initially, the raw data files, Affymetrix .cel files, are imported into an application and normalized or globally scaled to a specific target intensity. Next, the application must estimate the background-corrected signal along each probe on the array. This may include the estimation of signal, p-value, or other statistical scoring mechanism which will be carried into the next step. Finally, the application must employ one or more statistical algorithms that will detect the regions of potential DNA binding by the protein of interest by evaluating the signal or statistical measurements produced in the previous stage. Each probe along the interrogated site is reported and compared to neighboring probes with the intent of identifying a statistically significant region of binding activity. Typically, these regions are reported as lists of genomic coordinates, but other metrics such as ChIP fold enrichment score and false discovery rate (FDR) may be reported by the software. The output files from this stage of analysis are commonly of the .bed file format which may be visualized in the genome browsers that will be discussed later. Next, several of the most commonly used and some recently reported algorithms and analysis applications will be discussed. An early Chip-ChIP study using Affymetrix arrays was performed in collaboration with Affymetrix and employed the approach used in the Affymetrix GTRANS, or the later named, Tiling Analysis

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Software (TAS) (1). TAS is a freely available standalone Windows application that is not tied into the Affymetrix GCOS system. Because it is flat file based, the user only needs to point to the array .cel files to initiate an analysis. The application requires several library files, including BPMAP and GRC files that are supplied by Affymetrix. After downloading the appropriate library files for the subject arrays and creating the tiling array group (TAG file) within the software, the TAS workflow starts with the .cel file normalization. The software can perform a global median scaling to specified target intensity or optionally perform quantile normalization of the probe intensity data (2). Quantile normalization assumes that the data have the same underlying distribution, so it is most likely inappropriate to normalize all control and enriched samples together. The software allows the user to indicate the preference of a onesample or two-sample analysis (control and enriched IP sample) or the corresponding quantile normalization treatment. It is important to note that although TAS will perform analysis on a onesample experiment, the array must be of the PM/MM type or the results from the analysis will not be accurate. After normalization and scaling, TAS probe analysis is performed which will generate reports of absolute signal intensity and p-values. The first step of probe analysis employs a sliding window approach where bandwidth is set to half the expected IP DNA fragment length, which is typically near 500 bp. A larger bandwidth may increase statistical power, but it may also increase background noise. Next, TAS performs a non-parametric Wilcoxon signed-rank test (also known as a Mann–Whitney U-test) (3) which is a non-parametric test for samples lacking a normal distribution. The test ranks all probe pairs in the window and confirms that the sum of the ranks in the ChIP-enriched sample is higher than that of the control sample. The p-values generated from this test may be reported in a –10log10format which allows convenient viewing along with sequence, interval, or intensity data in a genome browser. TAS probe analysis also results in a file with signal intensity estimates at each probe, as generated by the Hodges–Lehmann estimator (4), which are the median of the pairwise averages of the PM–MM scores in the sliding window. Additionally, in a two-sample analysis, fold enrichment is also reported as a log2-fold change between the treatment and the control group signals. The final stage of probe analysis is an interval analysis which uses the output files from the previous steps to report discrete regions of potential target protein binding along the sequence of interest. The interval analysis is dictated by three parameters including a p-value or signal intensity threshold (depending on which is used for input), a max_gap parameter which results in a higher level of stringency with a smaller value and min_run which


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is more permissive with a smaller threshold value. The final set of the detection regions, or intervals, is output in a .bed file format and can be used as an input for downstream analysis or data visualization. Affymetrix Tiling Analysis Software could be considered a good starting point for the analysis of ChIP-Chip tiling data since the software is free, easy to install, and relatively straightforward to use. However, there is a learning curve to the software for those starting into ChIP-Chip analysis, so it is recommended to thoroughly review the documentation as well as the Affymetrix web talks that specifically address TAS with ChIP-Chip data. There are some caveats of the TAS package that should be considered. First, the software does not allow one-sample analysis for the tiling arrays that are not designed with MM probes. This may present challenges for those who would like to perform a small experiment without a control sample or for those who are checking QC of one array at a time. In addition, and perhaps more importantly, TAS does not account for probe affinity which is the variability in probe behavior based on sequence and GC content as well as high copy number probes that are specific to multiple sites of the genome. Some applications that address these concerns will be examined below. 2.1.2. HMMTiling

In order to address some of the potential shortcomings of the Affymetrix Tiling Analysis Software, Li et al. (5) developed a ChIP-Chip algorithm based on a hidden Markov model (HMM) (6) called HMMTiling. The software is offered as source code and runs in a Python command line environment on a Linux operating system only and is available for download at http:// genome.dfci.harvard.edu/xsliu/HMMTiling/. The authors state that a compiled binary version with a graphical user environment (GUI) will be released for both Windows and Linux, but at the time of this writing these alternatives are not yet available. HMMTiling was designed with the approach of developing a repeat-masked probe model that is defined using multiple samples from several experimental data sets of the user’s choice. The result is a model that estimates the behavior of each individual probe with the intention of minimizing the effects of poorly performing probes. In addition, the authors assert that the use of a HMM addresses the notion that the Wilcoxon rank test used in TAS requires additional replicates and a larger window size (which will consequently include more noisy probes or miss short binding regions) to evaluate the ChIP-enriched regions with a sufficiently small p-value The strategy of the HMMTiling software starts with a quantile normalization of the .cel files. Next, repetitive probes, those that are represented multiple times on the array as well as those that map multiple times to the genome, are filtered out before further

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analysis. Essentially, probes must have only one measurement within 1 kb of the genome, otherwise latter occurrences of the sequence are removed. Probes must also occur only once for each genomic position. Baseline probe behavior is then estimated by evaluating each probe across every sample in the data set. An HMM is then applied to estimate the probability of ChIP enrichment at each probe location, taking all replicates into consideration. A summary enrichment score is reported for the discovered ChIPenriched regions, and the data may be exported as a .bed file for downstream analysis and data visualization. HMMTiling is a good alternative to Affymetrix TAS since the algorithm takes baseline probe performance into consideration, further filters out repetitive probes that will potentially confound the data, and it obviates the need for the fixed-size sliding window of the Wilcoxon rank test. Additionally, the user has the option to pool together data from multiple independent data sets to estimate probe behavior. The authors claim that reasonable results may be attained from a single ChIP experiment so this application may be very useful to those who are testing multiple ChIP conditions and would rather not make use of multiple arrays during the optimization stages of their experiment. However, since this application is only available as a command line application for Linux operating systems with limited documentation, it may be out of reach for the majority of biologists at this time. Perhaps because of this, at this time, other applications described below may be a good alternative to researchers. 2.1.3. MAT

An algorithm titled model-based analysis of tiling-arrays (MAT) (7) was developed by the same group as the previously described HMMTiling application. This particular application is endorsed by Affymetrix as another alternative to their TAS application. MAT is open source and is available for download for Linux operating systems at http://chip.dfci.Harvard.edu/wli/MAT. The approach of MAT has some similarities to HMMTiling, but probe behavior modeling and ChIP-enriched region detection are performed very differently. The authors first remapped all of the 25-mer probe sequences in the BPMAP file of each tiling array type to the latest genome assembly while filtering out probes that map to more than one location in any 1 kb window, as well as those that map to the same genomic location. Probe behavior is estimated by assessing signal intensity, sequence, and copy number of all probes on each array, unlike HMMTiling where it requires multiple arrays to build the probe affinity model. Next, a probe affinity standardization method is employed to background correct and, in effect, normalize the arrays so that they may be directly compared. The authors report that quantile normalization is not necessary, and offers no further benefits after probe affinity standardization is performed.


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Once the arrays have been examined for probe affinity, ChIP region detection may be performed using a single ChIP-enriched sample only, multiple ChIP sample replicates, or multiple ChIP samples with input controls including those with unbalanced designs (i.e., one control to multiple ChIP-enriched samples). MAT uses a sliding window approach to identify enriched regions. The size of the window is user-adjustable and dependent on ChIP DNA fragment size. A trimmed mean of MAT-calculated t-values (where the upper and lower 10% of t-values from the window are removed) and a MATscore are calculated and reported. Computed t-values are standardized probe intensities that have been adjusted for the baseline predicted by the probe behavior model (sequence, copy number, etc.) and standard deviation of a pre-determined probe affinity bin. ChIP-enriched regions are then identified based on p-values, false discovery rate (FDR), or MATscores, depending on the user preference. Resulting MATscores for each probe and the genomic coordinates for enriched binding regions are exported as .bar files and .bed files, respectively, for downstream analysis and visualization. MAT is a powerful alternative to TAS that is reported to work sufficiently with as little as one ChIP-Chip sample, with no replicates or controls, similar to HMMTiling. Currently, the application is only available for Linux platforms running Python and it has a command line interface. This limitation will exclude many biologists, but those who have some experience with Linux may find the software tractable. The program expects a user-created text file called a .tag file which describes the experiment and a few library files that are downloadable from the MAT website. Launching MAT then requires a single line of code. 2.1.4. TileMap

TileMap is a free application developed by Ji and Wong (8) that also steps away from the sliding window approach of Affymetrix TAS. TileMap features user-selectable analysis options including a moving average (MA) or an HMM, a pseudo false discovery rate (FDR) calculation, and repeated probe filtering. TileMap is available in a command line interface for Windows and Unix operating systems at http://biogibbs.stanford.edu/jihk/TileMap/ index.htm, but recently the software was released as a graphical user interface along with the authors’ CisGenome project, available at http://www.biostat.jhsph.edu/hji/cisgenome/. TileMap employs a hierarchical empirical Bayes model to pool information from all probes on the array, followed by the generation of test statistics that are used to identify protein-binding patterns. The Bayes method allows one to estimate the probabilities of an individual probe by using information from empirical measurements of that probe and on the entire population (9). Neighboring probes are then combined using either a moving average (MA) or an HMM. The application is intended to

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accurately handle experiments with few replicates or complex multiple sample comparisons, and the authors claim that the algorithm runs >10 times faster than that of TAS. The implementation of TileMap involves a workflow that includes functions that normalize the raw data followed by log transformation of the PM or PM–MM values. In order to reduce noise, the application includes a repeat masking step that filters out any probe that occurs more than once in a 2 kb window. The algorithm then calculates an unbalanced mixture subtraction (UMS) which is similar to a local false discovery rate (FDR) for the MA or HMM model parameters. The output of TileMAP includes summaries for each probe with a .bed file containing the identified genomic binding regions. As usual, this file may be further processed by downstream applications. TileMap is navigated via command line interface for Windows and Unix and requires the user to set up two parameter files, but the documentation and examples may be adequate for many biologists to get started with analysis. However, an attractive alternative may be the CisGenome application since the features of TileMap have been incorporated into the interface along with a graphical user environment. 2.1.5. TiMAT

The TiMAT application essentially serves to address the issues of noisy probe data, calculate FDR, and identify and evaluate ChIPenriched intervals. TiMAT, freely available at http:// bdtnp.lbl.gov/TiMAT/, is an open source Java-based command line application that was specifically designed for ChIP-Chip data. It will run on Windows, Mac OS X, and Linux operating systems with the Java Runtime Environment (JRE) installed. TiMAT consists of a series of small programs that perform specific functions in the processing pipeline. The process involves multiple steps starting with the creation of a Tpmap file from an Affymetrix BPMAP file (this file must only be created once per array type). Next the .cel files are loaded and converted to a TiMAT-specific format, followed by normalization and scaling where replicates are averaged, log intensities are calculated, and subtraction of control values from ChIP-enriched values is performed. Next, window scores and optionally FDR are calculated using a sliding trimmed mean window. Then, enriched intervals are identified by employing a number of user-selectable filters including FDR, minimum interval length, and maximum gap. Finally, significant interval peaks are selected and the multiple reports are generated and exported from the application, however not of the typical .bed file format. The files will most likely have to be modified for downstream visualization in genome browsers. TiMAT uses a sliding window approach similar to that of Affymetrix TAS, but the content of the data files that are exported is much more robust in terms of scoring of intervals, FDR filtering,


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and peak finding. However, the software does require control samples and does not take into account the individual variation of each probe (probe affinity), nor does it address the repetitive probes on the array. In addition, the application must be installed and configured as a Java command line interface. The user will need to be comfortable working in DOS and addressing memory allocation by using the Java –Xmx flag or memory errors will likely occur. Additionally, the fact that each process of the pipeline requires the user to launch a different program, specifying multiple input files, may pose as a significant burden to the user. For these reasons, many biologists may not easily traverse the complete analysis pipeline using TiMAT and may be advised to work with one of the applications that include a graphical interface (TAS or CisGenome). 2.1.6. Tilescope

Tilescope is a graphical, web-based application developed by Zhang et al. (10) which is intended to be an integrated pipeline featuring an automated workflow. The application is freely downloadable at http://tilescope.gersteinlab.org and will run on any platform with a Java-enabled web browser. The processing pipeline for Tilescope involves three stages: data input, tile scoring, and feature identification. First, Affymetrix .cel and BPMAP files are uploaded through the use of a web form, and quantile normalization is then performed by default. Tile scoring is performed using a TAS-like sliding window approach with an added non-parametric statistical test to further assess the differences between the control and the ChIP-enriched samples at each probe. Both fold enrichment and p-value scores at each genomic location are reported. Binding sites are identified by one of three methods: the max-gap and min-run TAS-like metric, a novel iterative peak method, or an HMM method. Upon completion of data processing, a web page is displayed that includes a summarization of parameters, scatter plots, and histograms. Downloadable feature lists include genomic coordinates (including those for updated genome builds) along with the calculated metrics, as well as links to custom tracks on the UCSC genome browser. Because Tilescope is integrated and web based, it may be one of the easiest-to-use applications available to researchers who are just getting involved in ChIP-Chip analysis. It offers three choices for binding site identification as well as a seamless transition to data visualization in a genome browser. Although probe affinity and repetitive probes are not addressed here, the application may still offer a solid starting point for non-programmers who would like an alternative to TAS.

2.1.7. CisGenome

CisGenome is a publicly available integrated application that was developed by the authors of TileMap (8). The application is available with a graphical user interface (GUI) for Windows operating systems, as well as a command line interface for Linux and

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Mac OS. Both source code and binary executables for CisGenome are available at http://www.biostat.jhsph.edu/hji/cisgenome/index.htm. The CisGenome GUI enables a quick and easy approach to data analysis which starts with the loading of the appropriate BPMAP files and .cel files of interest. Next, the .cel files may be quantile normalized or analyzed using the MAT algorithm (7), and then binding site peak detection is performed using the TileMap algorithm. Once the analysis is complete, CisGenome can visualize the binding regions in the resulting output file from within the application. Tracks of interval data or gene annotations may be added to the genome browser and the user may traverse through each chromosome narrowing in on sites of interest. At the time of this writing, CisGenome was newly released and is currently in beta stages, but this application may be the most integrated, easy-to-use freely available package. The fact that it installs and runs as a graphical Windows application including a genome browser within the interface makes CisGenome an excellent application for the average biologist to use. In addition, the inclusion of the MAT and TileMap algorithms along with user-selectable parameters gives users the opportunity to easily step away from the Affymetrix TAS software. One caveat is that at this time the documentation for the graphical version of the software is limited to a basic tutorial, but the authors do offer a comprehensive manual for the command line version of the software which describes the implementation of the algorithms in detail. 2.1.8. Partek Genomics Suite

For researchers who are processing larger volumes of ChIP-Chip as well as other types of microarray samples, or for those who need the support and ease-of-use of a commercial analysis application, Partek Genomics Suite (Partek GS) may serve as an alternative to the freely available applications. Information as well as a free trial of the software for Windows, Mac, and Linux operating systems may be found at http://www.partek.com. Partek GS is designed to handle Affymetrix data for studies including gene expression, exon tiling, copy number analysis, SNP association, as well as tiling data. The application workflow starts with .cel file import and carries through to normalization, binding region detection, and visualization including mapping of binding regions to probe sets that are represented on the HGU133 Plus 2.0 human expression array. In addition, the software automatically provides links to external databases including NetAffx, UCSC Genome Browser, and the Affymetrix Integrated Genome Browser. Normalization and signal intensity estimation of the .cel files are achieved using the Robust Multiarray Average (RMA) algorithm (2) without a median polishing step, including quantile


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normalization and log2 conversion. Detection of potential ChIPbinding sites is then performed using a user-adjustable sliding window approach similar to that used in Affymetrix TAS. The benefits of Partek GS include its straightforward workflow and easily navigable interface, as well as its ability to handle multiple types of Affymetrix data, including the ability to merge data sets from these different array types within the same workflow. However, it should be noted that the software does not implement a probe affinity model or repetitive probe masking such as those included with MAT. If a commercial application that includes these features is preferred, Biotique Systems’ XRAY may be of interest. 2.1.9. Biotique Systems XRAY for ChIP-Chip

XRAY is an Excel add-in available commercially for Windows operating systems at http://www.orderxray.com. The graphical interface offers a wizard-driven workflow for the identification and scoring of binding sites. XRAY is targeted toward researchers on a budget who are comfortable with the Excel working environment. According to the Biotique Systems, XRAY is priced similar to the cost of a ‘‘couple of arrays.’’ XRAY may be a good choice for those who have limited funds and do not have the need for an application that supports multiple types of Affymetrix microarrays. XRAY includes built-in .cel file quality and normalization metrics, and users may select from the TAS (1), HMM (6), or MAT (7) algorithms for array processing and binding site identification. The resulting output may be viewed on the UCSC genome browser from within the application without the need to link externally to the site. Additionally, the software will export a document outlining the analysis methods employed in the array processing, and users may export the Excel charts to publications.

2.1.10. Genomatix ChipMiner

ChipMiner is a newly available application from Genomatix (http://www.genomatix.de) that is designed to handle any of the Affymetrix ChIP tiling arrays including promoter and whole genome sets. The graphical user interface of ChipMiner allows the user to import Affymetrix .cel files for enriched region detection analysis. The results are visualized against the overlapping genomic annotation including upstream and downstream loci. In addition to finding transcription factor binding sites, the software can identify phylogenetically conserved transcription factor binding motifs including a mechanism to compare these motifs to 10 vertebrate species. The final stage of the ChipMiner pipeline involves the display of all results as a pathway mining component.

2.2. Binding Site Visualization

Accurate signal estimation and binding site identification are central to ChIP-Chip analysis, but to garner meaningful results from the data it is necessary to further visualize and report the data

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using other downstream tools. Genome browsers are powerful interfaces that allow the simultaneous visualization of ChIP-Chip experimental data as a track against an annotated genome. Users may add additional annotation tracks containing information such as SNP sites, known genes, predicted genes, ESTs, or even data merged from gene expression data sets. The UCSC Genome Browser is a commonly used web-based genome browser that is available at http://genome.ucsc.edu/ index.html (11). The Genome Browser includes annotation tracks that are computed at UCSC from publicly available data, as well as tracks that are provided by collaborators. The user can view the data along the chromosome by zooming in or providing genomic coordinates from genome BLAST searches, etc. The browser includes clickable areas that allow the user to explore external databases from the NCBI and others such as the BLAT sequence alignment tool which may be used for further downstream analysis. Affymetrix also offers a freely available genome browser entitled Integrated Genome Browser (IGB) located at http:// www.affymetrix.com/partners_programs/programs/developer/ tools/download_igb.affx. Similar to the UCSC browser, IGB allows users to dynamically explore the genomic annotations by adding multiple tracks of user-generated data along with annotations from public databases. IGB is launched from within a browser as a Java application. Although similar in function to the UCSC browser, IGB has a completely different style of interface that allows rapid zooming and sliding across the chromosomal data. IGB is connected to multiple public databases, so the user may choose from multiple annotation tracks and genomes. In addition, IGB will open Affymetrix .chp files as generated from Tiling Analysis Software for a relatively seamless progression through the analysis pipeline. The software will also directly link to the NCBI Entrez nucleotide database and the UCSC Genome Browser. 2.3. Binding Site Analysis and Characterization

In addition to data exploration in genome browsers, it is extremely helpful to identify the genes that may be associated with the potential DNA-binding regions as well as to additionally characterize these sites. The cis-regulatory element annotation system (CEAS) (12) is an integrated web application that retrieves repeatmasked genomic sequences, calculates GC content, maps nearby genes, reports evolutionary conservation at the site of interest, and identifies transcription factor binding motifs from ChIP-Chip data. CEAS is available at http://ceas.cbi.pku.edu.cn. CEAS accepts input files with ChIP-binding regions in either the UCSC .bed format or the Sanger GFF format. Affymetrix BPMAP files are currently offered only for v35 of the NCBI genome assembly, but using the LiftOver program at the UCSC Genome Browser site (http://genome.ucsc.edu/cgi-bin/


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hgLiftOver), a user can convert .bed files exported from TAS or IGB to the updated v36 assembly needed to load into CEAS. Once the files are uploaded, the application will create conservation plots from a phylogenetic HMM which are then generated as phastCons thumbnails and exported to a .pdf file for review. In addition, CEAS will also search for the nearest RefSeq genes on both strands within 300 bp from the binding site of interest. This allows the researcher to potentially identify which particular genes may be affected by their protein of interest. CEAS reports the relative binding location within the gene (intron, exon, UTR, etc.), the length and GC content of the binding site, and it provides a link to the UCSC genome browser. In addition to the basic binding site characteristics, CEAS finds the enriched sequence motifs from within the binding sites using the precomputed motif-scanning methods TRANSFAC and JASPAR. Fold change, p-value, and other metrics are reported with each motif along with a helpful graphical sequence logo (covered below) which gives a quick visual indication of sequence conservation at each base in the binding motif. Additionally, CEAS can retrieve RepeatMasked genomic sequences at each binding site which allows the researcher to design qPCR primers or perform additional motif searching, which will be covered in more detail. Genome browsers typically allow the user to export the genomic sequence data associated with the potential binding sites. The next step of the ChIP-Chip analysis pipeline may include the processing of this sequence data in order to identify and report these potential regulatory elements as conserved motifs. Several motif discovery tools exist which allow the user to verify or identify de novo binding sites from the exported sequence data. Some common de novo search applications include MEME, AlignAce, and GLAM among many others which were previously reviewed and evaluated (13). Using a comprehensive scan of the sequence data exported from a genome browser or application such as CEAS, the user may quickly report a list of potential binding motifs from the ChIP-Chip data. Most of these applications report multiple potential motifs along with scoring metrics to allow the user to evaluate the search results. Once the potential binding motifs are identified, the biologist may then report them as a graphical sequence logo by utilizing the WebLogo application (14).WebLogo starts with an aligned set of potential binding site sequences and creates a graphic that represents each nucleotide by a color-coded stack of letters corresponding to each of the four bases. The height of each letter indicates the proportion of the sequence conservation of that site across the series of aligned DNA sequences. WebLogo can then export the graphic to one of many formats including GIF and PNG, as well as print-friendly vector-based formats such as EPS and PDF. WebLogo is a freely available web application that may be accessed at http://weblogo.berkeley.edu.

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3. Conclusions ChIP-Chip analysis is a microarray technique that is nascent and still evolving. Consequently, there are relatively few applications available for the analysis of ChIP-Chip data, and a standardized workflow is just emerging. Many applications and approaches to ChIP-Chip data analysis have been developed in recent years. The methods covered here are among the most well known and well supported in the scientific community. As work progresses with ChIP-Chip data, the most fruitful techniques will undoubtedly be improved by data management, simplification of the workflow, and by the addition of graphical user interfaces that allows use by biologists with limited programming experience. In its current state, ChIP-Chip data analysis may initially seem daunting, but if the user starts with an application that is appropriate for his or her level of computer expertise, budget, and particular experiment, working with ChIP-Chip data will be much easier and will result in more accurate data.

References 1. Cawley, S., Bekiranov, H., Ng, P., Kapranov, E., Sekinger, D., Kampa, A., Piccolboni, V., Sementchenko, J., Cheng, A. Williams. (2004) Unbiased Mapping of Transcription Factor Binding Sites along Human Chromosomes 21 and 22 Points to Widespread Regulation of Noncoding RNAs. Cell. 116, 499–509 S. 2. Bolstad B.M., Irizarry R.A., Astrand M., Speed T.P. (2003) A comparison of normalization methods for high density oligonucleotide array data based on variance and bias. Bioinformatics. 19,185–93. 3. Troyanskaya O.G., Garber M.E., Brown P.O., Botstein D., Altman R.B.. (2002) Nonparametric methods for identifying differentially expressed genes in microarray data. Bioinformatics. 18, 1454–61. 4. Hollander, M., Wolfe, D.A. (1999) Nonparametric Statistical Methods, 2nd edition. John Wiley and Sons, Inc., New York, NY. 5. Li, W., Meyer, C.A., Liu, X.S. (2005) A hidden Markov model for analyzing ChIP-chip experiments on genome tiling arrays and its application to p53 binding sequences. Bioinformatics. 21, 274–82. 6. Rabiner, L. (1989) A tutorial on hidden Markov models and selected applications in

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speech recognition. Proc IEEE. 77, 257–286. Johnson, W.E., Li, W., Meyer, C.A., Gottardo, R., Carroll, J.S., Brown, M., Liu, X.S. (2006) Model-based analysis of tiling-arrays for ChIPchip. Proc Natl Acad Sci U S A. 103,12457–62. Ji, H., Wong, W.H. (2005) TileMap: create chromosomal map of tiling array hybridizations. Bioinformatics. 21, 3629–36. Casella, G. (1985) An Introduction to Empirical Bayes Data Analysis. Am Stat. 39, 83–87 Zhang, Z.D., Rozowsky, J., Lam, H.Y., Du, J., Snyder, M., Gerstein, M. (2007) Tilescope: online analysis pipeline for highdensity tiling microarray data. Genome Biol. 8, R81. Kent, W.J., Sugnet, C. W., Furey, T. S., Roskin, K.M., Pringle, T. H., Zahler, A. M., Haussler, D. (2002) The Human Genome Browser at UCSC. Genome Res. 12, 996–1006. Ji, X., Li, W., Song, J., Wei, L., Liu, X.S. (2006) CEAS: cis-regulatory element annotation system. Nucleic Acids Res. 34, 551–554.


13. Tompa, M., Li, N., Bailey, T.L., Church, G.M., De Moor, B., Eskin, E., Favorov, A.V., Frith, M.C., Fu, Y., Kent, W.J., Makeev, V.J., Mironov, A.A., Noble, W.S., Pavesi, G., Pesole, G., Regnier, M., Simonis, N., Sinha, S., Thijs, G., van Helden, J., Vandenbogaert, M., Weng, Z., Workman, C.,

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Ye, C., Zhu, Z. (2005) Assessing Computational Tools for the Discovery of Transcription Factor Binding Sites. Nat Biotechnol. 23, 137–144. 14. Crooks, G.E., Hon, G., Chandonia, J.M., Brenner, S.E.. (2004) WebLogo: a sequence logo generator. Genome Res. 14, 1188–90.

Chapter 25 Non-Radioactive Assay Methods for the Assessment of Telomerase Activity and Telomere Length Partha P. Banerjee and Shankar Jagadeesh Abstract The telomeric repeat amplification protocol (TRAP) assay is a highly sensitive PCR based assay and is an important tool for understanding the role of telomerase in cancer. This assay measures an enzymatic activity where the amount of target is dependent upon the activity of the enzyme. This protocol consists of two steps: first, telomeric repeats are added to the substrate by the enzyme and second, the extended products will be amplified by Taq-DNA polymerase. The amplified TRAP assay products will be separated on 10% native PAGE and detected by SYBR Green I dye. Key words: Telomerase, TRAP assay, Telomere, SYBR Green I, Southern blot.

1. Introduction Telomerase, a ribonucleoprotein enzyme with specialized reverse transcriptase activity, catalyses the synthesis and extension of telomeric DNA (1). Telomerase is present in germ line cells, cancerderived cell lines, and spontaneously immortalized cells in culture. It is activated in 85–90% of malignant tumors but usually absent in normal somatic cells which results in the progressive loss of telomeres with each cell division (2). Activation of telomerase allows cells to overcome replicative senescence, therefore, be a rate-limiting step in cellular immortality and oncogenesis (3). The telomerase complex is composed of telomerase reverse transcriptase (TERT) (4), telomerase RNA component (TERC) (5), telomerase associated proteins (TEP1) (6), and chaperone proteins (p23, Hsp90) (7). TERC, TEP1, p23, and Hsp90 are expressed in a wide variety of cells, irrespective of the presence or absence of telomerase activity. On Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_25 Springerprotocols.com

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the other hand, a strong correlation is observed between hTERT mRNA expression and telomerase activity in a variety of epithelial cancers (8–11), indicating that measuring telomerase activity could be an important tool for the detection of tumorigenesis. Similarly, telomeres play a critical role in chromosomes structure and function. They prevent aberrant recombination (12) and attachment of chromosomes ends to the nuclear envelope by capping the ends of chromosomes (12). In the somatic cells, telomere loss occurs with each replication cycle due to incomplete DNA replication in the absence of telomerase (13). Critically shortened length leads to chromosomal abnormalities, which ultimately resulting in replicative senescence of cells. In contrast, in immortal cell lines telomere length will be stabilized by the presence of telomerase (14). Thus, stabilization of telomere length appears to be one of the earliest and most prevalent genetic alterations acquired in the multi-step process of malignant transformation. Therefore, telomere length assessment could be a useful biomarker for monitoring early diagnosis of cancer. Here, we will describe telomerase activity (TRAP) assay first and then telomere length assay. 1.1. The TRAP Assay

This assay has two steps (as shown in Fig. 25.1 below). In the first step, telomerase in cell or tissue extracts extends the TS oligonucleotide with three or more TTAGGG repeats (lower case). These

Fig. 25.1. A flow chart depicting the various steps in the TRAP assay.

Non-Radioactive Assay Methods

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telomerase products are specifically amplified by PCR with the upstream TS2 and the downstream primer ACX (15). The ACX primer has 6 bp ‘anchor’ at the 50 -end which is neither telomeric nor complementary to telomeric sequences, followed by sequences that hybridize to telomeric repeats. The presence of the 6 bp anchor and non-complementary nucleotides in the primer reduces primer dimer PCR artifact formation. The anchored primer also prevents 30 elongation of telomerase products by capping the 30 -end of telomerase product after the first PCR cycle. Thus the length of PCR products produced in a TRAP assay utilizing the ACX primer accurately reflects the activity of the telomerase being tested (Fig. 25.2). The TRAP internal control utilized in the assay, TSNT, is amplified by TS primer and its own dedicated return primer, NT, which is not a substrate for telomerase. Thus, the internal control is semi-competitive in that it shares only one of the primers, TS, used to amplify the telomerase products. False-negative results can be easily identified by the

Fig. 25.2. Determination of telomerase activity in prostate cancer cells (LNCaP) with or without treatments of a telomerase inhibitor for 3 days in culture. Telomerase was extracted from cell pellets and subjected to TRAP assay. NC, negative control using lysis buffer only.

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Fig. 25.3. Determination of telomere length in prostate cancer cells (LNCaP) with or without the treatment of a telomerase inhibitor for 3 days. N, normal (untreated) LNCaP cells; mean telomere length is 2.5 kb; TI, LNCaP cells treated with telomerase inhibitor; mean telomere length is 1.5 kb. MWM, molecular weight marker.

disappearance of the internal control band with the incorporation of internal control. This non-radioactive TRAP assay can be used to quantify the level of telomerase activity in normal and cancer cells as well as in tissues. 1.2. The Telomere Length Assay

This assay consists of multiple steps. The first step is the isolation of genomic DNA. The genomic DNA is then digested and the DNA fragments are separated by gel electrophoresis. Following Southern transfer of the DNA fragments to a nylon membrane. The blotter DNA fragment will be hybridized with digoxigenin


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(DIG-labeled probe), a chemiluminescent probe specific for telomeric repeats and incubated with a DIG-specific antibody coupled with alkaline phosphatase. The immobilized telomere probe is visualized by adding alkaline phosphatase metabolizing substrate, CDP-Star. Lastly, the mean telomere repeat fragment is determined by measuring the chemiluminescence signal using an imaging system or by visually comparing the mean size of the smear to the molecular weight marker (Fig. 25.3).

2. Materials 2.1. TRAP Assay

1. Phosphate-buffered saline (PBS): 10 g NaCl, 0.25 g KCl, 1.43 g Na2HPO4, and 0.25 g KH2PO4. Make up to 1000 ml w/dH2O

2.1.1. Buffers and Solutions

2. Wash buffer: 10 mM HEPES–KOH (pH 7.5), 1.5 mM MgCl2, 10 mM KCl, and 1 mM DTT. 3. Cell/tissue lysis buffer: 10 mM Tris–HCl (pH 7.5), 1 mM MgCl2, 1 mM EGTA, 0.1 mM PMSF, 5 mM 2-mercaptoethanol, 0.5% CHAPS, and 10% glycerol. 4. 5 TRAP buffer: 100 mM Tris–HCl pH 8.3, 7.5 mM MgCl2, 315 mM KCl, 0.025% Tween-20, 5 mM EGTA, and 0.5 mg/ml BSA. 5. 5 TBE buffer: 54 g of Tris base, 27.5 g of boric acid, and 20 ml of 0.5 M EDTA (pH 8.0). Make up to 1000 ml w/ dH2O

2.1.2. Oligonucleotides

1. TS primer:

50 – AATCCGTCGAGCAGAGTT – 30

2. ACX primer:

50 – GCGCGGCTTACCCTTACCCT TACCCTAACC – 30

3. NT primer:

50 – ATCGCTTCTCGGCCTTTT – 30

50 – AATCCGTCGAGCAGAGT TAAAAGGCCGAGAAGCGAT – 30 (primers were synthesized from Integrated DNA Technologies, Iowa) 4. TSNT primer:

2.1.3. 10% PAGE for Telomere Analysis

1. Running gel: 5 TBE buffer, 5.0 ml; 30% acrylamide, 16.6 ml; TEMED, 0.035 ml; 10% APS, 0.1875 ml; and H2O, 50 ml. 2. Stacking gel: Tris–HCl (pH 6.8), 0.698 ml; 30% acrylamide, 0.752 ml; TEMED, 0.0067 ml; 10% APS, 0.0336 ml; and H2O, 6 ml.

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2.1.4. Additional Reagents

1. SYBR Green I dye (Molecular Probes, Inc.) 2. Taq-DNA polymerase (New England Biolabs) 3. dNTP mix (Invitrogen)

2.2. The Telomere Length Assay

2.2.1. Buffers and Solutions

1. Phosphate-buffered saline (PBS): 10 g NaCl, 0.25 g KCl, 1.43 g Na2HPO4, and 0.25 g KH2PO4. Make up to 1000 ml w/dH2O 2. DNA buffer: 1 M Tris–HCl, pH 8.0, 5 ml; 0.5 M EDTA, 5 ml; distilled water 15 ml 3. TAE buffer: 0.04 M Tris–acetate, 0.001 M EDTA, pH 8.0 4. Sodium dodecyl sulfate (SDS), 10% 5. Phenol:chloroform:isoamylalcohol 25:24:1 saturated with 10 mM Tris, pH 8.0, 1 mM EDTA (Sigma) 6. Sodium acetate, 3 M, pH 5.2 7. Isopropanol (Molecular Biology grade, Fluka BioChemica) 8. Ethanol, 70% 9. Denaturation solution: 0.5 M NaOH and 1.5 M NaCl 10. Neutralization buffer: 0.5 M Tris-HCl, pH 7.5 11. 20 SSC 3 M NaCl, 0.3 M sodium citrate, pH 7.0 12. Stringent wash buffer I: 2 SSC, 0.1% SDS 13. Stringent wash buffer II: 0.2 SSC, 0.1% SDS 14. Blocking buffer: dilute an appropriate volume of 10 blocking buffer (Roche) 1:10 with 1 maleic acid buffer (Roche).

2.2.2. Additional Reagents

1. Proteinase K (10 mg/ml) (Sigma Aldrich) 2. Trypsin–EDTA (Mediatech Cellgro) 3. Agarose (GibcoBRL) 4. 0.25 HCl solution 5. DNA digestion kit (Roche) 6. Nylon membrane (Schleicher & Schuell)

3. Methods 3.1. Preparation of Cell Lysate for Telomerase Activity

1. Wash cells or tissue in ice-cold PBS and pellet at 1500 rpm for 5 min at 4C. 2. Resuspend cells or tissue in ice-cold wash buffer and centrifuge again at 1500 rpm for 5 min at 4C. 3. Resuspend cells in ice-cold lysis buffer. For tissue, homogenize gently on ice. Incubate on ice for 30 min.


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4. Centrifuge at 14,000 rpm for 20 min at 4C. 5. Carefully separate supernatant and measure the protein concentration. At this step, samples can be stored at –80C in small aliquots to avoid repeated freezing and thawing. 3.2. TRAP Assay

1. Prepare the following TRAP reaction in a sterile 0.1 ml PCR tube: 5 TRAP buffer 5.0 ml dNTP (50 mM) 1.0 ml TS primer 1.0 ml Taq-DNA polymerase 0.5 ml Protein lysate (1–5 mg) H2O to 25.0 ml 2. Set up the negative control reaction at the same time as the above test reaction, include all of the components listed above, but omit the protein lysate. 3. Mix the component gently and incubate the reaction for 30 min at 23C followed by 90C for 3 min in a thermocycler. 4. To each tube add the following: ACX primer 1.0 ml NT primer 1.0 ml TSNT primer 1.0 ml 5. Amplify the telomere repeats using the denaturation, annealing, and polymerization times and temperature listed below:

3.3. PAGE Analysis

Cycle number

Denaturation

Annealing

Polymerization

27 cycles

30 s at 94C

30 s at 50C

1.5 min at 72C

1. Use a vertical electrophoresis apparatus constructed to hold 22 cm glass plates for casting the gel. Spacers vary in thickness from 0.5 to 2.00 mm. However, 1 mm spacers are recommended to run TRAP products, as they produce the sharpest and flattest DNA bands. 2. After polymerization is complete, place the gel in the electrophoresis tank. Fill the reservoirs of the electrophoresis tank with 0.5 TBE buffer. 3. Mix an appropriate amount of DNA gel loading buffer with the TRAP samples and load the mixture into the wells using a gel loading micropipette. Run the gel until the marker dye migrates to the bottom of the plate. 4. Dilute SYBR Green I in TBE buffer (pH 8.0) 1:10,000 just before use (16).

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5. Transfer the gel to a shallow tray containing diluted SYBR Green I. Fifty milliliters of staining solution is required to completely submerge the 20/20 cm gel. Cover the tray with aluminum foil or place it in the dark and incubate the gel at room temperature with gentle agitation for 5 min. No destaining is required. 6. View the bands under UV in an imaging system. SYBR Green I is maximally excited at 497 nm and the fluorescence emission of SYBR Green I stain bound to DNA is at 520 nm. 3.4 The Telomere Length Assay

3.4.1 Isolation of Genomic DNA

1. Use a cell scraper or trypsin to detach cells from the tissue culture dish. Centrifuge the cells at 1200 rpm for 10 min at 10C. Remove the supernatant and resuspend the cell pellet in 1 PBS. Wash the cell pellet twice with 10 ml 1 PBS, centrifuging between washes. 2. Resuspend the cell pellet in 1 ml DNA buffer. Centrifuge the cells at 1200 rpm for 10 min at 10C. Remove the supernatant. 3. Add 300 ml of DNA buffer to resuspend the cell pellet. Then add 5 ml of proteinase K (10 mg/ml) and 15.25 ml of 10% SDS. Shake gently and incubate overnight at 45C. 4. Add 360 ml of phenol/chloroform/isoamylalcohol or a total amount equal to the volume of the supernatant. Shake by hand for 10 min at room temperature. Centrifuge at 3000 rpm for 10 min at 10C. 5. Transfer the supernatant to a new tube and measure the volume. Add 1/10 the volume of 3 M sodium acetate (pH 5.2) and 3 the volume of 100% isopropanol. Shake gently until the DNA is precipitated. 6. Use a sterile glass pipette to transfer the precipitated DNA into a tube with 200 ml of 70% ethanol. Place the tube inverted on a rack for 2 h. Transfer the DNA into a sterile eppendorf tube. 7. Centrifuge at 10,000 rpm for 10 min at 10C. Dry the DNA pellet in a SpeedVac for 5 min. Dissolve the DNA in 100–200 ml of sterile water. Place the tube in an eppendorf thermomixer shaker overnight at 37C. 8. Measure the DNA concentration and run 1–5 ml of the DNA on a 1% agarose gel.

3.4.2 Digestion of Genomic DNA

1. Prepare DNA digestion enzyme mixture by mixing 20 U/ml each of Hinf1 and Rsa1 (Roche). About 20 U/ml enzyme mix is needed for each sample. 2. Dilute 1–2 mg of purified genomic DNA with nuclease-free water to a final volume of 17 ml. 3. For each sample, add 2 ml of 10 digestion buffer (Roche) and 1 ml of enzyme mixture and mix gently.


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4. Incubate the reaction mixture at 37C for 2 h. 5. After the incubation add appropriate amount of DNA gel loading buffer to stop the reaction and quick spin vials. 3.4.3 Gel Electrophoresis

1. Prepare a 0.8% agarose gel in 1 TAE buffer. 2. Load digested genomic DNA samples. 3. Mix 4 ml DIG molecular weight marker (Roche), 12 ml nuclease-free water, and appropriate volume of DNA gel loading buffer. Load 10 ml on either side of the samples to measure length accurately. 4. Run gel at 5 V/cm in 1 TAE buffer until the bromophenol blue dye is separated about 10 cm from the starting wells. 5. Submerge the gel in HCl solution for 10 min until the bromophenol blue stain changes to yellow, with agitation, at room temperature. 6. Rinse the gel two times with H2O. 7. Submerge the gel in the denaturation solution for 2 15 min at room temperature. 8. Rinse the gel two times with H2O. 9. Submerge the gel in the neutralization solution for 2 15 min at room temperature.

3.4.4 Southern Transfer to Nylon Membrane

It is recommended that powder-free rubber gloves be worn and handle membrane with forceps only at the edges. 1. Set up the transfer in a large electrophoresis tray. Fill the tray with transfer buffer (10 SSC). Assemble the following items (gel-sized and saturated with transfer buffer) in the middle support. Carefully remove all air bubbles. – Wick (Whatman 3 MM paper): This should be the same width as your gel and long enough to drape into the transfer buffer. – Three pieces of Whatman 3MM paper – Gel (upside down) – Nylon membrane. (Handle with clean gloves and bluntended forceps. Do not adjust the membrane once it is placed on the gel.) Cut off the lower left hand corner of the Nylon membrane for orientation – Three pieces of Whatman 3MM paper – Dry 5 cm stack of paper towels cut to the size of the gel glass plate and then a weight on top 2. Surround the transfer tray with plastic wrap or parafilm to prevent evaporation of the transfer buffer. Let transfer overnight. 3. After the Southern transfer, fix the transferred DNA on the wet blotting membrane by UV-crosslinking (120 mJ).

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4. Wash the blotting membrane with 2 SSC. 5. If not used immediately for the hybridization, membrane can be air dried and store at 4C. 3.4.5 Hybridization and Chemiluminescence Detection

1. Prewarm 25 ml of DIG hybridization solution to 42C. 2. Prehybridize the blot by submersing the blot in 18 ml of prewarmed DIG hybridization buffer and incubate for 30–60 min at 42C with gentle agitation. 3. Prepare hybridization buffer solution by adding 1 ml telomere probe (Roche) in 5 ml fresh prewarmed DIG hybridization buffer and mix. 4. Discard prehybridization buffer and immediately add hybridization solution to the membrane. 5. Incubate for 3 h at 42C with gently agitation. 6. Wash the membrane two times with sufficient stringent wash buffer I for 5 min at room temperature with gentle agitation. 7. Wash the membrane two times with sufficient prewarmed stringent wash buffer II for 20 min at room temperature at 50C with gently agitation. 8. Rinse membrane in 25 ml 1 washing buffer for 5 min at room temperature in gentle agitation. 9. Incubate membrane in 25 ml freshly prepared blocking solution for 30 min at room temperature with gentle agitation. 10. Dilute appropriate volume of anti-DIG-AP (Roche) with blocking solution to a final concentration of 75 mU/ml. 11. Remove blocking solution and incubate membrane in 25 ml anti-DIG-AP solution for 30 min at room temperature with gentle agitation. 12. Wash membrane two times for 15 min with 25 ml of 1 washing buffer (Roche) at room temperature with gentle agitation. 13. Incubate membrane in 20 ml of detection buffer (Roche) at room temperature with gentle agitation. 14. Discard detection buffer and remove excess liquid by placing on absorbent paper. Do not dry the membrane. 15. Immediately place the wet membrane, the DNA side facing up, on a sheet of polyethylene wrap and quickly apply approximately 40 drops substrate solution (Roche) on to the membrane. 16. Immediately cover the membrane with the second sheet of polyethylene wrap to spread the substrate solution homogeneously and without air bubbles over the membrane.


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17. Expose membrane to the imaging device or to X-ray film for 5–20 min. 3.4.6 Telomere Repeat Fragment Analysis

After exposure of the blot to an X-ray film, an estimation of the mean TRF length can be obtained by visually comparing the mean size of the telomeric DNA to the molecular weight marker.

4. Notes 1. Add PMSF to cell/tissue lysis buffer just before use (Section 2.1.1). 2. Filter 5 TRAP buffer and store at –20C in aliquots (Section 2.1.1). 3. Taking into account the size of the electrophoresis glass plates and the thickness of the spacers, calculate the volume of gel required. Samples should be run in 10% PAGE with 22 cm long plates in order to see well-separated telomere products as shown in representative picture (Section 3.1.3). 4. SYBR Green I is a light sensitive dye. Protect the solution from light all the time (Section 3.1.3). 5. Prepare and pipette the genomic DNA digestion mixture on ice. Prepare master mix for all samples to be analyzed. (Section 3.2.2) 6. The volume of DIG hybridization solution, washing buffer, telomere probe, and blocking solution are recommended for 200 cm2 membrane size. (Section 3.2.5) 7. Prepare anti-DIG-AP solution before use and do not store. (Section 3.2.5) 8. Luminescence intensity of the membrane, after treated with detection buffer, will be high during the first hour. However, it continues for at least 24 h. (Section 3.2.5)

References 1. Greider, C.W., and Blackburn, E.H. 1989. A telomeric sequence in the RNA of Tetrahymena telomerase required for telomere repeat synthesis. Nature 337:331–337. 2. Kim, N.W., Piatyszek, M.A., Prowse, K.R., Harley, C.B., West, M.D., Ho, P.L., Coviello, G.M., Wright, W.E., Weinrich, S.L., and Shay, J.W. 1994. Specific association of human telomerase activity with immortal cells and cancer. Science 266:2011–2015.

3. Harley, C.B., and Villeponteau, B. 1995. Telomeres and telomerase in aging and cancer. Curr. Opin. Genet. Dev. 5 5:249–255. 4. Kilian, A., Bowtell, D.D., Abud, H.E., Hime, G.R., Venter, D.J., Keese, P.K., Duncan, E.L., Reddel, R.R., and Jefferson, R.A. 1997. Isolation of a candidate human telomerase catalytic subunit gene, which reveals complex splicing patterns in different cell types. Hum. Mol. Genet. 6:2011–2019.

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5. Feng, J., Funk, W.D., Wang, S.S., Weinrich, S.L., Avilion, A.A., Chiu, C.P., Adams, R.R., Chang, E., Allsopp, R.C., Yu, J., et al. 1995. The RNA component of human telomerase. Science 269:1236–1241. 6. Harrington, L., McPhail, T., Mar, V., Zhou, W., Oulton, R., Bass, M.B., Arruda, I., and Robinson, M.O. 1997. A mammalian telomerase-associated protein. Science 275:973–977. 7. Holt, S.E., Aisner, D.L., Baur, J., Tesmer, V.M., Dy, M., Ouellette, M., Trager, J.B., Morin, G.B., Toft, D.O., Shay, J.W., et al. 1999. Functional requirement of p23 and Hsp90 in telomerase complexes. Genes Dev. 13:817–826. 8. Nakano, K., Watney, E., and McDougall, J.K. 1998. Telomerase activity and expression of telomerase RNA component and telomerase catalytic subunit gene in cervical cancer. Am. J. Pathol. 153:857–864. 9. Boldrini, L., Faviana, P., Gisfredi, S., Zucconi, Y., Di Quirico, D., Donati, V., Berti, P., Spisni, R., Galleri, D., Materazzi, G., et al. 2002. Evaluation of telomerase in the development and progression of colon cancer. Int. J. Mol. Med. 10:589–592. 10. Park, T.W., Riethdorf, S., Riethdorf, L., Loning, T., and Janicke, F. 1999.

11.

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Differential telomerase activity, expression of the telomerase catalytic sub-unit and telomerase-RNA in ovarian tumors. Int. J. Cancer 84:426–431. Kanaya, T., Kyo, S., Takakura, M., Ito, H., Namiki, M., and Inoue, M. 1998. hTERT is a critical determinant of telomerase activity in renal-cell carcinoma. Int. J. Cancer 78:539–543. Blackburn, E.H. 2001. Switching and signaling at the telomere. Cell 106:661–673. Collins, K., and Mitchell, J.R. 2002. Telomerase in the human organism. Oncogene 21:564–579. Blasco, M.A. 2005. Telomeres and human disease: ageing, cancer and beyond. Nat. Rev. Genet. 6:611–622. Kim, N.W., and Wu, F. 1997. Advances in quantification and characterization of telomerase activity by the telomeric repeat amplification protocol (TRAP). Nucleic Acids Res. 25:2595–2597. Jagadeesh, S., Kyo, S., and Banerjee, P.P. 2006. Genistein represses telomerase activity via both transcriptional and posttranslational mechanisms in human prostate cancer cells. Cancer Res. 66:2107–2115.

Chapter 26 Detecting ATM-Dependent Chromatin Modification in DNA Damage and Heat Shock Response Sandeep Misri, Shruti Pandita, and Tej K. Pandita Abstract The ataxia telangiectasia-mutated gene product (ATM), whose loss of function is responsible for ataxia telangiectasia (A-T), is a protein kinase that interacts with several substrates and is implicated in mitogenic signal transduction, chromosome condensation, meiotic recombination, cell-cycle control and telomere maintenance (Pandita, Expert Reviews in Molecular Medicine 5:1–21, 2003; Pandita, Oncogene 21:611–618, 2002; Matsuoka et al., Science 316:1160–1166, 2007). The ATM protein kinase is primarily activated in response to DNA double-strand breaks (DSBs) caused by ionizing radiation (IR) or radiomimetic drugs (Pandita et al., Oncogene 19:1386–1391, 2000). ATM is also activated by heat shock, which occurs independent of DNA damage (Hunt et al., Can Res 69:3010–3017, 2007). ATM is observed at the sites of DNA damage, where it is autophosphorylated and is dissociated from its non-active dimeric form to the active monomeric form (Bakkenist and Kastan, Nature 421:499–506, 2003). The ATM protein appears to be a part of the sensory machinery that detects DSBs during meiosis or mitosis, or breaks consequent to the damage by free radicals. Recent studies support the argument that ATM activation is regulated by chromatin modifications (Gupta, Mol Cell Biol 25:5292–5305, 2005). This review summarizes the multiple approaches used to discern the role of ATM in chromatin modification in response to DNA damage as well as heat shock. Key words: Ataxia telangiectasia, telomerase, DNA double-stranded breaks, chromatin modification, heat shock.

1. Introduction Maintenance of genomic stability depends on appropriate cellular responses to DNA damage (1, 2). This response is based on a complex network of signaling pathways that activate numerous processes and ultimately lead to damage repair and cell survival or death. The precise sequence of events that follow the IR-induced Srikumar P. Chellappan (ed.), Chromatin Protocols: Second Edition, vol. 523 ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2009 DOI 10.1007/978-1-59745-190-1_26 Springerprotocols.com

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DNA damage in cells is still not well understood. It is clear that a complex series of proteins are recruited at different times and in a cell-cycle-dependent manner to stabilize the genome. The ATM protein plays a key role in the sensing, processing and signaling of IR-induced DNA damage through its interactions with various proteins as summarized in Fig. 26.1. Cells deficient in ATM have been shown to have a high frequency of spontaneous chromosomal aberrations, high rates of intrachromosomal recombination and error-prone recombination (3–6). Such cells have a higher initial and residual chromosomal aberrations in G1 and G2 phases after IR exposure as determined by premature chromosome condensation technology (7–9) and by karyotypic examination of metaphase chromosome spreads (9, 10). ATM phosphorylates a number of substrates, including Chk1 and Chk2, which in turn

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Fig. 26.1 ATM phosphorylates several substrates in cellular responses to DNA DSB repair. ATM kinase activity increases immediately after DSBs occur in DNA following exposure to IR. ATM mediates the early stages of the rapid induction of several signaling pathways, which include activation of the DNA-DSB pathway, regulation of the cell-cycle checkpoint controls, activation of stress responses and maintenance of telomeres. ‘P’ with solid arrows indicates reported phosphorylation events; dashed arrows represent possible signaling steps and do not imply direct interaction between proteins; ‘C’ indicates sequestering in cytoplasm; ‘R’ indicates repair complexes; and ‘T’ indicates a role for the protein in telomere metabolism. AP-1, apetala1 transcription factor; BRCA1, breast cancer susceptibility gene product 1; c-Abl, abelson protein tyrosine kinase; CDK, cyclin-dependent kinase; CHK, checkpoint kinase; FANCD2, fanconi anemia protein; JNK, Jun N-terminal protein kinase; MRE11, meiotic recombination 11 gene product; MDM2, mouse double minute 2 (p53-binding protein); NBS1, Nijmegen breakage syndrome 1 protein (p95); SMC1, structural maintenance of chromosome 1; RAD50, a radiation-damage-repair-associated protein; TRF1, telomere-repeat-finding factor 1; hTERT, human catalytic unit of telomerase; hMOF, the human ortholog of the Drosophila MOF gene (males absent on the first).

Detecting ATM-Dependent Chromatin Modification

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target other proteins to induce cell-cycle arrest and facilitate DNA repair (Fig. 26.1). Some of these molecular targets include ATR, c-Abl, Chk-1, Chk-2, RPA, BRCA1, BRCA2, NF-B/IB alpha, beta-adaptin, MDC1, H2AX and autophosphorylation of ATM itself. Thus, ATM is a hierarchical kinase, capable of initiating many pathways simultaneously (3, 4, 11). ATM has a critical role in maintaining chromosome condensation in the vicinity of recombination intermediates (3, 12). ATM phosphorylates structural maintenance chromosome protein ‘‘Smc1’’, necessary for sister chromatid cohesion (3, 13, 14). Several studies indicate that the MRN complex as well as TRF2 either influences activation of ATM or serve as a modulator/amplifier of ATM activity (1, 3, 15–17). Simultaneous studies on DNA and chromosome damage after IR exposure suggested that A-T cells are more proficient in translating the DNA damage into chromosome damage (7, 8). It is possible that A-T cells have an inherent chromatin alteration that allows a more efficient translation of DNA damage into chromosome damage following IR exposure (18). ATM is held inactive in unirradiated cells as a dimer or higher-order multimer, with kinase bound to a region surrounding serine 1981, and IR exposure induces rapid autophosphorylation of ATM of serine 1981 that causes dimer dissociation and initiates cellular ATM kinase activity. It is proposed that the conversion of dimer to monomer status of ATM is regulated by the status of chromatin structure (19). Chromatin-modifying factor ‘‘hMOF’’ influences IR-induced initiation of ATM kinase activity (20). Furthermore, recent studies have revealed that heat shock which induces chromatin alterations independent of DNA damage, also activates ATM (10). 1.1. Chromatin Structure and Radiation Sensitivity

Chromatin, the physiological packaging structure of histones and DNA, is now gaining appreciation as a relevant target of signaling pathways. Chromatin structure is an important factor for determining protein–DNA interactions, with consequences for DNA metabolism and transcription control. After DNA damage, eukaryotic cells orchestrate a complex array of responses. The coordination of checkpoint-mediated cell-cycle arrest and the repair of damaged DNA are essential for cell survival. Chromatin structural alterations like histone modification by phosphorylation, acetylation or deacetylation have all been linked to the repair of damaged DNA. Histone H2AX, the major isoform in yeast and a minor H2A species in mammals, is phosphorylated at the carboxy-terminal serine 139 immediately following Spo11-induced DSB formation (21). Phosphorylated H2AX (termed-H2AX) appears within minutes of damage over large chromatin regions extending tens of kb in yeast and up to 2 Mb in mammalian cells (22). The analysis of

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H2AX-deficient mice has demonstrated a role for H2AX in a variety of responses to DSBs, including DNA repair, checkpoint signaling and immunoglobulin gene class switching (23). H2AX/ mice exhibit male-specific sterility, which is likely due to defects in chromatin remodeling during meiosis. How histone modifications participate in either the repair process or the checkpoint control is unclear. There is very limited information available about the role of histone modifying factors in IR response. Identification of chromatin-modifying factors that play a role in IR response will help to understand the mechanisms how chromatin modifications induced by IR trigger ATM activation. Protocols presented in this chapter describe how to analyze DNA damage using different methods. 1.2. ChromatinModifying Factors and DNA Repair

DNA repair process involves alterations in chromatin structure, which are required for direct access to DNA. This process is facilitated by the chromatin-modifying factors, which detach DNA from histones and other bound proteins. Post-translational covalent modifications of histones have emerged as key regulatory events in DNA damage response. A widespread modification is acetylation catalyzed by histone acetyl-transferases and reversed by deacetylases. Reversible acetylation of four lysines (K) at positions 5, 8, 12 and 16 in the amino-terminal tail of histone H4 occur in vivo in all eukaryotes (24, 25). Hyperacetylation of histone H4 could lead to unfolding of the nucleosomal fiber (26) and acetylation of histone H4 at K16 occurs on the hyperactive male X chromosome of Drosophila polytene chromosomes (27). Recent studies revealed that histone H4-K16 controls chromatin structure and protein’s interactions (28). Tip60 (Tat-interacting protein) interacts with ATM, acetylates histones (H2A, H3, H4) and plays a role in DNA repair (29, 30). In Drosophila Tip60 acetylates nucleosomal phospho-H2Av and exchanges it with an unmodified H2Av (31). Bird et al. (32) reported that the acetylation of histone H4 by Esa1 (essential SAS2-related acetyltransferase) is required for DNA repair in yeast and suggested that a similar modification may function in mammalian cells. ATM has been reported to regulate the function of some histone modifying factors. Gupta and coworkers identified hMOF as an ATM-interacting protein and have provided evidence that functional hMOF participates in the activation of ATM in response to DNA damage (20).

1.3. Determination of ATM-Interacting Proteins

Various approaches are used to determine the interactions between the proteins. For in vivo analysis, most commonly used is immunoprecipitation following by immunoblotting. These techniques are also described below.


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2. Materials 2.1. Immuno precipitation and Coimmunoprecipitation

1. Protein extract, protein A/G beads and antibodies. 2. Immunoprecipitation (IP) buffer: 50 mM Tris–HCl pH 8.0, 150 mM NaCl, 1% NP-40. 3. Protease inhibitor cocktail (100 ): 50 mg/ml PMSF, 1 mg/ ml aprotinin, 1 mg/ml leupeptin, 1 mg/ml pepstatin in 100% ethanol; store at –20C. 4. Sample loading buffer: 50 mM Tris–HCl pH 6.8, 100 mM dithiothreitol (DTT), 2% SDS, 0.2% bromphenol blue, 20% glycerol.

2.2. SDS-PAGE Analysis

1. SDS-PAGE stacking gel (10-ml total volume): 7.0 ml H2O, 1.25 ml 1 M Tris–HCl pH 6.8, 1.5 ml 30% acrylamide solution, 100 ml 10% SDS, 100 ml 10% APS, 10 ml TEMED. 2. 8% SDS-PAGE separating gel (10-ml total volume): 4.6 ml H2O, 2.5 ml 1.5 M Tris–HCl pH 8.8, 2.7 ml 30% acrylamide solution, 100 ml 10% SDS, 100 ml 10% adenosine 50 -phosphosulfate (APS), 6 ml TEMED. 3. 12% SDS-PAGE separating gel (based on 10-ml total volume): 3.3 ml H2O, 2.5 ml 1.5 M Tris–HCl pH 8.8, 4.0 ml 30% acrylamide solution, 100 ml 10% SDS, 100 ml 10% APS, 4 ml TEMED. 4. SDS-PAGE running buffer (10 ): 30.3 g Tris–HCl, 144 g glycine, 10 g SDS pH to 8.3 to 1 l with water. 5. Coomassie stain solution (1 l total volume): 1.25 g Coomassie brilliant blue (R250), 500 ml methanol, 100 ml glacial acetic acid, 400 ml H2O. 6. Coomassie destain solution: 100 ml methanol, 100 ml glacial acetic acid, 800 ml H2O.

2.3. Western Blot Analysis

1. Blocking solution (5% milk powder in TBST [pH 7.4]). 2. Dulbecco’s modified Eagle medium (DMEM). 3. ECL (enhanced chemiluminescent) detection kit. 4. Electrotransfer buffer 25 mM Tris, 192 mM glycine, 0.01% SDS and 20% methanol. 5. 2 Laemmli SDS sample buffer: 4 ml 10%w/v SDS, 2 ml glycerol, 1.2 ml 1 M Tris–Cl (pH 6.8) and 2.8 ml water. 6. Phosphate-buffered saline (PBS) (pH 7.1): 1.37 M NaCl, 27 mM KCl, 100 mM Na2HPO4, 10 mM CaCl2.2H20 and 5 mM MgCl2.6H20. To prepare 1 l of either 1 or 10 PBS, dissolve the reagents listed above in 800 ml of H2O.

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Adjust the pH to 7.4 (or 7.2, if required) with HCl and then add H2O to 1 l 7. Primary and secondary (Horseradish peroxidase [HRP]-conjugated) antibodies 8. TBST (pH 7.4) 20 mM Tris–HCl, 150 mM NaCl and 0.2 % Tweeen-20 9. Centrifugation tube (1.5 ml) 10. Electrotransfer equipment 11. Enhanced chemiluminescent detection equipment 12. Water bath (boiling) 13. Nitrocellulose membrane 14. SDS-polyacrylamide gel electrophoresis equipment. 2.4. ATM Kinase Assay

ATM is a serine/threonine kinase and specifically phosphorylates SQ/TQ sites. To determine the substrates of ATM phosphorylation, in vitro kinase assays are performed. Cells are broken open in lysis buffer. Afterwards, lysates were precleaned for 1 h at 4C with 10 mg rabbit purified IgG and 40 ml protein A/G beads. Endogenous ATM proteins are immunoprecipitated with anti-ATM antibodies and protein A/G sepharose. Immunoprecipitants are washed three times with lysis buffer, once with lysis buffer and 0.5 M LiCl and three times with kinase buffer. Immunoprecipitant beads are mixed with kinase buffer containing 10 mC [ -32P-ATP] and 1 mg GST-p5311–101, incubated for 20 min at 30C. The kinase reactions are stopped with SDS –PAGE loading dye, and reaction mixtures are separated by SDS – PAGE. 1. Lysis buffer : 50 mM Tris pH 7.5, 150 mM NaCl, 1% Tween 20, 0.2% NP-40, 1 mM NaF, 1 mM NaVO4, 50 mM glycerophosphate, 10% glycerol, 1 mM phenylmethylsulfonyl fluoride, 2 mg/ml pepstatin A, 5 mg/ml leupeptin, 10 mg/ml aprotinin and 1 mM dithiothreitol (DTT) 2. Anti-ATM antibodies and protein A/G sepharose 3. 0.5 M LiCl 4. Kinase buffer: 50 mM HEPES pH 7.5, 150 mM NaCl, 1% Tween 20, 0.2% NP-40, 1 mM NaF, 1 mM NaVO4, 1 mM DTT and 10 mM MnCl2

2.5. Immunostaining

1. Antibodies, primary and secondary. 2. Blocking solution (5% normal goat serum in PBTD). 3. Paraformaldehyde (PFA) (4%) in 1 PBS: paraformaldehyde, EM grade, PBS (10 ), NaOH (1 M) and HCl (1 M); for a 4% paraformaldehyde solution, add 4 g of EM grade paraformaldehyde to 50 ml of H2O. Add 1 ml of 1 M NaOH and stir gently on a heating block at 60C until the paraformaldehyde


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is dissolved. Add 10 ml of 10 PBS and allow the mixture to cool to room temperature. Adjust the pH to 7.4 with 1 M HCl (1 ml), then adjust the final volume to 100 ml with H2O. Filter the solution. 4. 1 phosphate-buffered saline (PBS) and Vectashield mounting medium (Vector Labs). 5. Coverslips (No. 1 thickness) and microslides (1.0 mm thick), precleaned. 6. Microscope, confocal or fluorescent or light. 7. PAP pen (Sigma) and tubes (1.5 ml microcentrifuge). 2.6. Chromosomal DNA Strand Break Analysis

Two different assays are used to measure DNA strand breaks after treatment with heat shock or IR exposure or heat shock + IR exposure. Biochemical methods such as (a) pulsed field gel electrophoresis (PFGE) (10) and (b) neutral filter elution (8, 20). Cytological approaches involve either the analysis of chromosomes at metaphase or the interphase using the premature chromosome condensation technique as described previously (7, 23).

2.6.1. Pulsed Field Gel Electrophoresis (PFGE)

1. Tris–EDTA buffer (10 mmol/l Tris–HCl, 1 mmol/L Na2EDTA [pH 8]) 2. 0.8% agarose gels, ethidium bromide, NaCl and Triton X-100 3. CHEF DRIII system (Bio-Rad, Hercules, CA) 4. Charge-coupled device transillumination

2.6.2. Neutral Filter Elution

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Filter elution is a commonly used technique for measuring DNA strand breakage. The assay uses a filter to mechanically impede passage of long (unbroken) DNA strands during elution. Doublestrand DNA breaks are detected with ‘‘neutral’’ filter elution (performed at pH 9.6). This pH is nondenaturing thus no DNA strand unwinding occurs. The eluting solution is composed of 0.05 M Tris–Cl/0.05 M glycine/0.025 M EDTA (disodium salt)/2% (w/v) sodium dodecyl sulfate. The pH of the eluting solution is adjusted to pH 9.6 with 10 N NaOH. DNA quantitation is conducted as above by prelabeling cells with radiolabeled thymidine or by using fluorometry. 1. Cells of interest and appropriate growth medium for cells 2. [Methyl 3H]thymidine (20 Ci/mmol) or [2–14C]thymidine (50 mCi/mmol) for radioactive determination 3. Unlabeled thymidine and test compound 4. Phosphate-buffered saline (PBS), ice-cold: 137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2HPO4. 7H2O and 1.4 mM KH2PO4

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5. Cell lysis solution, pH 9.6: 2% (w/v) sodium dodecyl sulfate (SDS), 0.025 M disodium EDTA and 0.1 M glycine (optional). Adjust pH to 9.6 with 1 N NaOH 6. Cell lysis solution (above) containing 0.5 mg/ml proteinase K 7. Alkaline elution solution, pH 12.3: 0.1% (w/v) sodium dodecyl sulfate, 20 mM tetrasodium EDTA, 0.8% tetrapropylammonium hydroxide. Adjust pH to 12.3 with 20% (w/v) tetrapropylammonium hydroxide in water 8. Scintillation fluid (e.g., Ecolume, ICN Biomedicals) containing 0.7% (v/v) acetic acid, 0.2 M tetrasodium EDTA pH 10 (pH adjusted with 1 N NaOH) 9. 1 N HCl 10. 0.4 N NaOH 11. 1 M potassium dihydrogen phosphate 12. 17 mM KH2PO4 13. 150 mM Hoechst 33258 stock solution 14. Tabletop centrifuge and 2.0-mm polycarbonate filter (25-mm diameter) 15. 1 l vacuum flask, 60C oven or water bath 16. Alkaline elution funnel apparatus (e.g., Millipore) 17. Aluminum foil or black paper cylinder 18. Multichannel peristaltic pump and suitable Tygon tubing 19. Fraction collector 20. Glass scintillation vials and liquid scintillation counter or fluorometer

3. Methods 3.1. Immuno precipitation and Coimmunoprecipitation

The beads selected to bind the antibodies depend on the species and subclass of the antibody used (e.g., protein A beads for rabbit, mouse IgG2a, IgG2b and IgG3; and protein G beads for mouse IgG1 and most subclasses of rat IgGs). 1. Add 5–10 mg of specific antibody or control antibody to the eppendorf tube containing the cold extract (approx. 300 mg). 2. Adjust the volume to 500 ml with IP buffer and add 5 ml of protease inhibitor cocktail. Incubate at 4C for 1–4 h. 3. Wash 50 ml of protein G slurry by adding 450 ml cold IP buffer and spinning for 5 s in a microcentrifuge. Remove supernatant and repeat the washing step twice. Resuspend the beads in 50 ml of cold IP buffer.


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4. Add the prewashed 50 ml of protein G slurry to the extract. Incubate for 1 h at 4C on a rolling wheel and spin for 5 s at 4C. Carefully remove the supernatant completely and wash the beads three to five times with 500 ml of IP buffer. 5. After the last wash, aspirate supernatant and add 50 ml of sample loading buffer to bead pellet. Vortex and heat to 100C for 5 min. Spin at 10,000 g for 5 min collect the supernatant and load onto an SDS-polyacrylamide gel. 3.2. SDS-PAGE

1. Remove medium from the cells. Rinse the cells once with 200 ml of PBS and add 200 ml of trypsin–EDTA. Incubate for 1 min at 37C; suspend the cells and add 800 ml of DMEM medium to quench the trypsin. 2. Transfer the suspended cells to a chilled 1.5-ml centrifugation tube. Collect the cells by centrifugation at 700 g for 4 min at 4C. Resuspend the cell pellet in ice-cold PBS and centrifuge again. 3. Treat the cells with ice-cold buffer containing 50 mM Tris–HCl pH 7.5, 150 mM NaCl, 1% NP40, 1 mM DTT, 1 mM EDTA, 50 mM glycerophosphate, 10% glycerol, 1 mM phenylmethylsulfonyl fluoride, 2 mg/ml pepstatin A, 5 mg/ml leupeptin, 10 mg/ml aprotinin and 1 mM dithiothreitol (DTT) at 4C for 30 min. Centrifuge again. 4. Remove the supernatant and add 25 ml of 90C 2X Laemmli SDS sample buffer to the cell pellet obtained. Incubate the sample for 3 min in a boiling water bath and vortex. 5. Separate the proteins by SDS-polyacrylamide gel electrophoresis using an acrylamide concentration appropriate to resolve the molecular weight of the targeted protein 6. Transfer proteins from the gel to a nitrocellulose membrane using electrotransfer buffer.

3.3. Immunoblotting

1. Incubate the membrane in blocking solution (PBS solution containing 0.05% Tween-20 and 1% nonfat dried milk, to block non-specific binding) for 1 h at 37C. 2. Replenish the blocking solution with fresh blocking solution and add the primary antibody at the appropriate dilution. Incubate for 1–2 h at 37C. Wash the blot four times with TBST for 10 min. For ECL detection, incubate the blot with either HRPconjugated secondary antibody at a suitable dilution in blocking solution for 1–2 h at 37C. Perform ECL detection.

3.4. ATM Kinase Assay

1. Cells are broken open in lysis buffer (50 mM Tris pH 7.5, 150 mM NaCl, 1% Tween 20, 0.2% NP-40, 1 mM NaF, 1 mM NaVO4, 50 mM glycerophosphate, 10% glycerol, 1 mM

404


phenylmethylsulfonyl fluoride, 2 mg/ml pepstatin A, 5 mg/ml leupeptin, 10 mg/ml aprotinin and 1 mM dithiothreitol (DTT). 2. Lysates are precleaned for 1 h at 4C with 10 mg rabbit purified IgG and 40 ml protein A/G beads. 3. Endogenous ATM proteins are immunoprecipitated with antiATM antibodies and protein A/G sepharose. Immunoprecipitants are washed three times with lysis buffer, once with lysis buffer and 0.5 M LiCl, and three times with kinase buffer (50 mM HEPES pH 7.5, 150 mM NaCl, 1% Tween 20, 0.2% NP-40, 1 mM NaF, 1 mM NaVO4, 1 mM DTT, 10 mM MnCl2). 4. Immunoprecipitant beads are mixed with kinase buffer containing 10 [mC 32P]ATP and 1 mg GST-p531–101, incubated for 20 min at 30C. The kinase reactions are stopped with SDS-PAGE loading dye and reaction mixtures are separated by SDS – PAGE. 5. Transferred ATM and substrates are visualized and quantitated on a PhosphorImager. 6. ATM proteins are detected using anti-ATM antibodies. Several approaches are used to determine ATM autophosphorylation and ATM-dependent phosphorylation. These include mass spectrometry, Western blotting and cytological analysis. First phosphorylation sites of protein substrate are identified by MALDI-TOF mass spectroscopy. Once the residue(s) are identified, phosphospecific antibodies are generated. Western blotting or immunostaining is used to detect the phosphorylated form of proteins. 3.5. Immunostaining

1. Cells are grown on coverslips and washed with PBS. 2. Incubate the samples in 4% PFA overnight at 4C or for 4–6 h at room temperature. 3. Rinse the specimens three times in 1 PBS for 5 min each (total time of 15 min). 4. Permeabilize the cells for 20 min in 0.5 % NP-40. Circle the area of interest on each slide with a hydrophobic PAP pen. Gently pipette 200–300 ml of blocking solution onto each slide. Incubate the slides for 1–2 h at room temperature. 5. Remove the blocking solution by draining the excess. Add 200–300 ml of primary antibody diluted in blocking solution to each slide. Incubate the slides overnight at 4C. 6. Remove the primary antibody by rinsing the slides three times in PBS for 10 min each at room temperature. Drain the excess PBS and add 200–300 ml of secondary antibody diluted in blocking solution. Incubate the slides for 1–2 h at room temperature. 7. Remove the secondary antibody by rinsing the slides three times in PBS for 10 min each at room temperature. Drain as


405

much PBS from the slides as possible. Add one drop of Vectashield mounting medium directly onto each section. 8. Carefully place a coverslip onto each slide. Allow Vectashield to harden for at least 3 h at room temperature before imaging. Image the slides on a confocal or fluorescent microscope. 3.6. Chromosomal DNA Strand Break Analysis

For analyzing the chromosome DNA damage biochemical as well as cytological approaches are used. The biochemical approaches are described here mainly.

3.6.1. Pulse Field gel Electrophoresis

1. In the PFGE, cells are harvested, embedded in agarose plugs and lysed in situ. 2. Plugs are washed in Tris–EDTA buffer (10 mmol/l Tris–HCl, 1 mmol/l Na2EDTA [pH 8]) and PFGE is carried out with a CHEF DRIII system in 0.8% agarose gels. 3. The gels are run at 14C with linearly increasing pulse times from 50 to 5,000 s for 192 h at field strength of 1.5 V/cm. 4. Subsequently, gels are ethidium bromide stained and photographed with a charge-coupled device camera system under UV transillumination. 5. Quantitative analysis to determine the fraction of DNA entering the gel provides a measure for the relative number of double-strand breaks. Control cell DNA is normalized to zero and DNA of cells treated with 10 Gy is given a value of 1.

3.6.2. Neutral Filter Elution

1. Label cells with [14C] or [3H]thymidine – Expose 105 cells/ml in suspension or a monolayer culture of 106 cells to 0.1 mCi/ ml [methyl 3H]-thymidine plus 105 M unlabeled thymidine or 0.01 to 0.02 mCi/ml of [2–14C]-thymidine for 24 h. Grow and label an adequate number of cells 2. Recover cells in suspension (by centrifuging 5 min at 200 g, 4C) or in monolayer (by rinsing twice with ice-cold PBS, scraping the cells with a rubber policeman and centrifuging 5 min at 200 g, 4C) and remove the supernatant. 3. Rinse pellets with ice-cold PBS, centrifuge again, remove the supernatant and transfer an appropriate number of cells into the appropriate medium for treatment with test agent. 4. Assemble and prepare filter apparatus – Secure the filter within the funnel apparatus and attach to a suction flask that is in turn connected to a vacuum line. Add 20 ml of ice-cold PBS to the funnel. Apply a full vacuum for a few seconds to expel air from the filter. Adjust vacuum to a slow flow rate. 5. Apply cells to filter – Add 0.5–2 106 cells, diluted in 20 ml ice-cold PBS, to the funnel. Continue to apply a gentle vacuum to pull the cells onto the filter.

406


6. Just before the solution level reaches the filter, add 5–10 ml of additional ice-cold PBS to wash the cells. Repeat twice. During the last wash, turn off the vacuum when the fluid is 1 cm above the filter and allow the last of the solution to flow through the filter by gravity. Cover the funnel apparatus, with foil or a black paper cylinder. 7. Lyse the cells – Disconnect the funnel/filter apparatus from the suction flask. Fill the upper chamber of the filter holder with cell lysis solution, pH 9.6, by using a pipettor with a plastic tip that will fit through the filter orifice. Withdraw the tip slightly to allow for air bubbles to escape from the upper chamber. Pipet 1 ml at a time, slowly. Add a total of 5 ml of cell lysis solution. Allow the lysis solution to drip by gravity into a scintillation vial. 8. Digest protein – Connect the outflow tubes to tubing leading to a peristaltic pump (with the pump turned off at this point). Fill the upper chamber of the filter holder with 2 ml of cell lysis solution containing 0.5 mg/ml of proteinase K, using the procedure described above for adding the cell lysis solution. Pump through the filter at 0.035 ml/min. Turn off the pump. 9. Elute-damaged DNA – Aspirate excess proteinase K solution from the filter funnel and add 5 ml of alkaline elution solution, pH 12.3, by slowly running the solution down the wall of the funnel. Pour an additional 30–40 ml of the solution into the funnel. Turn on the pump at a speed of 0.035 ml per min and collect fractions into scintillation vials at 90- or 180min intervals for 15 h. Turn pump off. 10. Quantitate DNA using scintillation counting of radiolabeled DNA – Pour off any remaining elution solution. 11. Turn pump on to maximum speed to empty solution remaining in the line and filter holder. Collect this fraction into a scintillation vial. 12. Turn off the pump and remove the filter. Place it in a scintillation vial containing 0.4 ml of 1 N HCl. Cap the vial and heat at 60C in oven or water bath for 1 h to depurinate the DNA. 13. Add 2.5 ml of 0.4 N NaOH and shake vigorously. Allow to stand for 1 h at room temperature. Shake again. 14. Flush the filter holder and line tubing with 10 ml of 0.4 N NaOH by pumping at high speed. Collect into another scintillation vial. Wash again with 2.5 ml 0.4 N NaOH and count to check if all radioactivities have been removed. 15. Add 10 ml of scintillation fluid (e.g., Ecolume, ICN Biomedicals) containing 0.7% (v/v) acetic acid to each scintillation vial.


407

16. Count the radioactivity in each of the vials containing the filters, rinse solution and eluted fractions. 17. Calculate results by adding the radioactivity (cpm) from the filter to the radioactivity measured in the wash of the filter holder. Add this amount to the cumulative amount of radioactivity in the fractions. 18. Plot data on a semilogarithmic plot of DNA retention (i.e., the fraction of total DNA retained on the filter [log scale]) versus elution time 3.7. Cytogenetic Approach

This method is cell-based analysis of individual chromosome damage provides the most sensitive assay for determining the induction of DNA strand breaks after stress. Essentially, cells are treated with colcemid after IR exposure. Chromosome aberrations are assessed by counting chromatid breaks and gaps per cell. The details of cytogenetical approaches for assessing chromosomal DNA damage are well described in the past (5, 7, 8, 20, 33–39).

4. Notes 1. Chromatin structural perturbations induced by DNA doublestrand breaks have been proposed to serve as a trigger for ATM activation as determined by ATM autophosphorylation and enhanced kinase activity. Two areas of research are being actively pursued to determine which type of chromatin alterations are responsible for the ATM activation and how such alterations can activate ATM function. 2. Hyperthermia (heat shock) enhances ATM autophosphorylation and its kinase activity, which occurs independent of DNA double-strand breaks. 3. To determine the cellular ATM activation, cells must be physiologically viable as determined at least by trepan blue dye exclusion assay. 4. -H2AX foci are the consequence of chromosomal DNA double-strand breaks, however, this does not stand true for heatinduced -H2AX foci, because hyperthermia does not induce DNA double-strand breaks. 5. Both neutral filter elution as well as pulse field gel electrophoresis allow to measure gross chromosomal DNA double-strand breaks, whereas cytogenetic approaches allow to measure DNA double-strand breaks of individual chromosomes of each cell. Thus cytogenetic approach to quantitate chromosomal DNA damage is more sensitive.

408


6. Ionizing radiation-induced chromosomal DNA breaks correlate with the number of -H2AX foci, however, -H2AX foci are seen in S-phase cells without exposure to DNA-damaging agents.

Acknowledgments The authors thank the former and current members of the laboratory, who have carried the work presented in this manuscript. This work was supported by funds from NIH (CA10445, CA123232)

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INDEX A

Analysis of Uniquely Positioned mononucleosomes........................................ 109 Materials .................................................................. 112 Methods................................................................... 113 Notes........................................................................ 121 Antibodies Anti-alpha tubulin ................................................... 312 Anti-BrdU ................................................................. 31 Anti-DIG .................................................................. 72 Anti LacI ................................................................... 35 Anti-NeuN .............................................................. 319 Anti-Phospho-histone H3 Serine 10 ...................... 312 Anti-rabbit, biotinylated.......................................... 312 Methyl specific......................................................... 243 Secondary................................................................... 33 Antigen retrieval ............................................................ 319 Arginine deiminase........................................................ 250 Array normalization and probe analysis ........................ 369 Array processing, ChIP-on-chip ................................... 354 Assay, ATM kinase ............................................... 400, 403 Assay, ChIP-on-chip............................................. 341, 343 Materials .................................................................. 344 Methods................................................................... 347 Notes........................................................................ 361 Assay, Chromatin Immunoprecipitation (ChIP).................. 141, 153, 239, 323, 324, 329 Alternate protocols .................................................. 332 Materials .................................................................. 325 Methods................................................................... 327 Notes........................................................................ 334 On tissues................................................................. 333 Assay, Comet ................................................................. 169 Materials .................................................................. 170 Methods................................................................... 171 Notes........................................................................ 174 Assay, histone deacetylase............................................ 279, 327, 335 Materials .................................................................. 281 Methods................................................................... 284 Notes........................................................................ 291 Assay, histone demethylase.................... 249, 252, 256, 257 Materials .................................................................. 251 Methods................................................................... 253 Notes........................................................................ 258 Assay, histone transfer ................................................... 266

Acrylamide, 30%............................................................ 206 Agarose gel electrophoresis, see Electrophoresis, Agarose gel Agarose, low gelling temperature .................................. 171 Alexa Fluor 488 goat anti-mouse IgG .......................... 312 Alexa Fluor 568 goat anti-rabbit IgG ........................... 312 Analysis of arginine methylation ................................... 235 Materials .................................................................. 238 Methods................................................................... 240 Notes........................................................................ 245 Analysis of ATM-dependent chromatin modification ................................................. 395 Materials .................................................................. 399 Methods................................................................... 402 Notes........................................................................ 407 Analysis of ChIP-on-chip data ...................................... 367 Analysis of DNA Topology in Yeast Chromatin ..................................................... 93 Materials .................................................................... 96 Methods..................................................................... 97 Notes........................................................................ 104 Analysis of genomic aberrations .................................... 177 Materials .................................................................. 178 Methods................................................................... 184 Notes........................................................................ 198 Analysis of genomic methylation................................... 217 Materials .................................................................. 219 Methods................................................................... 221 Notes........................................................................ 232 Analysis of histone chaperones ...................................... 263 Materials .................................................................. 264 Methods................................................................... 267 Notes........................................................................ 276 Analysis of histone H3 phosphorylation by immunostaining ........................................... 311 Materials .................................................................. 312 Methods................................................................... 314 Notes........................................................................ 320 Analysis of Reconstituted Chromatin ..................................................... 11 Materials .................................................................... 13 Methods..................................................................... 15 Notes.......................................................................... 21

411

CHROMATIN PROTOCOLS

412 Index

Assay, histone ubiquitination and deubiquitination........................... 298, 303, 304 Materials .................................................................. 298 Methods................................................................... 299 Notes........................................................................ 306 Assay for premature chromosome condensation .................................................. 83 Materials .................................................................... 85 Methods..................................................................... 86 Notes.......................................................................... 87 Assay, telomerase activity (TRAP)........................ 384, 389 Materials .................................................................. 387 Methods................................................................... 388 Notes........................................................................ 393 Assay, telomerase length........................................ 386, 390 Materials .................................................................. 387 Methods................................................................... 388 Notes........................................................................ 393 Assay, template relaxation ............................................. 272 Assay, in vitro methylation ............................................ 236 Assay, in vitro methyltransferase ........................... 242, 252 Assay, in vitro replication ...................................... 203, 212 Materials .................................................................. 206 Methods................................................................... 209 Notes........................................................................ 213 Assay, in vitro transcription................................... 267, 274 Assay, in vivo Chromatin Decondensation ..................... 27 Materials .................................................................... 29 Methods..................................................................... 31 Notes.......................................................................... 38 Ataxia telangiectasia (AT) ............................................. 395 Ataxia telangiectasia-mutated gene product (ATM) ......................................................... 395 Interacting proteins.................................................. 398

B Baculovirus..................................................................... 240 Beads, Protein A ............................................................ 330 Beads, Protein G............................................................ 330 Binding site analysis and characterization ..................... 378 Binding site visualization............................................... 377 Biotinylated peptides ..................................................... 243 Biotique Systems XRAY for ChIP-on-chip ................. 377 Blocking of non-specific binding .................................. 319 Blotting, Southern ........................................................... 99 Bright-field analysis....................................................... 316 Bromodeoxyuridine.......................................................... 31 Buffer Hybridization............................................................. 97 Lysis ......................................................................... 170 Nuclei lysis ............................................................... 112 Permeabilization ........................................................ 30 Phosphate................................................................... 85 Pre-extraction ............................................................ 30

C Calyculin A .......................................................... 83, 84, 85 CARM1................................................................. 236, 244 Carnoy’s fixative............................................................... 85 ChIP-on-chip ........................................................ 341, 343 ChIP, see Assay, Chromatin immunoprecipitation Chloroquine diphosphate ........................................ 96, 103 Chromatin Analysis of reconstituted............................................ 19 Assembly extracts......................................................... 1 Associated factors, non-histone............................... 145 Decondensation assays......................................... 27, 31 Donor chromatin – chicken erythrocytes .................................................. 114 Immunoprecipitation ...................... 141, 157, 323, 324, 328, 334 Optically highlighted............................................... 125 Photoactivated ......................................................... 133 Preparation............................................................... 114 Reconstituted ......................................................... 2, 11 Reconstitution on immobilised DNA .............................................................. 17 Reconstitution using purified histones ...................... 18 Remodeling.......................................................... 28, 31 Repair....................................................................... 141 Template .................................................................. 274 See also Assay, Chromatin immunoprecipitation Chromosal DNA strand break analysis ......................... 401 Chromosome Condensation, drug induced...................................... 86 Condensation, premature .......................................... 83 Metaphase........................................................ 177, 184 CisGenome .................................................................... 375 Cis-regulatory element annotation system .................... 378 Citrate buffer, pH 6.0.................................................... 314 Cloning of PCR Amplicons ............................................ 58 Comets Moment ................................................................... 174 Quantification.......................................................... 173 Comparative Genomic Hybridization, 177, see Analysis of genomic aberrations Complex, pre-replication............................................... 203 Creatine Phosphokinase .................................................... 4 Crosslinking Agents ...................................................................... 169 Formaldehyde .......................................................... 145 Protein DNA ........................................................... 144 Cytometric analysis of DNA damage............................ 161 Materials .................................................................. 162 Methods................................................................... 163 Notes........................................................................ 166 Cytometry, laser scanning.............................................. 164

CHROMATIN PROTOCOLS

Index 413

D

E

Data analysis methods, ChIp-on-Chip assays .............. 367 Deacetylases Class I ...................................................................... 280 Histone .................................................................... 279 Purification .............................................................. 280 Deamination, bisulfite ..................................................... 50 Demethylase, histone..................................................... 249 Deubiquitination, histone............................................. 295, 297, 303 40 , 6-diamidino-2-phenylindole (DAPI) ................ 30, 163 3’3 Diamnino benzidine (DAB) Peroxidase substrate kit ................................................................. 313 Diethyl pyrocarbonate (DEPC) ...................................... 71 Dimers, cyclobutane pyrimidine............................ 141, 143 DNA Biotinylation .............................................................. 15 Bisulfite deamination................................................. 56 Breaks....................................................................... 125 Connector ................................................................ 103 Coupling to Dynabeads ............................................. 16 Deaminated template........................................... 52, 58 Double-strand breaks, detection.............................. 170 Double-stranded breaks......................................... 161, 170, 395 Extraction, from blood .................................... 180, 186 Extraction, from cell lines........................................ 187 Extraction, from Paraffin embedded tissue..................................................... 180, 188 Fragmentation, apoptotic ........................................ 166 Immobilisation..................................................... 13, 15 Isolation, total yeast ............................................. 49, 54 Methyltransferase....................................................... 41 Preparation................................................................. 98 Sequencing................................................................. 59 Single strand breaks, detection ................................ 170 Template, closed circular ........................................... 93 Topology.................................................................... 93 DNA Methyltransferase Probing of Chromatin Structure ........................................................ 41 Materials .................................................................... 47 Methods..................................................................... 52 Notes.......................................................................... 59 DNAse I .................................................................... 31, 37 Double labeling of cells.................................................. 316 Drosophila preblastoderm embryos .......... 1–25, 247, 267, 398, 409 Chromatin assembly activity.................................... 4, 6 Dechorionation ........................................................ 3–4 Extract preparation ...................................................... 4 Harvesting.................................................................... 3 Dye, loading..................................................................... 96 Dynabeads........................................................................ 13

Electrophoresis Agarose ...................... 7, 20, 96, 98, 115, 170, 171, 182 Low gelling temperature.......................................... 171 Polyacrylamide ........................... 63, 207, 213, 400, 403 Pulse-field ........................................................ 405, 407 Topoisomer resolving ................................................ 98 Embryo, mouse ................................................................ 71 End-labeling, indirect.................................................... 147 Endonuclease, DNase1.................................................. 119 Extraction of immunoprecipitated DNA ...................... 329 Extracts, cytosolic .......................................................... 209 Extracts, nuclear............................................................. 209

F Fixation, cells ................................................................. Flavin adenine dinucleotide, (FAD).............................. Flow cytometry .............................................................. Flow cytometry, multiparameter ............................. Instrument settings .................................................. Formaldehyde, 37% ....................................................... Fugene 6.........................................................................

163 249 163 161 165 325 128

G GeneChip Human Promoter 1, 0 Array ....................... 361 GeneChip hybridization, wash and stain kit................. 345 GeneChip sample clean up module............................... 345 GeneChip WT double-Stranded DNA Terminal Labling kit ................................................... 345 Genomatix ChipMiner.................................................. 377 Genomic DNA, digestion ............................................. 390 Giemsa dye ...................................................................... 86 Glass beads, acid washed ................................................. 96 Green Fluorescent Protein ............................................ 126 Photoactivatable variant........................................... 126

H HEK293T cells.............................................................. 281 Hirt extraction protocol................................................. 210 Histone Acetylation............................................................... 263 Chaperone................................................................ 263 Core ......................................................................... 127 Deacetylase 7 complex ............................................. 282 Deacetylase enzymatic assay .................................... 287 Deacetylase (HDAC) .............................................. 279 Deubiquitination.............................. 295, 297, 298, 303 H2AX .............................................................. 126, 397 H2AX phosphorylation ........................................... 161 H2B ................................................................. 127, 135 H3 ............................................................................ 311 H3 phosphorylation......................................... 311, 312 Linker histone removal............................................ 116

CHROMATIN PROTOCOLS

414 Index

Histone (continued) Octamer ................................................................... 251 Phosphorylation ....................................................... 312 Purification by chromatography .............................. 255 Purified ...................................................................... 18 Sumoylation ............................................................. 295 Ubiquitination.......................................... 295, 296, 298 HMMTiling .................................................................. 371 Hoechst 33342....................................................... 127, 314 Hybridization Buffer, ChIP-on-chip assay..................................... 346 Closed circular DNA............................................... 100 Comparative Genomic............................................. 177 in situ ................................................... 68, 73, 183, 194 Telomerase assay...................................................... 390 Whole mount............................................................. 68 Hydroxy radicals ............................................................ 119

I IGEPAL CA-630 ......................................................... 345 Image, differential interference contrast (DIC) ............ 130 Immunofluorescence............................................ 32, 70, 76 Immunohistochemical analysis ...................................... 318 Immunolabeling..................................................... 128, 131 Immunoprecipitation and co-immunoprecipitation ...................... 399, 402 Immunoprecipitation reaction, Chromatin ........... 328, 348 Immunostaining..................... 257, 314, 318, 319, 400, 404 Paraffin embedded sections ..................................... 319 Preparation of tissue ................................................ 318 Irradiation, gamma ........................................................ 169 Irradiation, UV .............................................................. 147 Isolation Genomic DNA ........................................................ 390 Nuclei....................................................................... 114

J JmjC domain proteins.................................................... 250

K Kanamycin ..................................................................... 207 Kinase assay, ATM................................................ 400, 403 Klenow DNA polymerase................................................ 21

M Magnetic beads .............................................................. 361 Mass spectrometry ......................................................... 241 Medium Mounting ............................................................. 30, 34 Melphalan ...................................................................... 172 Metabolic labeling of cells ............................................. 244 Metaphase chromosome preparation............................. 184 Methotrexate.................................................................... 29 Methylation Arginine ................................................................... 235 Genomic .................................................................. 217 in vitro.............................................................. 239, 255 Methyltransferase Accessibility protocol for individual templates (MAP-IT)................................................ 44, 45 Accessibility protocol (MAP) .............................. 42, 44 Cytosine-5.................................................................. 42 DNA .......................................................................... 41 Histone .................................................................... 236 Mapping..................................................................... 50 Mapping of DNA-Protein Interactions .................... 56 Protein arginine ....................................................... 235 Microarray...................................................................... 342 Micrococcal nuclease ........................ 3, 8, 15, 20, 115, 267, 273, 328, 360 Microscopy Bright-field .............................................................. 316 Epifluorescence ........................................ 171, 195, 315 Laser scanning confocal ................................... 128, 130 Model-based analysis of tiling arrays (MAT) ............... 372 Monitoring DNA breaks in chromatin in vivo............. 125 Materials .................................................................. 128 Methods................................................................... 129 Notes........................................................................ 136 Mononucleosomes ......................................................... 109 Purification of .......................................................... 301 Reconstitution.................................................. 117, 302 uH2A containing ..................................................... 301 Mounting sections on slides .......................................... 318 Mouse embryo for in situ hybridization .......................... 71 Preparation of ............................................................ 71 Tissue sections ........................................................... 73

N L Lac operator ..................................................................... 28 Laser microirradiation ................................................... 129 Linking number ..................................................... 101, 102 LSD1 ............................................................................. 249 Lysate, ChIP assay......................................................... 327 Lysate, telomerase activity ............................................. 388 Lysis Buffer, ChIP assays ...................................... 326, 346

Neutral filter elution of DNA ............................... 401, 405 Nick translation ..................................................... 181, 191 NPM1 ............................................................................ 263 Nuclear extract, HeLa ........................................... 265, 268 Nucleophosmin.............................................................. 275 Nucleosome........................................................................ 1 Assembly .................................................................. 111 Mapping........................................................... 118, 119

CHROMATIN PROTOCOLS

Index 415

Positioning ............................................................... 111 Positioning sequence ....................................... 113, 121 Purification ...................................................... 251, 254 Repeat length............................................................... 9 Separation of ................................................................ 4 Spacing factors ............................................................. 1

O Okadaic acid .............................................................. 84, 88 Oligonucleosomes.......................................................... 306

P Paraformaldehyde .................................................. 314, 320 Partek Genomics Suite .................................................. 376 PEPI-1 plasmid ............................................................. 210 Peroxidase activity, endogenous .................................... 319 PG5ML array................................................................. 264 PHO5 promoter ............................................................... 44 Phosphatase, alkaline ....................................................... 69 Phosphorylation, H2AX......................... 29, 126, 131–135, 138, 161, 397 Photoactivation .............................................................. 127 Photobleaching .............................................................. 126 Polycomb proteins ......................................................... 295 Polymerase chain reaction Bisulfite Sequencing ................................ 217, 220, 230 Degenerate oligonucleotide primed................. 181, 189 Ligation mediated.................................................... 356 Methylation specific................................. 217, 220, 229 Real-time quantitative ............................................. 155 Whole Genome Amplification................................ 359 Polymerase II, RNA ...................................................... 141 Polymerase, Taq DNA ..................... 52, 58, 112, 113, 146, 181, 189, 190, 383, 388, 389 PRCLac vector ................................................................ 30 Preblocking Staph A beads............................................ 331 Preclearing of ChIP lysates.................................... 331, 355 Preparation of Chromatin assembly extracts Materials ...................................................................... 3 Methods....................................................................... 5 Notes............................................................................ 7 Preparation of donor chromatin .................................... 114 Primer design, PCR............................................... 222, 227 Propidium iodide ........................................................... 164 Protein A....................................................................... 154, 330 G .............................................................................. 330 HMGN...................................................................... 67 Nucleosomal binding ................................................. 67 Proteinase K............................................................. 79, 170 Purification of core histones .................................. 265, 269 Purification of histone deacetylases ............... 280, 284, 285 Purification of hyperacetylated core histones ................ 269

Q QIAPrep miniprep Kit .................................................. 211

R Radiation, ionizing ........................................ 162, 165, 396 Radiation sensitivity....................................................... 397 Reaction, histone deubiquitination ............................... 303 Repair analysis By ChIP assay .......................................................... 153 By indirect end-labeling .......................................... 151 Repair, global genome ................................................... 142 Repair, nucleotide excision ............................................ 141 Repair, transcription coupled......................................... 141 Replication origins ...................................................................... 203 in vitro...................................................................... 203 RNA DIG labeled ............................................................... 70

S Saccharomyces cerevisiae............................................... 45, 94 See also yeast Saline, Tris buffered ................................................ 69, 313 Phosphate buffered ......................................... 162, 313, 344, 401 See also TBS SDS-PAGE........................................................... 399, 403 Sections, paraffin embedded.......................................... 319 Immunohistochemical analysis of............................ 319 Sephadex G-50 columns................................................ 156 Sepharose, Protein G fast-flow ..................................... 345 Sf 9 cells ......................................................................... 285 Simian virus ............................................................. 40, 203 Sirtuins, (SIRT1–7)....................................................... 280 Sodium azide ................................................................. 320 Sodium bisulphite conversion........................................ 219 Southern transfer ........................................................... 391 SSC, 20X ......................................................................... 96 Staph A cells .................................................................. 330 Steroid receptor coactivator ........................................... 236 Substrate, histone deacetylase................................ 282, 286 Sucrose gradient............................................................. 254

T TBS.......................................................................... 69, 313 See also Saline, TRIS buffered TBS-Tween 20 (TTBS) .................................................. 69 Telomerase Activity..................................................................... 383 Associated proteins .................................................. 383 Length.............................................................. 383, 390 RNA component (TERC) ...................................... 383 TileMap ......................................................................... 373

CHROMATIN PROTOCOLS

416 Index

Tilescope ........................................................................ 375 Tiling analysis software, Affymetrix.............................. 369 Tiling Arrays, Affymetrix .............................................. 367 TiMAT .......................................................................... 374 Topoisomerase I, Drosophila ................................ 264, 267 Catalytic domain...................................................... 264 Topoisomerase II ........................................................... 169 Topotecan ...................................................................... 165 Transcription coupled repair, method to study ............. 141 Materials .................................................................. 147 Methods................................................................... 149 Notes.............................................................................. 155 Transcription, in vitro............................................ 267, 274 Transfection ............................................................. 31, 128 Transformation Bacteria .............................................................. 48, 211 Yeast..................................................................... 48, 53 TRAP Assay .......................................................... 384, 389 Trichloroacetic acid (TCA) ........................................... 251 Triton X-100, 10% ........................................................ 313

U Ubiquitination................................................................ 295 Ultra-violet irradiation................................................... 147 U-test, Mann-Whitney ................................................. 370 UV Transilluminator ....................................................... 47

V Vectastain Elite Kit........................................................ 313 Visualization of HMGN Nucleosomal proteins ............. 67 Materials .................................................................... 68 Methods..................................................................... 70 Notes.......................................................................... 77

W Wash buffers, ChIP assay...................................... 336, 346 Wash, Church Gilbert..................................................... 97 WebLogo application .................................................... 379 Western blot analysis ............................................. 257, 399 Whatman 3 mm paper................................................... 120 Whole mount in situ hybridization ........................... 68, 70 Wilcoxon signed-rank test............................................. 370 Wizard DNA Clean-up System .................................... 219

X X-chromosome inactivation .......................................... 249

Y Yeast DNA topology ........................................................... 93 MTase expressing ...................................................... 52 T85 strain................................................................. 298 See also Saccharomyces cerevisiae

Chromatin Protocols (Methods in Molecular Biology)

Chromatin Protocols, 2nd Edition (Methods in Molecular Biology Vol 523)

Chromatin Protocols (Methods in Molecular Biology Vol 119)

Chromatin Remodeling (Methods in Molecular Biology, v833)

Yeast Protocols (Methods in Molecular Biology) (Methods in Molecular Biology)

Molecular Toxicology Protocols (Methods in Molecular Biology)

Molecular Imaging: Methods and Protocols (Methods in Molecular Biology)

Molecular Chaperones: Methods and Protocols (Methods in Molecular Biology, v787)

Molecular Chaperones: Methods and Protocols (Methods in Molecular Biology 787)

Molecular Profiling: Methods and Protocols (Methods in Molecular Biology, 823)

Arabidopsis Protocols (Methods in Molecular Biology (Cloth))

Photosynthesis Research Protocols (Methods in Molecular Biology)

Genomics Protocols (Methods in Molecular Biology)

DNA Sequencing Protocols (Methods in Molecular Biology)

Epitope Mapping Protocols (Methods in Molecular Biology)

B Cell Protocols (Methods in Molecular Biology)

Retinoid Protocols (Methods in Molecular Biology)

Dictyostelium discoideum Protocols (Methods in Molecular Biology)

Matrix Metalloproteinase Protocols (Methods in Molecular Biology)

Parasite Genomics Protocols (Methods in Molecular Biology)

PCR Cloning Protocols (Methods in Molecular Biology)

Protein Targeting Protocols (Methods in Molecular Biology)

Matrix Metalloproteinase Protocols (Methods in Molecular Biology)

Cytokine Protocols (Methods in Molecular Biology)

PCR Protocols (Methods in Molecular Biology)

Protein Phosphatase Protocols (Methods in Molecular Biology)

Transmembrane Signaling Protocols (Methods in Molecular Biology)

Pyrosequencing Protocols (Methods in Molecular Biology)

YAC Protocols (Methods in Molecular Biology)

DNA Repair Protocols (Methods in Molecular Biology)

DNA Methylation Protocols (Methods in Molecular Biology)

Chromatin Protocols (Methods in Molecular Biology)

Chromatin Protocols, 2nd Edition (Methods in Molecular Biology Vol 523)