Protein Acetylation: Methods and Protocols [1st ed.] 978-1-4939-9433-5;978-1-4939-9434-2

Table of contents :
Front Matter ....Pages i-xiii
Front Matter ....Pages 1-1
High-Resolution Mass Spectrometry to Identify and Quantify Acetylation Protein Targets (Birgit Schilling, Jesse G. Meyer, Lei Wei, Melanie Ott, Eric Verdin)....Pages 3-16
Isolation of Proteins on Nascent Chromatin and Characterization by Quantitative Mass Spectrometry (Paula A. Agudelo Garcia, Miranda Gardner, Michael A. Freitas, Mark R. Parthun)....Pages 17-27
Detection and Quantitation of Acetylated Histones on Replicating DNA Using In Situ Proximity Ligation Assay and Click-It Chemistry (Pavlo Lazarchuk, Sunetra Roy, Katharina Schlacher, Julia Sidorova)....Pages 29-45
Front Matter ....Pages 47-47
Quantification of In Vitro Protein Lysine Acetylation by Reversed Phase HPLC (Catherine W. Njeri, Onyekachi E. Ononye, Lata Balakrishnan)....Pages 49-56
Measurement and Analysis of Lysine Acetylation by KAT Complexes In Vitro and In Vivo (Anahita Lashgari, Jean-Philippe Lambert, Jacques Côté)....Pages 57-77
Site-Specific Lysine Acetylation Stoichiometry Across Subcellular Compartments (Anastasia J. Lindahl, Alexis J. Lawton, Josue Baeza, James A. Dowell, John M. Denu)....Pages 79-106
Lysine Acetylation of Proteins and Its Characterization in Human Systems (David K. Orren, Amrita Machwe)....Pages 107-130
Front Matter ....Pages 131-131
Molecular and Cellular Characterization of SIRT1 Allosteric Activators (Michael B. Schultz, Conrad Rinaldi, Yuancheng Lu, João A. Amorim, David A. Sinclair)....Pages 133-149
Assessment of SIRT2 Inhibitors in Mouse Models of Cancer (Yashira L. Negrón Abril, Irma Fernández, Robert S. Weiss)....Pages 151-171
Front Matter ....Pages 173-173
Using Yeast as a Model Organism to Study the Functional Roles of Histone Acetylation in DNA Excision Repair (Amelia J. Hodges, Steven A. Roberts, John J. Wyrick)....Pages 175-190
Biochemical and Cellular Assays to Assess the Effects of Acetylation on Base Excision Repair Enzymes (Shrabasti Roychoudhury, Suravi Pramanik, Hannah L. Harris, Kishor K. Bhakat)....Pages 191-206
Analysis of DNA Processing Enzyme FEN1 and Its Regulation by Protein Lysine Acetylation (Onyekachi E. Ononye, Catherine W. Njeri, Lata Balakrishnan)....Pages 207-224
Examining the Role of HDACs in DNA Double-Strand Break Repair in Neurons (Ping-Chieh Pao, Jay Penney, Li-Huei Tsai)....Pages 225-234
Front Matter ....Pages 235-235
Experimental Approaches to Investigate the Role of Helicase Acetylation in Regulating R-Loop Stability (Chenlin Song, Ingrid Grummt)....Pages 237-253
Rapid Detection of p53 Acetylation Status in Response to Cellular Stress Signaling (Marina Farkas, Steven B. McMahon)....Pages 255-262
Front Matter ....Pages 263-263
Isolation and Quantification Brain Region-Specific and Cell Subtype-Specific Histone (De)Acetylation in Cognitive Neuroepigenetics (Craig Myrum, Peter R. Rapp)....Pages 265-277
Functional Analysis of HDACs in Tumorigenesis (Melissa Hadley, Satish Noonepalle, Debarati Banik, Alejandro Villagra)....Pages 279-307
Back Matter ....Pages 309-310

Citation preview

Methods in Molecular Biology 1983

Robert M. Brosh, Jr. Editor

Protein Acetylation Methods and Protocols

Methods

in

M o l e c u l a r B i o lo g y

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

For further volumes: http://www.springer.com/series/7651

Protein Acetylation Methods and Protocols

Edited by

Robert M. Brosh, Jr. NIH Biomedical Research Center, National Institute on Aging, National Institutes of Health, Baltimore, MD, USA

Editor Robert M. Brosh, Jr. NIH Biomedical Research Center National Institute on Aging National Institutes of Health Baltimore, MD, USA

ISSN 1064-3745     ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-4939-9433-5    ISBN 978-1-4939-9434-2 (eBook) https://doi.org/10.1007/978-1-4939-9434-2 © Springer Science+Business Media, LLC, part of Springer Nature 2019 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Cover caption: Detection of acetylated histones on newly replicated DNA using Click-It reaction to attach biotin molecules to EdU bases in DNA, and Proximity Ligation Assay to generate fluorescent signal at sites where EdU bases and acetylated histones are close to each other. This Humana Press imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 233 Spring Street, New York, NY 10013, U.S.A.

Preface Over the past several decades, modulation of protein function by posttranslational modifications has attracted great interest in multiple areas of biology. The covalent and enzymatic modification of proteins post their biosynthesis provides an often reversible and acute means of regulating vital processes in cells. The functional roles of proteins can be modulated by various covalent modifications including phosphorylation, sumoylation, ubiquitination, glycosylation, and acetylation. Such protein modifications affect various aspects including protein folding, subcellular targeting, interactions with other molecules, catalytic function, and signaling capacity. This book is dedicated to protein acetylation, which has emerged as a highly provocative means of functional regulation. The leading experts in the field of protein acetylation were recruited for this book to contribute experimental strategies and useful laboratory protocols that represent a comprehensive collection of great utility and significance. I anticipate that this Methods in Molecular Biology book on Protein Acetylation will be widely read and applied in almost every biological setting of experimentation, as it is increasingly evident that acetylation of chromatin-associated proteins and nucleic acid-interacting and nucleic acid-modifying proteins plays extremely important roles in not only genome metabolism but more broadly cellular homeostasis, tissue integrity and function, and organismal fitness. This book on protein acetylation describes both fundamental and more elaborate protocols to measure and assay protein acetylation and its consequences in biological systems. The book begins with three chapters on molecular and cellular techniques to detect, quantify, and isolate acetylated protein targets. Schilling et  al. describe a high-resolution and high-throughput mass spectrometry approach using minimal protein lysate and an optimized affinity enrichment strategy with a label-free quantification workflow to assess protein acetylation (Chapter 1). This is followed by a contribution from the Parthun lab that describes experimental procedures to evaluate acetylation of histones during chromatin assembly, a highly relevant topic in the burgeoning field of epigenetics (Chapter 2). The combinatorial approach involves state-of-the-art iPOND, quantitative mass spectrometry, and SILAC methodologies to characterize acetylation in nascent chromatin. This first section of the book is wrapped up with an exemplary chapter from the Sidorova lab describing experimental procedures to detect and quantify acetylated histones that occur during DNA replication using an in situ Click Chemistry to label DNA and Proximity Ligation Assay to specifically visualize the labeled DNA with a modified histone of choice that is recognized by a modification-specific antibody (Chapter 3). The approach is valuable to measure not only histone modifications during active replication but also acetylation when the replication fork stalls due to damage or pharmacological nucleotide depletion. The focus of this unique collection of experimental procedures dealing with protein acetylation is then targeted in four chapters on measurement and analysis of lysine acetylation and functional consequences. Lysine acetylation is enacted by a group of enzymes designated lysine acetyltransferases (KATs). This section begins with a chapter from the Balakrishnan laboratory which describes a reverse-phase HPLC-based strategy that assesses substrate consumption and product formation at the same time (Chapter 4). Its value

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resides in the strong reproducibility and application toward kinetic analysis of KATs. Recognizing the importance of studying native KAT complexes, the Côté lab offers a useful protocol to purify KAT complexes from human cells based on genome editing and tandem affinity purification (TAP) (Chapter 5). They also describe experimental procedures to study the isolated KAT complexes using a lysine acetyltransferase assay and detection of acetylated lysines by small-scale affinity purification or anti-acetyl lysine antibody immunopurification. Finally, they present a general procedure to make homogeneous and site-­ specific acetylated recombinant protein in bacteria so that functional consequences of acetylation can be studied with a suitable quantity of purified protein. To address the difficulty of determining lysine acetylation stoichiometry across subcellular compartments in eukaryotic cells, the Denu lab provides a strategy of subcellular fractionation with offline prefractionation to determine acetylation stoichiometry using data-independent mass spectrometry (Chapter 6). This approach is highly valuable for investigation of acetylation in various organelles including mitochondria where the posttranslational modification may occur in low stoichiometry. This chapter is followed by one from the Orren lab that describes established and novel experimental approaches and methodologies to detect lysine acetylation or deacetylation of specific target proteins or groups of proteins and how these events are regulated (Chapter 7). A unique class of deacetylation enzymes are the sirtuins, which are discussed in two chapters dedicated to protocols for detection and characterization of sirtuin targets. The Sinclair lab is especially interested in sirtuins as they represent a class of enzymes that have been implicated in healthy aging and longevity. In their chapter, Schultz et  al. describe experimental approaches to identify and characterize sirtuin-activating compounds (STACs) (Chapter 8). This includes the purification of the prominent lysine deacetylase SIRT1, in vitro assays with recombinant SIRT1 and allosteric activators, and mitochondrial assays for SIRT1 activators in cells. Following this chapter, the Weiss lab presents a series of procedures to investigate small molecule inhibitors of another sirtuin known as SIRT2 implicated in tumorigenesis using genetically engineered and xenograft mouse models of cancer (Chapter 9). Experimental procedures to detect and measure the consequences of acetylation for DNA repair and more broadly DNA processing enzymes and processes are presented in two modules. Beginning with protein targets that influence DNA repair, the Wyrick lab discusses techniques exploiting the model and genetically tractable unicellular organism yeast to investigate the functional consequences of histone acetylation on DNA nucleotide excision repair (NER) (Chapter 10). Procedures to measure the repair of cyclobutane pyrimidine dimers (CPDs) in bulk chromatin as well as individual chromatin loci in yeast that have been manipulated for their histone acetyltransferase (HAT) activity are described. Next, the Bhakat group presents experimental assays to assess the effect of acetylation on a key player in the base excision repair (BER) pathway, i.e., abasic endonuclease 1 (APE1) (Chapter 11). This narrative includes in vitro assays measuring APE1 DNA substrate binding and repair (catalytic) activity, as well as DNA damage repair in  vivo as measured by comet assay and cell survival after genotoxic exposure, and subcellular localization of APE1 by immunostaining. Ononye et  al. present experimental protocols to analyze the effects incurred by acetylation of the structure-specific nuclease known as flap endonuclease 1 (FEN1), a key processing enzyme implicated in BER and lagging strand maturation during DNA replication (Chapter 12). Following these chapters, the Tsai lab describes experimental assays to assess the effect of HDAC inhibition on DNA double-strand break (DSB) repair in mouse primary cortical neurons (Chapter 13).

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Reaching beyond specific DNA repair proteins or pathways, the influence of protein acetylation on processes that affect genomic stability and the cellular stress response is described in two subsequent chapters. Song and Grummt present experimental protocols to assay acetylation of a target DNA helicase known as DDX21 that is implicated in the resolution of three-stranded nucleic acid structures designated R-loops composed of both DNA and RNA arising from transcription; such structures can be problematic when encountered by the replication fork (Chapter 14). Assays to measure the functional consequences of DDX21 acetylation on its helicase activity are presented. Aside from catalytic enzymes like helicases or nucleosomal proteins such as histones, certain DNA-interacting factors such as the tumor suppressor and cell cycle regulator p53 can be acetylated as well. Farkas and McMahon present a procedure for the rapid detection of p53 acetylation status that becomes modulated by cellular stress (Chapter 15). Having covered various facets of acetylation and its modulation of biological processes at the molecular and cellular levels, as well as studies in model organisms including yeast and mouse, the final two chapters of the collection delve into two areas of disease and aging research of great interest, namely, neuroepigenetics and tumorigenesis. Myrum and Rapp present techniques to isolate and quantify brain-specific and cell subtype-specific acetylated histone proteins which play a role in the regulation of gene expression that profoundly influence long-term memory (Chapter 16). Hadley et  al. describe functional tests of HDACs implicated in tumorigenesis which may be applied to assess the efficacy of HDAC inhibitors (Chapter 17). These final chapters emphasize the translational value of protein acetylation targets and their modulation via biological processes and pharmacological intervention. Altogether, the compiled 17 chapters in this book provide a comprehensive set of experimental techniques and useful strategies to examine the molecular, cellular, tissue, and organismal consequences of protein acetylation. I firmly believe that this unique collection provides an extremely valuable resource to novices and experts alike who find that their research takes them in directions to characterize how the regulatory switches of protein acetylation events are manifested in significant ways and by diverse mechanisms in biological systems. I wish to thank all the authors for their outstanding chapters. Their efforts are very much appreciated, and it was a great pleasure to work with them in producing this book. I also acknowledge John Walker, the series editor, for his helpful advice and insight as well as David Casey, the editor of Springer Protocols, for his assistance and guidance. Finally, I wish to thank my teachers and mentors throughout my academic and scientific career who have taught me the mechanics and usefulness of the scientific method, the practical value of keeping a good lab notebook, and the benefits of creative thinking. While the list extends beyond those mentioned here, I especially thank my grade school teacher Mr. Gerry Robinson (St. Vincent de Paul School (Wheeling)), high school teacher Mr. Carl Hocke (Central Catholic High School (Wheeling)), undergraduate professors Drs. Milton Smith and Robert Paysen (Bethany College), and graduate advisor Dr. Steven Matson (University of North Carolina at Chapel Hill). Baltimore, MD, USA

Robert M. Brosh, Jr.

Contents Preface. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .     v Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .   xi

Part I Detection, Quantification, and Isolation of Acetylated Protein Targets   1 High-Resolution Mass Spectrometry to Identify and Quantify Acetylation Protein Targets . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .   3 Birgit Schilling, Jesse G. Meyer, Lei Wei, Melanie Ott, and Eric Verdin   2 Isolation of Proteins on Nascent Chromatin and Characterization by Quantitative Mass Spectrometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .  17 Paula A. Agudelo Garcia, Miranda Gardner, Michael A. Freitas, and Mark R. Parthun   3 Detection and Quantitation of Acetylated Histones on Replicating DNA Using In Situ Proximity Ligation Assay and Click-It Chemistry. . . . . . . .  29 Pavlo Lazarchuk, Sunetra Roy, Katharina Schlacher, and Julia Sidorova

Part II Lysine Acetylation   4 Quantification of In Vitro Protein Lysine Acetylation by Reversed Phase HPLC. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Catherine W. Njeri, Onyekachi E. Ononye, and Lata Balakrishnan   5 Measurement and Analysis of Lysine Acetylation by KAT Complexes In Vitro and In Vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anahita Lashgari, Jean-Philippe Lambert, and Jacques Côté   6 Site-Specific Lysine Acetylation Stoichiometry Across Subcellular Compartments. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anastasia J. Lindahl, Alexis J. Lawton, Josue Baeza, James A. Dowell, and John M. Denu   7 Lysine Acetylation of Proteins and Its Characterization in Human Systems. . . . David K. Orren and Amrita Machwe

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Part III Sirtuin Targets of Acetylation   8 Molecular and Cellular Characterization of SIRT1 Allosteric Activators . . . . . . 133 Michael B. Schultz, Conrad Rinaldi, Yuancheng Lu, João A. Amorim, and David A. Sinclair   9 Assessment of SIRT2 Inhibitors in Mouse Models of Cancer . . . . . . . . . . . . . . 151 Yashira L. Negrón Abril, Irma Fernández, and Robert S. Weiss

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Part IV Modulation of DNA Repair by Acetylation 10 Using Yeast as a Model Organism to Study the Functional Roles of Histone Acetylation in DNA Excision Repair. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Amelia J. Hodges, Steven A. Roberts, and John J. Wyrick 11 Biochemical and Cellular Assays to Assess the Effects of Acetylation on Base Excision Repair Enzymes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Shrabasti Roychoudhury, Suravi Pramanik, Hannah L. Harris, and Kishor K. Bhakat 12 Analysis of DNA Processing Enzyme FEN1 and Its Regulation by Protein Lysine Acetylation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Onyekachi E. Ononye, Catherine W. Njeri, and Lata Balakrishnan 13 Examining the Role of HDACs in DNA Double-Strand Break Repair in Neurons. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ping-Chieh Pao, Jay Penney, and Li-Huei Tsai

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Part V DNA Interactive Proteins and Processing Enzymes Affected by Acetylation 14 Experimental Approaches to Investigate the Role of Helicase Acetylation in Regulating R-Loop Stability. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 237 Chenlin Song and Ingrid Grummt 15 Rapid Detection of p53 Acetylation Status in Response to Cellular Stress Signaling. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 255 Marina Farkas and Steven B. McMahon

Part VI Influence of Acetylation Processes on Neuroepigenetics and Tumorigenesis 16 Isolation and Quantification Brain Region-Specific and Cell Subtype-Specific Histone (De)Acetylation in Cognitive Neuroepigenetics. . . . . . . . . . . . . . . . . . 265 Craig Myrum and Peter R. Rapp 17 Functional Analysis of HDACs in Tumorigenesis. . . . . . . . . . . . . . . . . . . . . . . 279 Melissa Hadley, Satish Noonepalle, Debarati Banik, and Alejandro Villagra Index. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 309

Contributors Paula A. Agudelo Garcia  •  Department of Biological Chemistry and Pharmacology, The Ohio State University, Columbus, OH, USA João A. Amorim  •  Department of Genetics, Blavatnik Institute, Paul F. Glenn Center for the Biology of Aging, Harvard Medical School, Boston, MA, USA; CNC—Center for Neuroscience and Cell Biology, University of Coimbra, Coimbra, Portugal; IIIUC— Institute of Interdisciplinary Research, University of Coimbra, Coimbra, Portugal Josue Baeza  •  Wisconsin Institute of Discovery, University of Wisconsin-Madison, Madison, WI, USA; Biomolecular Chemistry Department, School of Medicine and Public Health, University of Wisconsin-Madison, Madison, WI, USA; Epigenetics Institute, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA, USA; Department of Biochemistry and Biophysics, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA, USA Lata Balakrishnan  •  Department of Biology, School of Science, Indiana University Purdue University Indianapolis, Indianapolis, IN, USA Debarati Banik  •  The George Washington University Cancer Center, N.W. George Washington University, Washington, DC, USA Kishor K. Bhakat  •  Department of Genetics, Cell Biology and Anatomy, University of Nebraska Medical Center, Omaha, NE, USA; Fred and Pamela Buffet Cancer Center, University of Nebraska Medical Center, Omaha, NE, USA Jacques Côté  •  St. Patrick Research Group in Basic Oncology, Quebec City, QC, Canada; Laval University Cancer Research Center, Quebec City, QC, Canada; Centre de Recherche du Centre Hospitalier Universitaire (CHU) de Québec-Université Laval, Quebec City, QC, Canada John M. Denu  •  Wisconsin Institute of Discovery, University of Wisconsin-Madison, Madison, WI, USA; Biomolecular Chemistry Department, School of Medicine and Public Health, University of Wisconsin-Madison, Madison, WI, USA; Morgridge Institute for Research, University of Wisconsin-Madison, Madison, WI, USA James A. Dowell  •  Wisconsin Institute of Discovery, University of Wisconsin-Madison, Madison, WI, USA Marina Farkas  •  Department of Biochemistry and Molecular Biology, Sidney Kimmel College of Medicine, Thomas Jefferson University, Philadelphia, PA, USA Irma Fernández  •  Department of Biomedical Sciences, Cornell University, Ithaca, NY, USA Michael A. Freitas  •  Department of Cancer Biology and Genetics, The Ohio State University, Columbus, OH, USA Miranda Gardner  •  Department of Cancer Biology and Genetics, The Ohio State University, Columbus, OH, USA Ingrid Grummt  •  Division of Molecular Biology of the Cell II, German Cancer Research Center (DKFZ), DKFZ-ZMBH Alliance, Heidelberg, Germany Melissa Hadley  •  The George Washington University Cancer Center, N.W. George Washington University, Washington, DC, USA

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Hannah L. Harris  •  Department of Genetics, Cell Biology and Anatomy, University of Nebraska Medical Center, Omaha, NE, USA Amelia J. Hodges  •  School of Molecular Biosciences, Washington State University, Pullman, WA, USA Jean-Philippe Lambert  •  Laval University Cancer Research Center, Quebec City, QC, Canada; Centre de Recherche du Centre Hospitalier Universitaire (CHU) de Québec-­ Université Laval, Quebec City, QC, Canada; Département de Médecine Moléculaire, Université Laval, Quebec City, QC, Canada Anahita Lashgari  •  St. Patrick Research Group in Basic Oncology, Quebec City, QC, Canada; Laval University Cancer Research Center, Quebec City, QC, Canada; Centre de Recherche du Centre Hospitalier Universitaire (CHU) de Québec-Université Laval, Quebec City, QC, Canada Alexis J. Lawton  •  Wisconsin Institute of Discovery, University of Wisconsin-Madison, Madison, WI, USA; Biomolecular Chemistry Department, School of Medicine and Public Health, University of Wisconsin-Madison, Madison, WI, USA Pavlo Lazarchuk  •  Department of Pathology, University of Washington, Seattle, WA, USA Anastasia J. Lindahl  •  Wisconsin Institute of Discovery, University of Wisconsin-Madison, Madison, WI, USA; Biomolecular Chemistry Department, School of Medicine and Public Health, University of Wisconsin-Madison, Madison, WI, USA Yuancheng Lu  •  Department of Genetics, Blavatnik Institute, Paul F. Glenn Center for the Biology of Aging, Harvard Medical School, Boston, MA, USA Amrita Machwe  •  Department of Toxicology and Cancer Biology and Markey Cancer Center, University of Kentucky College of Medicine, Lexington, KY, USA Steven B. McMahon  •  Department of Biochemistry and Molecular Biology, Sidney Kimmel College of Medicine, Thomas Jefferson University, Philadelphia, PA, USA Jesse G. Meyer  •  Buck Institute for Research on Aging, Novato, CA, USA Craig Myrum  •  Laboratory of Behavioral Neuroscience, Neurocognitive Aging Section, National Institute on Aging, National Institutes of Health, Baltimore, MD, USA Yashira L. Negrón Abril  •  Department of Biomedical Sciences, Cornell University, Ithaca, NY, USA Catherine W. Njeri  •  Department of Biology, School of Science, Indiana University Purdue University Indianapolis, Indianapolis, IN, USA Satish Noonepalle  •  The George Washington University Cancer Center, N.W. George Washington University, Washington, DC, USA Onyekachi E. Ononye  •  Department of Biology, School of Science, Indiana University Purdue University Indianapolis, Indianapolis, IN, USA David K. Orren  •  Department of Toxicology and Cancer Biology and Markey Cancer Center, University of Kentucky College of Medicine, Lexington, KY, USA Melanie Ott  •  Gladstone Institutes, University of California, San Francisco, San Francisco, CA, USA Ping-Chieh Pao  •  Picower Institute for Learning and Memory, Massachusetts Institute of Technology, Cambridge, MA, USA; Department of Brain and Cognitive Sciences, Massachusetts Institute of Technology, Cambridge, MA, USA Mark R. Parthun  •  Department of Biological Chemistry and Pharmacology, The Ohio State University, Columbus, OH, USA Jay Penney  •  Picower Institute for Learning and Memory, Massachusetts Institute of Technology, Cambridge, MA, USA; Department of Brain and Cognitive Sciences, Massachusetts Institute of Technology, Cambridge, MA, USA

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Suravi Pramanik  •  Department of Genetics, Cell Biology and Anatomy, University of Nebraska Medical Center, Omaha, NE, USA Peter R. Rapp  •  Laboratory of Behavioral Neuroscience, Neurocognitive Aging Section, National Institute on Aging, National Institutes of Health, Baltimore, MD, USA Conrad Rinaldi  •  Department of Genetics, Blavatnik Institute, Paul F. Glenn Center for the Biology of Aging, Harvard Medical School, Boston, MA, USA Steven A. Roberts  •  School of Molecular Biosciences, Washington State University, Pullman, WA, USA; Center of Reproductive Biology, Washington State University, Pullman, WA, USA Sunetra Roy  •  Department of Cancer Biology, University of Texas MD Anderson Cancer Center, Houston, TX, USA Shrabasti Roychoudhury  •  Department of Genetics, Cell Biology and Anatomy, University of Nebraska Medical Center, Omaha, NE, USA Birgit Schilling  •  Buck Institute for Research on Aging, Novato, CA, USA Katharina Schlacher  •  Department of Cancer Biology, University of Texas MD Anderson Cancer Center, Houston, TX, USA Michael B. Schultz  •  Department of Genetics, Blavatnik Institute, Paul F. Glenn Center for the Biology of Aging, Harvard Medical School, Boston, MA, USA Julia Sidorova  •  Department of Pathology, University of Washington, Seattle, WA, USA David A. Sinclair  •  Department of Genetics, Blavatnik Institute, Paul F. Glenn Center for the Biology of Aging, Harvard Medical School, Boston, MA, USA; Department of Pharmacology, School of Medical Sciences, The University of New South Wales, Sydney, NSW, Australia Chenlin Song  •  Division of Molecular Biology of the Cell II, German Cancer Research Center (DKFZ), DKFZ-ZMBH Alliance, Heidelberg, Germany Li-Huei Tsai  •  Picower Institute for Learning and Memory, Massachusetts Institute of Technology, Cambridge, MA, USA; Department of Brain and Cognitive Sciences, Massachusetts Institute of Technology, Cambridge, MA, USA Eric Verdin  •  Buck Institute for Research on Aging, Novato, CA, USA Alejandro Villagra  •  The George Washington University Cancer Center, N.W. George Washington University, Washington, DC, USA Lei Wei  •  Buck Institute for Research on Aging, Novato, CA, USA Robert S. Weiss  •  Department of Biomedical Sciences, Cornell University, Ithaca, NY, USA John J. Wyrick  •  School of Molecular Biosciences, Washington State University, Pullman, WA, USA; Center of Reproductive Biology, Washington State University, Pullman, WA, USA

Part I Detection, Quantification, and Isolation of Acetylated Protein Targets

Chapter 1 High-Resolution Mass Spectrometry to Identify and Quantify Acetylation Protein Targets Birgit Schilling, Jesse G. Meyer, Lei Wei, Melanie Ott, and Eric Verdin Abstract The dynamic nature of protein posttranslational modification (PTM) allows cells to rapidly respond to changes in their environment, such as nutrition, stress, or signaling. Lysine residues are targets for several types of modifications, including methylation, ubiquitination, and various acylation groups, especially acetylation. Currently, one of the best methods for identification and quantification of protein acetylation is immunoaffinity enrichment in combination with high-resolution mass spectrometry. As we are using a relatively novel and comprehensive mass spectrometric approach, data-independent acquisition (DIA), this protocol provides high-throughput, accurate, and reproducible label-free PTM quantification. Here we describe detailed protocols to process relatively small amounts of mouse liver tissue that integrate isolation of proteins, proteolytic digestion into peptides, immunoaffinity enrichment of acetylated peptides, identification of acetylation sites, and comprehensive quantification of relative abundance changes for thousands of identified lysine acetylation sites. Key words Acetylation, Posttranslational modifications, Mass spectrometry, Data-independent acquisition, Quantification

1  Introduction Proteomics technology has become the best method to determine changes in relative protein abundances, but also to measure changes in protein posttranslational modifications (PTM). Many unique protein forms with different PTMs, called proteoforms, may exist in parallel, and the quantity of each proteoform is often highly dynamic. Lysine acetylation is one of the most common PTMs among cellular proteins and regulates a variety of physiological processes including enzymatic activity, protein–protein interactions, gene expression, and subcellular localization [1]. Modification of lysine residues by acetylation was first studied over 50  years ago [2, 3]. In recent years, more extensive proteomic acetylation studies revealed previously unappreciated roles for lysine acetylation in the regulation of diverse cellular pathways,

Robert M. Brosh, Jr. (ed.), Protein Acetylation: Methods and Protocols, Methods in Molecular Biology, vol. 1983, https://doi.org/10.1007/978-1-4939-9434-2_1, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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and particularly of mitochondrial proteins [4]. Furthermore, it has been demonstrated how drastically the mitochondrial acetylome can be altered, such as through deacetylation by the sirtuin SIRT3 or via nutritional changes [5, 6]. Due to relatively low levels of acetylation modifications and low lysine site occupancies, mass spectrometric identification of acetylation sites is not trivial, and antibody-based enrichment strategies have proven to be highly effective to gain deeper insights into the dynamic acetylome. However, these workflows are challenging due to multiple processing steps that must be highly quantitative and reproducible, and there is a great need for more standardized protocols. Additionally, affinity enrichment protocols often require a high amount of protein lysate input material, which can be a challenge when limited amounts of starting materials are available. For example, the original Cell Signaling Technologies “PTMScan Acetyl-Lysine Motif [Ac-K] Immunoaffinity Bead” protocol recommends the use of 10–20 mg protein lysate as starting material. To overcome these limitations and enable more routine and robust identification and quantification of protein acetylation sites, here we present a detailed protocol describing a high-throughput and quantitative proteomic approach using 1–5 mg protein lysate for acetyl peptide enrichment (also see Note 1). Our method utilizes optimized affinity enrichment in combination with a comprehensive, label-free data-independent acquisition (DIA) mass spectrometric workflow, and several open-source data processing software tools (see Fig. 1). This protocol can accommodate relatively low amounts of starting material and results in the discovery of regulated acetylation sites in proteins via an unbiased acetylomics approach, thus enabling subsequent experiments to decipher the biological significance of site-specific protein acetylation. Tissues, such as liver (or similarly, isolated mitochondria or cell lines), are lysed and soluble protein is obtained (1–20 mg), which is subsequently proteolytically digested into tryptic peptides. Acetylated peptides are immunoaffinity enriched using anti-acetyl-­lysine antibodies. Enriched peptides are then analyzed by nanoflow liquid chromatography coupled to tandem mass spectrometry (nLC-MS/MS) in data-independent acquisition (DIA) mode. Quantification and analysis is performed using open-source software tools, including Skyline.

Fig. 1 Acetyl-lysine immunoaffinity enrichment and label-free quantification workflow

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2  Materials 2.1  Tissue Lysis and Tryptic Protein Digestion

1. Mouse liver tissue (wild-type B57BL). 2. 2 mL Safe-Lock Tubes (Eppendorf). 3. Lysis buffer: 8 M Urea in 100 mM triethylammonium bicarbonate (TEAB), pH 8.5 containing 1× HALT protease inhibitor cocktail (Pierce), deacetylase inhibitors: 5 μM trichostatin A (TSA), 5  mM nicotinamide, and 75  mM sodium chloride (NaCl). 4. QIAGEN TissueLyser II and stainless steel beads, 5  mm (QIAGEN). 5. Bioruptor sonicator (Diagenode). 6. Bicinchoninic acid (BCA) protein assay (Pierce Thermo Scientific). 7. Reducing Reagent: 1  M dithiothreitol (DTT), freshly prepared in deionized water/MilliPORE (referred to as H2O). 8. Alkylation Reagent: 200  mM iodoacetamide (IAA), freshly prepared in H2O. 9. Dilution buffer: 50 mM TEAB in H2O. 10. Digestion enzyme: (Promega).

2.2  Desalting of Proteolytic/Tryptic Peptides After Digestion (Oasis/ HLB)

Modified

sequencing-grade

trypsin

1. Oasis HLB (Hydrophilic-Lipophilic-Balanced) 1  cc Vac Cartridge, 30 mg Sorbent per Cartridge, 30 μm particle size (Waters Technologies Corporation). 2. Extraction manifold, 20-port vacuum manifold (Waters Technologies Corporation). 3. HPLC MS-grade acetonitrile (ACN) and water (H2O) (Burdick and Jackson). 4. HPLC MS-grade formic acid (FA) (Sigma-Aldrich). 5. HLB Solvent A: 0.2% FA in HPLC-MS grade H2O. 6. HLB Solvent B: 80% ACN/20% of 0.2% FA in HPLC-MS grade H2O.

2.3  Anti-Acetyl Immunoaffinity Enrichment

1. PTMScan Acetyl-Lysine Motif [Ac-K] Immunoaffinity Beads (Cell Signaling Technologies). 2. PTMScan Immunoaffinity (IAP) Buffer: 50  mM MOPS, 10  mM Na3PO4, 50  mM NaCl in water at pH  7.2 (Cell Signaling Technologies). 3. 1× Phosphate-buffered saline (PBS): 0.01  M phosphate-­ buffered saline (0.0027 M KCl, 0.138 M NaCl) at pH 7.4 at 25 °C (Sigma-Aldrich). 4. Wide-bore 200 μL pipet tips (VWR International).

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2.4  Small-Scale Acetyl-Peptide Desalting Prior to MS Analysis

1. Empore Octadecyl (C18) 47 mm Extraction Disks (3 M). 2. 18-Gauge blunt-tipped needle and plunger. 3. VWR 200 μL low-binding pipet tips (VWR International). 4. MulTI SafeSeal Sorenson 0.65  mL microcentrifuge tubes (VWR International). 5. Snap Cap Low Retention 1.5 and 2 mL graduated microcentrifuge (Eppendorf) tubes (Thermo Scientific). 6. StageTip Solvent A: 0.2% FA in HPLC-MS grade H2O. 7. StageTip Solvent B: 0.2% FA in 50% HPLC-MS grade ACN in HPLC-MS H2O.

2.5  Chromatography and Mass Spectrometry: Nanoflow HPLC-MS/ MS

1. All HPLC-MS/MS buffers are “HPLC-MS grade” (all Burdick and Jackson). 2. Mobile Phase A: 2% ACN/98% water/0.1% formic acid (v/v/v). 3. Mobile Phase B: 98% ACN/2% water/0.1% formic acid (v/v/v). 4. Nanoflow liquid chromatography: Ultra Plus nano-LC 2D HPLC (Eksigent) connected to a cHiPLC system (Eksigent) with a C18 pre-column chip (200  μm × 0.4  mm ChromXP C18-CL chip, 3 μm, 120 Å, SCIEX), and an analytical C18 column chip (75 μm × 15 cm ChromXP C18-CL chip, 3 μm, 120 Å). 5. Mass Spectrometer: quadrupole time-of-flight (QqTOF): TripleTOF 6600 system (SCIEX) or other high-resolution mass spectrometry systems.

3  Methods 3.1  Tissue Lysis and Tryptic Protein Digestion

1. Harvest mouse liver and take ~50 mg tissue (wet weight) and process immediately or freeze at −80 °C. 2. Chill TissueLyser adapter sets to −20 °C or set on dry ice. 3. Add frozen tissue and one stainless steel bead to each of the labeled and chilled Safe-Lock tubes (Eppendorf) over dry ice. 4. Add 400  μL ice-cold lysis buffer containing protease and deacetylase inhibitors to each of the tubes containing tissue and bead. 5. Vortex briefly to ensure that the lysis buffer volume covers the whole tissue. If the tissue is large and is not fully submerged add more lysis buffer in increments of 50 μL until the tissue is fully covered with buffer. 6. Place tubes on chilled adapter sets and ensure balancing the tubes among the 2 adapters. Homogenize with TissueLyser II at 30 Hz two times for 3 min at 4 °C.

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7. Remove bead with tweezer. Clean tweezer with deionized water and HPLC-grade methanol and dry in order to prevent cross-contamination. 8. Spin briefly to collect all volume to the bottom of the tube. 9. Sonicate on Bioruptor sonicator (Diagenode) for 10 cycles of 30 s ON/30 s OFF at 4 °C at high power. 10. Centrifuge homogenized tissue lysate for 10  min, at 14,000 × g, at 4 °C. 11. Transfer supernatant to new 1.5 mL tubes while avoiding any lipid and fat layer above the cleared lysate and any pellet at the bottom of the tube. 12. Determine protein concentration using the BCA assay. 13. Remove an aliquot of lysate containing 5 mg of soluble protein according to BCA assay (or more input material if available, e.g., 20  mg, see Note 1), and add DTT to a final concentration of 4.5 mM to reduce disulfide bonds for 30 min at 37 °C with 3.5 × g agitation (Eppendorf ThermoMixer). 14. Cool reduced protein lysate to room temperature (RT), and add IAA to a final concentration of 10  mM to alkylate free thiols. Allow reaction to proceed at RT in the dark for 30 min. 15. Dilute reduced and alkylated proteins tenfold with 50  mM TEAB. 16. Add trypsin to initiate protein digestion (enzyme:protein ratio = 1:50, wt/wt) at 37 °C overnight with 3.5 × g agitation (Eppendorf ThermoMixer). 17. Quench the digestion by addition of FA to a final concentration of 1% FA. 18. Remove the lipids and undigested proteins by centrifugation at 1800 × g for 10 min at RT, and desalt the supernatant containing peptides (see Subheading 3.2). 3.2  Desalting Proteolytic/Tryptic Peptides Using Oasis HLB Cartridges

1. Apply vacuum to the Oasis HLB 1 cc cartridges (30 mg sorbent, max. binding capacity: 5 mg) using the vacuum extraction manifold and condition the cartridges twice with 800 μL of organic HLB Solvent B. 2. Equilibrate cartridges three times with 800  μL of aqueous HLB Solvent A. 3. Load the acidified tryptic peptides onto the cartridge (here from 5 mg protein digest). 4. Wash the bound peptides three times with 800  μL of HLB Solvent A. 5. Remove the HLB cartridges from the vacuum manifold and place into 1.5 mL microcentrifuge tubes. Elute the peptides by sequential addition of 800 μL followed by 400 μL HLB Solvent B.

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6. Mix the elution with vortexing, and then remove 2.4  μL (1/500th or 10 μg) for independent and parallel protein-level quantification (also see Note 2). 7. Concentrate/dry the eluted peptides completely using a SpeedVac (see Note 3). 3.3  Anti-Acetyl Immunoaffinity Enrichment

1. For immunoaffinity enrichment [5, 7], resuspend all the dried peptides with 1.4 mL ice-cold IAP buffer and mix by pipetting. Avoid vortexing as this may create bubbles. 2. Check that the pH of the resuspended peptide solution is between pH 7–8 by pipetting 2 μL on pH paper. If the pH deviates add more IAP buffer (in small increments of 50 μL) or adjust pH otherwise. 3. Centrifuge the resuspended peptides at 10,000 × g for 5 min at 4 °C. A small pellet may appear, but the majority of peptides will be soluble. Keep peptide solution on ice during preparation of the antibody-bead conjugate (see Note 4). 4. Prepare the PTMScan Acetyl-Lysine antibody beads for peptide affinity enrichment, add 1 mL cold 1× PBS buffer to one tube of antibody-conjugated beads, pipette up and down three times for mixing. The ideal ratio of PTMScan Acetyl-Lysine Motif antibody-conjugated beads to peptide starting material should be ¼ of a tube of antibody beads for 5 mg of peptides (see Note 1). 5. Transfer the slurry of antibody-conjugated beads to a new 1.5 mL microcentrifuge tube and centrifuge at 2000 × g for 30 s at RT to prevent beads from sticking to the side of the tube. Remove the PBS buffer by aspiration, leaving some volume to avoid disruption of the beads. 6. Wash the antibody beads with 1 mL cold 1× PBS and centrifuge at 2000 × g for 30 s at RT. Remove the majority of PBS by aspiration. 7. Repeat the PBS wash step twice more for a total of four 1 mL 1× PBS washes. 8. Resuspend the washed beads from one tube of PTMScan Acetyl-­L ysine antibody in 440 μL PBS, mix several times by pipetting with wide-bore 200 μL pipette tip, and take out four 100 μL aliquots of bead suspension into 1.5 mL microcentrifuge tubes ensuring the master mixture remained thoroughly mixed. In order to ensure consistent bead quantities in the four 100 μL aliquots, about 40 μL of beads will remain in the original tube (see Note 5). Centrifuge the aliquoted beads at 2000 × g for 30 s at RT. Visually ensure that each tube has a similar quantity of antibody-conjugated beads. Remove all 1× PBS by aspiration using a 0.2 mm gel loading flat pipet tip. 9. Transfer the resuspended peptides from step 3 directly onto the prepared PTMScan Acetyl-Lysine Motif antibody-­ conjugated beads.

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10. Incubate the peptides and antibody-conjugated bead mixture(s) at 4  °C overnight on an end-over-end rotator or gentle mixer. 11. After incubation, centrifuge the peptide/bead mixtures at 2000 × g at 4 °C for 30 s. 12. Remove the supernatant containing the unbound peptides and save for possible further applications. 13. Wash the peptide-bound beads with 1  mL cold IAP buffer, mix by inverting the tube 5 times, then centrifuge at 2000 × g, 4 °C for 30 s. Remove the IAP wash solution by aspiration, but leave a small volume to avoid bead disruption. 14. Repeat the IAP wash step once for a total of two IAP washes. 15. Wash the peptide-bound beads with 1 mL ice-cold HPLC-MS water, mix by inverting 5 times, then centrifuge at 2000 × g, 4 °C for 30 s. Remove the water wash solution by aspiration, leaving a small volume to avoid bead disruption. 16. Repeat the water wash twice for a total of three water washes. 17. After the last water wash, centrifuge once more for 30  s at 2000 × g and 4 °C to collect any remaining volume to the bottom. Aspirate the remaining water with a 0.2 mm gel loading flat pipet tip while avoiding the beads. 18. Add 55  μL 0.15% TFA in HPLC-MS water to the peptide-­ bound beads. Incubate at RT for 10 min. Tap the bottom of the tubes to mix intermittently. 19. Centrifuge the mixture for 30 s at 2000 × g, RT. Remove the eluted peptides with 0.2 mm gel loading flat pipet tip and save in a 0.65 mL microcentrifuge tube. 20. Add 45  μL 0.15% TFA in HPLC-MS water to the peptide-­ bound beads. Incubate the mixture at RT for 10  min with intermittent agitation by tapping the bottom of the tubes. 21. Centrifuge the mixture for 30 s at 2000 × g, RT. Remove the second elution by pressing a 0.2 mm gel loading flat pipet tip to the bottom of the tubes and combine with the first elution. 22. Centrifuge the eluted peptides at 12,000 × g at RT for 5 min to pellet any beads that may have carried over. Store eluted peptides on ice for immediate desalting. 3.4  Small-Scale Acetyl-Peptide Desalting Prior to LC-MS—C18 StageTips (or ZipTips)

1. Prepare the C18 StageTips for desalting as described by Rappsilber et al. [8]: assemble a set of three disks (punched out with a 18-gauge needle from an Octadecyl C18 Extraction Disk membrane) in a low-binding 200  μL pipet tip—held together in a 0.65 mL Eppendorf tube with a hole in the bottom allowing for solvent flow upon centrifugation into a 2 mL collection tube.

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2. Condition the StageTip with 100 μL of 100% ACN by passing the supernatant through the assembly by centrifugation at 3000 × g for 1 min. Wash with 100 μL of Stage Tip Solvent B by centrifugation at 3000 × g for 1 min (see Note 6). 3. Equilibrate the StageTip with 100 μL of Stage Tip Solvent A by centrifugation at 3000 × g for 1.5 min. Repeat this step for a total of two equilibration washes. 4. Load the acidified immunoaffinity peptide elution from Subheading 3.3, step 22 onto the StageTip and centrifuge at 3000 × g for 1.5 min. 5. Wash the peptides bound to the StageTip with 100  μL of Solvent A by centrifugation at 3000 × g for 1.5 min. Repeat this step for a total of two washes. 6. Elute the peptides with 50 μL of Stage Tip Solvent B into a new Eppendorf tube, and centrifuge at 3000 × g for 3 min to ensure all elution volume passes through. Subsequently, dry the peptide eluate completely in vacuo. 7. Resuspend the peptides in an appropriate volume of mobile phase A of your LC-MS system, e.g., 7 μL of 2% ACN/98% water/0.1% formic acid (v/v/v), add a retention time standard, for example, 0.1 μL of indexed retention time standard (iRT from Biognosys or other standards). Vortex the peptide solution for 10 min at 4 °C, and then centrifuge for 2 min at 12,000 × g and 4 °C. Transfer the supernatant to an autosampler vial for nano LC-MS/MS (see Note 7). 3.5  Nanoflow LC-MS/MS Analysis

1. Samples are analyzed by reverse-phase HPLC-ESI-MS/MS using the Eksigent Ultra Plus nano-LC 2D HPLC system combined with a cHiPLC System, directly connected to a quadrupole time-of-flight SCIEX TripleTOF 6600 mass spectrometer (SCIEX). Typically, mass resolution for precursor ion scans is ~45,000 (TripleTOF 6600), and fragment ion resolution is ~15,000 (“high sensitivity” product ion scan mode, see Note 8). After injection, peptide mixtures are transferred onto a C18 pre-column chip and washed at 2 μL/min for 10  min with the Mobile Phase A (loading solvent). Subsequently, peptides are transferred to the analytical column ChromXP C18-CL chip and eluted at a flow rate of 300 nL/ min typically with a 2–3 h gradient using aqueous and acetonitrile solvent buffers (Mobile Phases A and B). 2. Data-dependent acquisitions (DDA). For spectral library building, initial data-dependent acquisitions (DDA) are carried out to obtain MS/MS spectra for the 30 most abundant precursor ions (100  ms per MS/MS) following each survey MS1 scan (250 ms), yielding a total cycle time of 3.3 s.

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3. Data-independent acquisitions (DIA). For label-free relative quantification, all study samples are analyzed by data-­ independent acquisitions (DIA), using a 64 variable window SWATH acquisition strategy [9, 10]. Briefly, instead of the Q1 quadrupole transmitting a narrow mass range through to the collision cell, windows of variable width (5–90 m/z) are passed in incremental steps over the full mass range (m/z 400–1250). The cycle time of 3.2 s includes a 250 ms precursor ion scan followed by 45 ms accumulation time for each of the 64 DIA-­ SWATH segments. 3.6  Traditional MS Workflow A: Identification and Quantification of Acetylation Sites Using DDA & DIA

1. Mass spectrometric data from data-dependent acquisitions (DDA) is analyzed with the database search engine ProteinPilot 5.0 (SCIEX) using parameters such as trypsin digestion, cysteine alkylation set to iodoacetamide, lysine acetylation, and species Mus musculus; false discovery rates of 1% are used (see Note 9). 2. Using the database search engine results generated above, MS/MS spectral libraries are generated in Skyline daily v 4.1.1 [11], an open-source data processing workspace for quantitative proteomics; DIA raw data files are imported into Skyline and both MS1 precursor ion scans as well as MS2 fragment ion scans are extracted for all acetylated peptides present in the spectral libraries [12]. In Skyline, typically 6–10 MS2 fragment ions are extracted per acetylated peptide based on ranking from the corresponding MS/MS spectra in the spectral libraries, and fragment peak areas are summed per peptide. 3. Relative quantification of acetylation levels comparing different conditions, for example, knockout versus wild-type strains, can be performed directly in Skyline using integrated statistical algorithms. Statistical assessment of peak selection can be done within Skyline using mProphet [13], which was adjusted to specifics of DIA data. Alternatively, the corresponding extracted acetylation site peak areas can be exported and subjected to other open-source programs, such as mapDIA [14] which is specialized for processing and statistical analysis of quantitative proteomics data from DIA-MS (see Note 8).

3.7  Alternative MS Workflow B: Identification and Quantification of Acetylation Sites Using Exclusively DIA with PIQED Software [15]

In addition to the presented traditional protocol describing MS data acquisition (see Subheading 3.6), an alternative workflow (see Note 10) is presented that does not require DDA but only DIA, which can be advantageous when working with very low amounts of starting material or many replicates. 1. Data from DIA mass spectrometry only, without spectral libraries generated by DDA (as described under Subheading 3.6), can alternatively be analyzed using the automated pipeline PIQED and a corresponding java interface in which spectral

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libraries are generated directly from DIA data files that are subsequently also used for quantification [15]. Briefly, the raw data is converted to mzXML file formats, pseudo-MS/MS spectra are generated using DIA-Umpire [16], and acetylated peptides are identified by database search with MS-GF+ [17], Comet [18], and X! Tandem (thegpm.org). Subsequently, files are automatically imported into Skyline [12] and peak area results and outputs from acetylated peptides are submitted to mapDIA [14] for statistical analysis as part of the PIQED pipeline. 3.8  Anticipated Results

1. Depending on experimental design we typically identify and quantify 1000–2000 acetylation sites from an enrichment of 5 mg of protein lysate. See Fig. 2. 2. Numbers of acetylation sites vary per protein, ranging mostly between 1–20 detected sites per protein. 3. Workflow reproducibility is assessed between replicates, and we typically measure coefficients of variations CV  Duplicate (see Note 7). 3. Select the newly created copy, go to Image > Type > 8-bit. 4. With the same image selected, go to Process > Filters > Gaussian blur (see Note 8). Set the value at 3 in the pop-up window. 5. With the same image selected, go to Adjust > Threshold, set appropriate threshold to differentiate nuclei from background (Otsu preset usually works well, see Note 9). 6. OPTIONAL: go to Process > Binary > Watershed (see Note 10). 7. Go to Analyze > Set Measurements. Make sure Area (see Note 11), Standard deviation, Mean grey value, and Integrated density are selected. From the dropdown menu redirect to the image on which Thresholding was done. 8. Go to Analyze > Analyze particles. Select sizes 100 to Infinity (see Note 12), check Exclude on edges and Add to Manager. 9. Go to Analyze > Set measurements, redirect to the original DAPI channel file. 10. Click on ROI manager, Ctrl+A to select all ROIs and click Measure. A new window with results should appear. 11. Repeat steps 8–10 for the green (EdU) and red (PLA) channels.

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12. Select Red (PLA) channel, Image > Duplicate (see Note 13). 13. Go to Image > Type > 8-bit, then Image > Adjust > Threshold. Set an appropriate threshold that displays all unique foci (Intermodes preset usually works well). 14. Go to Process > Find Maxima. From the dropdown menu select Single Points, and check Light background and Preview Selection. 15. After clicking OK a new window should appear (see Note 14). Click Analyze > Set measurements, and redirect to this new window. 16. Select ROI manager, and click Measure. 17. Results page will now contain data for each nucleus. Press Ctrl+A to select all data, and paste into an Excel spreadsheet. Close all open images and results table without saving. 18. Repeat steps 1–17 for the next image. 3.9  Automating Image Processing

Image analysis can be automated by recording a macro of the steps delineated above. 1. With an image open, go to Plugins > Macros > Record. 2. Once the Recorder window is open, perform the steps above. 3. When done, click Create button. Save the macro and install by clicking Plugins > Macros > Install. Installed macros will appear under Plugins > Macros, at the bottom of the menu. 4. Open an image you want to process and click on the installed macro.

4  Notes 1. DUOLINK reagents are a convenient alternative to generating PLA probes and amplification reagents in-house. For the latter approach, the procedure is described in detail in [22]. 2. If tolerated, higher concentrations of EdU increase the level of incorporation of EdU into DNA and the probability of proximity of an EdU moiety to a target antigen [19]. Increasing EdU concentration should be considered when troubleshooting a low PLA signal. Shorter pulses of EdU reduce the level of incorporation but also limit detection to a closer vicinity of replication forks rather than to a longer tract of newly replicated DNA. 3. Blocking and antibody dilution buffer alternatives are supplied with Duolink PLA probes. If a user-preferred incubation buffer for IF is available for the primary antibodies, we recommend trying this buffer.

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4. PLA-rated antibodies are not required but are recommended. IF-rated antibodies are required. Optimization of antibody dilution and verification of its performance in IF is a good place to start. 5. Draining coverslips of excess wash liquid before every incubation with reagents is critical to avoiding low/absent PLA signal. No liquid should be pooled on a coverslip; however, it should not be dry. 6. Bio-formats function allows to work with images that are stored as more than one file. Bio-Formats offers options like grouping files with similar names, or distinguishing files based on unique identifiers. These features can be used to streamline the process of image analysis. 7. It is important to create a copy of a given channel because some of the actions of the next steps cannot be undone. 8. Gaussian Blur. The next step will allow to evaluate what is “positive” and “negative,” i.e., set a threshold for positive signal (Threshold function). Using the Threshold function may be challenging if the image has a lot of jagged edges. Gaussian Blur allows to smooth out the edges of nuclei and perform thresholding more accurately. Gaussian Blur should not be done on original files that will be used to collect quantitative data; it should only be performed on duplicates. 9. This step should be performed on a duplicate DAPI channel image that has been Gaussian blurred. In general, every pixel in an 8 bit image will hold a value between 0 and 255. This value is the “brightness” of the pixel. Thresholding allows to set a cutoff pixel value that will enable the software to “find” and mark nuclei in the image. In a typical image of DAPI-­ stained nuclei with a big differential between the signal and the background, presets such as OTSU work very well. 10. When cell density in an image is high, the software will interpret adjoining nuclei as a single nucleus. If this happens often enough it may skew the results. Watershed function draws a 1-pixel-wide line between objects that is considered adjoined. Note, however, that it may also interpret an irregularly shaped nucleus as separate, adjoined nuclei. Ultimately, it is up to a user to make a call to watershed or not. 11. The software does not read the scaling information of the image by default. This information can be found in a metadata file that is generated for each image by the image acquisition software. However, in our case the absolute dimensions of cells are less important than relative differences between the samples. For example, if comparing two cell lines it is important to know if they have the same or different nuclear sizes in order

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to choose mean or total nuclear fluorescence as a readout. Thus, it is recommended to record nuclear area data per nucleus. In addition, Area value allows to track one and the same nucleus through different channels because it will remain a constant. To set the scale, go to Analyze > Set Scale. For a relative scale, 1 pixel can be set as 1 μm. 12. Size value minimum of at least 100 pixels will exclude most noise and false positive signals. Selecting different size cutoff values may help to exclude aggregates or broken cells. Exclude on edges box decides if the software will consider something that is halfway outside of the picture as an ROI. Add to manager box needs to be selected for ROIs to be stored in the ROI manager. 13. This and next steps measure PLA foci number in the nuclei. When PLA signals are sparse or of moderate to low brightness, it may be valuable to count PLA foci in each nucleus rather than measuring total signal strength per nucleus. To do this, first select the channel that has the PLA signal, then run Thresholding. The selected threshold preset needs to display as many individual foci as possible without having them merge together. 14. In the new image the software has replaced each focus with a single pixel with a value 255. If you perform measurements on this new image with the ROIs created earlier you will see in the Results tab that for each ROI the software summed up the measurements. That is, if a nucleus had one focus, the value will be 255, if there were two foci the value will be 510 and so on. Downstream processing in Excel will require dividing this column by 255 to get actual count of PLA foci per nucleus.

5  Discussion of Data Analysis and Presentation Data on signal intensity and foci counts, compiled in Excel, can be analyzed in Excel or with any statistical and graphing software. Nonparametric statistical tests should be used to assess the significance of differences between samples. Figure 2a, b, d shows examples of detection of acetylated histones H4 (H4K12ac) and H3 (H3K9ac). One aspect of detection of proteins on EdU-labeled DNA is that only a fraction of cells are in S phase during a typical pulse-labeling. Cells outside S phase will remain EdU-negative and thus will not display any PLA signal. These EdU-negatives should be distinguished from experimental negatives resulting from the too-rare or absent proximity of the antigens under study, or from technical failure. In other cases, in order to properly quantify PLA signal, one needs to assess the level

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Fig. 2 Detection and quantitation of histone H4K12ac and H3K9ac at newly replicated DNA. (a) A representative example of simultaneous detection of EdU incorporation (with AlexaFluor488 azide) and H4K12ac/EdU PLA (with biotin azide) in SV40-transformed human fibroblast line GM639 labeled with EdU for 30 min and fixed. Note that PLA signal is predominantly limited to EdU-positive cells. (b) A blow-up of a single nucleus showing H4K12ac/EdU PLA and total EdU signals. (c) Left panel: Scatterplots of unfiltered EdU (green) and PLA (red) channel fluorescence intensities per nucleus measured in the images similar to (a). Click-It reactions were performed with biotin azide only or with mixtures of indicated molar ratios of biotin and AlexaFluor488 azides. Right panel: the indicated data from the left panel are separated into EdU-negative and EdU-positive subsets based on the EdU signal cutoff. Data means are indicated by black lines. (d) An example of detection of H3K9ac at newly replicated DNA in GM639 fibroblasts. Only the PLA signal is visualized in this experiment. (e) Scatterplots of PLA foci numbers measured in the images represented by (d). Left panel: Click-It reactions were performed with biotin azide only or with mixtures of indicated molar ratios of biotin and AlexaFluor488 azides. The data derived with mixed azides are subsetted into EdU-negative and EdU-positive groups. Right panel: based on the observed number of PLA foci in EdU-negative cells, inferences can be made as to what fraction of PLA-negatives in the biotin azide-only set comes from EdU-negative (non S phase cells). Arrows above the scatterplots illustrate the origin of the subsetted data

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of EdU incorporation. Lower (or higher) EdU incorporation in experiment versus control samples may lead to a spuriously low (or high) PLA signal. There are several ways to account for these contingencies. “Spiking” biotin azide with a sub-stoichiometric dose of a fluorescent azide (Subheading 3.3, step 4) allows detecting EdU-­ positive, S phase cells without sacrificing too much of the PLA signal (Fig. 2a–c). The molar ratio of the two azides should be optimized since PLA signal intensity is obviously affected by introduction of a fluorescent azide that reduces the pool of EdU available for PLA interaction (Fig. 2c). However, the approach helps to identify and measure the “true” PLA signal distribution in S phase cells. For example, in Fig. 2c the EdU signal distribution revealed by AlexaFluor488 green fluorescence is clearly bimodal, corresponding to the signal levels in the S phase, EdU-positive cells, and EdU-negative cells (background). In the right panel of Fig. 2c, the PLA data from the spiked samples are now subsetted based on EdU status. It is clear that this step adjusts the PLA signal distribution to remove PLA-negatives coming from non-S phase cells and reveal the true signal. In Fig. 2e PLA signal of H3K9ac is measured as foci number rather than mean fluorescence intensity per nucleus, and, as can be seen in the left panel in a non-spiked sample quite a few cells show no or very few foci. By comparing the foci numbers in spiked and non-spiked samples, it can be inferred that less than 5 PLA foci for this particular antibody and conditions likely represent background staining, and the PLA data in a non-spiked sample can be subsetted accordingly. This experimentally verified cutoff can be then used throughout the experimental series while working with non-spiked samples. Accounting for EdU incorporation can serve another purpose when quantifying PLA data. In Fig. 3a, the level of H4K12ac at replication forks is measured in cells treated with histone deacetylase HDAC1 and 2 inhibitor, CI994. Previous work, including ours, showed that this treatment increases the level of histone H4K12 acetylation at replication forks and reduces the speed of fork progression, leading to less EdU incorporation per cell [8, 9]. Both of these effects can be detected by measuring EdU and PLA signals (Fig. 3a). However, reduced EdU incorporation in the CI994-treated sample masks the extent by which the increased H4K12ac abundance increases the PLA signal. In this case, normalizing PLA signal to EdU signal for each cell can better highlight the difference. There are cases where spiking biotin azide with fluorescent azide may be less advantageous. One obvious counter-indication is when working with low/rare PLA signals. Also, when performing a chase after an EdU pulse, residual incorporation of progressively

Fig. 3 Using normalization of PLA data in addressing biological questions. (a) Simultaneous detection of EdU (green) and H4K12ac/EdU PLA (red) signals allows to adjust PLA signal values based on differences in EdU incorporation between experiment and control. GM639 fibroblasts were treated with 6 μM CI994 for 6 h,

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more diluted EdU into DNA may contribute to the overall EdU signal but not to PLA signal due to a decreased density of these extra EdU moieties in DNA, skewing the data after PLA to EdU normalization. For these and other situations, a validated alternative to spiking is to measure EdU incorporation in a duplicate set of samples [19, 27, 28]. Specifically, PLA-competent EdU in DNA can be quantified by clicking EdU to biotin azide and performing PLA between mouse and rabbit anti-biotin antibodies (Fig. 3b, c). In this application, the H3K9ac/EdU PLA signals are normalized to the median of the EdU/EdU PLA signals. Using this method, we can demonstrate, for example, that p53 null HAP-1 cells show a drastic decrease in H3K9ac level at hydroxyurea-stalled DNA replication forks compared to wild-type (wt) HAP-1 cells, consistent with our previous finding that p53 affects DNA replication restart [28]. Notably, decreased H3K9 acetylation in p53-deficient compared to p53-proficient cells has previously been described at sub-telomeric regions after etoposide treatment, suggesting a common mechanism [29]. In summary, findings shown in Fig. 3 exemplify the utility of the described method to measure localized histone modifications at actively replicating DNA as well as sites where replication forks have stalled due to nucleotide depletion or damage.

Acknowledgments This work was supported by NIH grants GM115482 and CA215647 to J.S. and Cancer Prevention Research Institution of Texas grant R1312 to K.S. K.S. was also supported by fellowships from Rita Allen Foundation and Andrew Sabin Family Foundation, and Cancer Prevention Research Institution of Texas Scholarship in Cancer Biology.

Fig. 3 (continued) labeled with 20 μM EdU for 30 min, and fixed. Click-It reactions were performed with biotin azide at 50:1 molar excess over AlexaFluor488 azide. Left panel: beanplots of signal intensity data in EdUpositive cells. Right panel: H4K12ac/EdU PLA signal in each nucleus was normalized to EdU signal and plotted as beanplots. Statistical significance was determined in K-S tests. Y axes are at log scale. (b) A representative example of H3K9ac/EdU PLA (top panels) and EdU/EdU PLA (bottom panels) in HAP-1 wild-type and p53-null cells pulsed with 125 μM EdU for 8 min, then incubated with 0.2 mM of hydroxyurea for 4 h. PLA assays were performed on duplicate samples. (c) Scatterplots of data measured in the images represented by (b). In the right panel, H3K9ac/EdU signal in each nucleus of a given sample was normalized to the sample-specific median EdU/EdU PLA signal. Statistical significance was measured in Mann-Whitney tests. Note the log scale of Y axes in the non-normalized data. Arrows above the scatterplots in (a) and (c) illustrate the lineage of the normalized data

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References 1. Alabert C, Groth A (2012) Chromatin replication and epigenome maintenance. Nat Rev Mol Cell Biol 13:153–167 2. Gong F, Miller KM (2013) Mammalian DNA repair: HATs and HDACs make their mark through histone acetylation. Mutat Res 750:23–30 3. Galvani A, Thiriet C (2015) Nucleosome dancing at the tempo of histone tail acetylation. Genes (Basel) 6:607–621 4. Gong F, Chiu LY, Miller KM (2016) Acetylation reader proteins: linking acetylation signaling to genome maintenance and cancer. PLoS Genet 12:e1006272 5. Alabert C, Jasencakova Z, Groth A (2017) Chromatin replication and histone dynamics. In: Masai H, Foiani M (eds) DNA replication: from old principles to new discoveries. Springer Singapore, Singapore, pp 311–333 6. Nagarajan P, Ge Z, Sirbu B, Doughty C, Agudelo Garcia PA, Schlederer M, Annunziato AT, Cortez D, Kenner L, Parthun MR (2013) Histone acetyl transferase 1 is essential for mammalian development, genome stability, and the processing of newly synthesized histones H3 and H4. PLoS Genet 9:e1003518 7. Ge Z, Nair D, Guan X, Rastogi N, Freitas MA, Parthun MR (2013) Sites of acetylation on newly synthesized histone H4 are required for chromatin assembly and DNA damage response signaling. Mol Cell Biol 33:3286–3298 8. Bhaskara S, Jacques V, Rusche JR, Olson EN, Cairns BR, Chandrasekharan MB (2013) Histone deacetylases 1 and 2 maintain S-phase chromatin and DNA replication fork progression. Epigenetics Chromatin 6:27 9. Kehrli K, Phelps M, Lazarchuk P, Chen E, Monnat R Jr, Sidorova JM (2016) Class I histone deacetylase HDAC1 and WRN RECQ helicase contribute additively to protect replication forks upon hydroxyurea-induced arrest. J Biol Chem 291:24487–24503 10. Annunziato AT, Seale RL (1983) Histone deacetylation is required for the maturation of newly replicated chromatin. J Biol Chem 258:12675–12684 11. Benson LJ, Gu Y, Yakovleva T, Tong K, Barrows C, Strack CL, Cook RG, Mizzen CA, Annunziato AT (2006) Modifications of H3 and H4 during chromatin replication, nucleosome assembly, and histone exchange. J Biol Chem 281:9287–9296 12. Sirbu BM, Couch FB, Feigerle JT, Bhaskara S, Hiebert SW, Cortez D (2011) Analysis of pro-

tein dynamics at active, stalled, and collapsed replication forks. Genes Dev 25:1320–1327 13. Alabert C, Bukowski-Wills J-C, Lee S-B, Kustatscher G, Nakamura K, de Lima Alves F, Menard P, Mejlvang J, Rappsilber J, Groth A (2014) Nascent chromatin capture proteomics determines chromatin dynamics during DNA replication and identifies unknown fork components. Nat Cell Biol 16:281–293 14. Filippakopoulos P, Knapp S (2014) Targeting bromodomains: epigenetic readers of lysine acetylation. Nat Rev Drug Discov 13:337–356 15. Chiu LY, Gong F, Miller KM (2017) Bromodomain proteins: repairing DNA damage within chromatin. Philos Trans R Soc Lond Ser B Biol Sci 372:20160286 16. Fujisawa T, Filippakopoulos P (2017) Functions of bromodomain-containing proteins and their roles in homeostasis and cancer. Nat Rev Mol Cell Biol 18:246–262 17. Sirbu BM, McDonald WH, Dungrawala H, Badu-Nkansah A, Kavanaugh GM, Chen Y, Tabb DL, Cortez D (2013) Identification of proteins at active, stalled, and collapsed replication forks using isolation of proteins on nascent DNA (iPOND) coupled with mass spectrometry. J Biol Chem 288:31458–31467 18. Sirbu BM, Couch FB, Cortez D (2012) Monitoring the spatiotemporal dynamics of proteins at replication forks and in assembled chromatin using isolation of proteins on nascent DNA. Nat Protocols 7:594–605 19. Roy S, Luzwick JW, Schlacher K (2018) SIRF: quantitative in situ analysis of protein interactions at DNA replication forks. J Cell Biol 217:1521–1536 20. Petruk S, Sedkov Y, Johnston DM, Hodgson JW, Black KL, Kovermann SK, Beck S, Canaani E, Brock HW, Mazo A (2012) TrxG and PcG proteins but not methylated histones remain associated with DNA through replication. Cell 150:922–933 21. Petruk S, Cai J, Sussman R, Sun G, Kovermann SK, Mariani SA, Calabretta B, McMahon SB, Brock HW, Iacovitti L et al (2017) Delayed accumulation of H3K27me3 on nascent DNA is essential for recruitment of transcription factors at early stages of stem cell differentiation. Mol Cell 66:247–257.e245 22. Soderberg O, Gullberg M, Jarvius M, Ridderstrale K, Leuchowius KJ, Jarvius J, Wester K, Hydbring P, Bahram F, Larsson LG et al (2006) Direct observation of individual

Acetylated Histone Detection in Situ on Replicating DNA endogenous protein complexes in situ by proximity ligation. Nat Methods 3:995–1000 23. Fredriksson S, Gullberg M, Jarvius J, Olsson C, Pietras K, Gustafsdottir SM, Ostman A, Landegren U (2002) Protein detection using proximity-dependent DNA ligation assays. Nat Biotechnol 20:473–477 24. Iannascoli C, Palermo V, Murfuni I, Franchitto A, Pichierri P (2015) The WRN exonuclease domain protects nascent strands from pathological MRE11/EXO1-dependent degradation. Nucleic Acids Res 43:9788–9803 25. Weibrecht I, Gavrilovic M, Lindbom L, Landegren U, Wahlby C, Soderberg O (2012) Visualising individual sequence-­ specific protein-DNA interactions in situ. New Biotechnol 29:589–598 26. Zhang W, Xie M, Shu MD, Steitz JA, DiMaio D (2016) A proximity-dependent assay for spe-

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cific RNA-protein interactions in intact cells. RNA (New York, NY) 22:1785–1792 27. Taglialatela A, Alvarez S, Leuzzi G, Sannino V, Ranjha L, Huang JW, Madubata C, Anand R, Levy B, Rabadan R et al (2017) Restoration of replication fork stability in BRCA1- and BRCA2-deficient cells by inactivation of SNF2-­ family fork remodelers. Mol Cell 68:414–430. e418 28. Roy S, Tomaszowski KH, Luzwick JW, Park S, Li J, Murphy M, Schlacher K (2018) p53 orchestrates DNA replication restart homeostasis by suppressing mutagenic RAD52 and POLtheta pathways. Elife 7:e31723 29. Tutton S, Azzam GA, Stong N, Vladimirova O, Wiedmer A, Monteith JA, Beishline K, Wang Z, Deng Z, Riethman H et al (2016) Subtelomeric p53 binding prevents accumulation of DNA damage at human telomeres. EMBO J 35:193–207

Part II Lysine Acetylation

Chapter 4 Quantification of In Vitro Protein Lysine Acetylation by Reversed Phase HPLC Catherine W. Njeri, Onyekachi E. Ononye, and Lata Balakrishnan Abstract Protein lysine acetylation is a reversible posttranslational modification that is catalyzed by a group of enzymes that are collectively referred to as lysine (K) acetyltransferases (KATs). These enzymes catalyze the transfer of the acetyl group from acetyl coenzyme A (Ac-CoA) to the ε-amino group of lysine amino acid. Protein lysine acetylation plays a critical role in the regulation of important cellular processes and it is therefore paramount that we understand the catalytic mechanisms of these enzymes. While there is a variety of methods that have been developed to analyze the enzymatic properties of KATs, majority of the proposed methods have considerable limitations. We describe here a reversed phase HPLC based method that monitors substrate consumption and product formation simultaneously. This method is highly reproducible and optimally suited for the determination of accurate kinetic parameters of KATs. Key words In vitro protein lysine acetylation, Lysine acetyltransferases (KATs), Reversed phase high performance liquid chromatograph (HPLC)

1  Introduction Histone acetyltransferases (HATs) or lysine acetyltransferases (KATs) are a group of enzymes that catalyze the transfer of an acetyl group of acetyl-CoA to a specific lysine residue on the protein substrate [1]. This posttranslational modification neutralizes the positively charged lysine residue, which consequently affects the functional and structural properties of the modified protein. Modification of histone tails by acetylation weakens the interaction between the histone proteins and DNA [2]. This weakened interaction exposes the DNA, making it more accessible to biological machineries [3]. In addition to acetylating histone proteins and upregulating transcription, HATs have been shown to acetylate a variety of nonhistone proteins [4–6]. The functional consequence of lysine acetylation on nonhistone proteins has been a subject under intense investigation due to its implication in a variety of

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crucial biological processes including DNA replication and repair, cell cycle progression, apoptosis, tumorigenesis, and a myriad of other processes [7, 8]. There are several biochemical assays that have been developed for the determination of the kinetic properties of acetyltransferases [1]. These assays have been designed to either detect the acetylation of the protein substrate or the formation of the free CoASH product. In one of the assays that involves the measurement of the acetylated protein product, the acetyl-CoA substrate used is radioactively labeled with either 3H or 14C. Upon acetylation, the radiolabeled protein product formed is then analyzed either by autoradiography or by liquid scintillation counting [9]. This method, however, is costly and the use of radioactive material poses potential health risks and environmental hazards [9]. Additionally, if the level of acetylation is relatively low, detection of acetylation via autoradiography may require longer exposure times. Mass spectrometry can also be used to analyze in vitro acetylated peptides and proteins [10]. The main drawbacks to this method is that it can be very expensive, requires specialized technical expertise and specific instrumentation. Methods that are geared toward the measurement of the free CoASH product generated in an acetyltransferase reaction can either utilize fluorescent probes that target the thiol group of CoASH or can utilize enzyme-coupled reactions [1]. Although these methods tend to be straightforward and relatively inexpensive, the quality of the assays can potentially be limited due to interferences that might arise from reagents in the sample [1]. For instance, trace amounts of thiol containing compounds in the sample can be a source of significant errors in fluorometric assays that target the thiol group of CoASH. The reversed phase HPLC method outlined in this protocol is designed in such a way that the acetyl-CoA substrate and the CoASH product can be monitored simultaneously by recording absorbance at 260 nm. Additionally, changing the wavelength to 215 nm allows one to monitor and differentiate the acetylated and unmodified protein substrates. In the assay outlined in this protocol, the H3 peptide is in vitro acetylated by the HAT domain of human p300. The reaction mixture is then injected into the HPLC system which monitors the UV spectra of the various components of the reaction mixture. Each compound in the reaction mixture is eluted from the C18 column at different rates allowing the compounds to be chromatographically separated. The respective absorbance peaks are recorded and the corresponding areas under the peak are used to determine the concentration of substrate consumed and product formed. This method is highly dependable and can be applied to determine the efficiency of an acetyltransferase in modifying a specific recombinant protein.

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2  Materials 1. All the reagents used in the experiments should be of the highest available purity. 2. The instrument used to perform these experiments was Varian ProStar HPLC system equipped with Varian Galaxie Chromatography Data System software version 1.9.3.2. 3. Acetyl-CoA sodium salt was dissolved in nuclease-free water. 4. Coenzyme A Free acid was dissolved in nuclease-free water. 5. KH2PO4 and K2HPO4 dissolved in water to make 1 M potassium phosphate buffer pH 7.0. 6. Acetonitrile, HPLC grade. 7. Acetic Acid (30%). 8. H3 and H4 peptides (Active Motif) were resuspended in 1× histone acetyltransferase (HAT) buffer (recipe below). 9. 1× histone acetyltransferase (HAT) buffer: 50 mM Tris–HCl [pH  8.0], 1  mM dithiothreitol, 10  mM sodium butyrate, 10  mM sodium chloride, 1  mM phenylmethylsulfonyl fluoride, and 10% (v/v) glycerol. 10. Reversed phase HPLC column used is TARGA C18 5  μm, 150 × 4.6 mm. 11. Hamilton syringe for HPLC. 12. HPLC washing Solvent A: 50  mM potassium phosphate [pH 7.0]. 13. HPLC elution Solvent B: 60% Acetonitrile in 50 mM potassium phosphate [pH 7.0].

3  Methods 3.1  In Vitro Acetylation Reaction

1. The HAT used in this assay was commercially obtained, but recombinantly expressed and purified acetyltransferases can also be used. Purity of these acetyltransferases should be confirmed before using them in the reaction. 2. Initial in  vitro acetylation reactions should be done at 1:1:10 ratio (peptide/protein:HAT:acetyl-CoA). Depending on the efficiency of acetylation reactions, these ratios can be subsequently adjusted. A typical HAT assay consists of the 1× HAT buffer in a final volume of 20 μL, HAT (100 nM), protein to be acetylated (100  nM), and a range of concentrations of acetyl-­ CoA (1–10 μM). The order of addition of the reagents in a typical acetylation reaction is highly important (refer to Note 1). 3. First, the appropriate volume of the 1X HAT buffer is added to a 1.5 mL Eppendorf tube. This is followed by the peptide

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or protein to be acetylated, and then the HAT.  Finally, the reaction is initiated by the addition of an appropriate concentration of acetyl-CoA. 4. Following the addition of acetyl-CoA, gently tap the reaction mixture to ensure efficient mixing of reagents. 5. Incubate the reactions at 37 °C for 30 min. 6. When the acetylation reaction is complete, terminate the reaction by the addition of 30% acetic acid. 7. The sample is now ready to be analyzed by HPLC. 1. The solvents that are used in this assay must be filtered through a 0.2 μm filter and degassed prior to use. Before sample injection into the HPLC system, the column needs to be equilibrated in the following order; 30  min with distilled water; 30 min with 100% solvent B, and 30 min with 100% solvent A (refer to Notes 2–4).

3.2  Reversed Phase HPLC Assay Preparation

2. An elution gradient is then established (Fig. 1) to determine the ratio between solvent A and solvent B that will provide the best separation of the compounds of interest (refer to Note 5). 3. Table 1 describes an elution gradient used to separate acetyl-­ CoA and CoASH at a flow rate of 1 mL/min. This gradient is designed such that it facilitates the separation of acetyl-CoA and CoASH based on the difference in their retention times. 4. The gradient also includes a column cleaning step in which the gradient steeps up to 100% solvent B allowing the elution of all the compounds present in the sample. To ensure that the column is completely clean, wash the column with 100% B for 3–5 min. 5. The last step is the re-equilibration step where the column is washed with 100% solvent A for at least 1 min. 1. After the elution gradient has been created and the column has been adequately equilibrated, inject 10 μL of an appropriate concentration (in μM) of Acetyl-CoA. Monitor the elution

3.3  Quantification of CoASH

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Fig. 1 The elution gradient designed for the separation of acetyl-CoA from CoASH

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Table 1 HPLC method for the analysis of acetyl-CoA and CoASH Time, min

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Fig. 2 (a) A representative chromatogram showing the elution profile of acetyl-CoA. (b) A representative chromatogram showing the elution profile of CoASH

of acetyl-CoA from the column by recording absorbance at 260 nm (refer to Note 6). In our assay, the acetyl-CoA peak is recorded at a retention time of 9.6 min (Fig. 2a). 2. In order to accurately quantify the amount of free product formed by the acetylation reaction, inject varying concentrations of CoASH and record absorbance at 260  nm. In our assay, free CoASH elutes faster than Acetyl-CoA with a retention time of 8.4  min (Fig.  2b). The area under the curve is

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Fig. 3 A representative chromatogram showing the separation of CoASH and acetyl-CoA

integrated using the Varian Galaxie Chromatography Data System software. 3. Plot the determined areas against the respective concentrations of CoASH to create a standard curve that will be used to quantify the amount of CoASH formed from the acetylation reactions. 3.4  Analysis of the In Vitro Acetylation Reactions

1. Inject 10 μL of each acetylation reaction into the HPLC system and monitor the absorbance at 260  nm. As previously mentioned, the HPLC assay method is extremely versatile in the sense that both the CoASH formed and the Acetyl-CoA used can be monitored in a single profile (Fig. 3). 2. The area under the curve of both acetyl-CoA and CoASH is then determined and the amount of product formed is evaluated using the previously created standard curve.

4  Notes 1. In any acetylation reaction, it is crucial that you add the reagents to the reaction tube in the correct order. We have noticed that if you add acetyl-CoA to the HAT before addition of the target protein to be acetylated, the amount of acetylated protein product is significantly low, due to HAT autoacetylation. As such, always mix the HAT with the protein to be acetylated first in the reaction buffer, before the addition of acetyl-CoA.

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2. It is important to properly equilibrate your column prior to injecting your samples for analysis. The column might contain some compounds from previous experiments that might interfere with your assays. 3. Initially washing your column with water prevents precipitation of salts and other compounds that might still be in the column. 4. The solvents used with the HPLC system must always be filtered prior to their use. This includes the distilled water that is used at the beginning to wash the column. Filtering your solvents prolongs the life of your column as it reduces the buildup of particulate matter and also improves peak resolution. 5. Designing a good elution gradient method is crucial to the resolution of your samples. This is purely experimental and varies from assay to assay based on the solvent systems used. 6. The wavelength that was used in the assay described here was set to 260 nm. In case you want to evaluate the acetylated and the unmodified protein, set the wavelength to 215 nm when designing the elution method.

Acknowledgments This work was supported by NIH Grant GM0938328 and New Faculty Start-Up Funds from Indiana University Purdue University Indianapolis (IUPUI). References 1. Gadhia S, Shrimp JH, Meier JL, McGee JE, Dahlin JL (2004) Histone acetyltransferase assays in drug and chemical probe discovery. In: Sittampalam GS, Coussens NP, Brimacombe K et al (eds) Assay guidance manual. Eli Lilly & Company and the National Center for Advancing Translational Sciences, Bethesda, MD 2. Hong L, Schroth GP, Matthews HR, Yau P, Bradbury EM (1993) Studies of the DNA binding properties of histone H4 amino terminus. Thermal denaturation studies reveal that acetylation markedly reduces the binding constant of the H4 “tail” to DNA.  J Biol Chem 268(1):305–314 3. Yang XJ, Seto E (2008) Lysine acetylation: codified crosstalk with other posttranslational modifications. Mol Cell 31(4):449–461. https:// doi.org/10.1016/j.molcel.2008.07.002

4. Close P, Creppe C, Gillard M, Ladang A, Chapelle JP, Nguyen L, Chariot A (2010) The emerging role of lysine acetylation of non-nuclear proteins. Cell Mol Life Sci 67(8):1255–1264. https://doi.org/10.1007/ s00018-009-0252-7 5. Glozak MA, Sengupta N, Zhang X, Seto E (2005) Acetylation and deacetylation of non-­ histone proteins. Gene 363:15–23. https:// doi.org/10.1016/j.gene.2005.09.010 6. Choudhary C, Kumar C, Gnad F, Nielsen ML, Rehman M, Walther TC, Olsen JV, Mann M (2009) Lysine acetylation targets protein complexes and co-regulates major cellular functions. Science (New York, NY) 325(5942):834–840. https://doi. org/10.1126/science.1175371 7. Di Martile M, Del Bufalo D, Trisciuoglio D (2016) The multifaceted role of lysine acetyla-

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tion in cancer: prognostic biomarker and therapeutic target. Oncotarget 7(34):55789–55810. https://doi.org/10.18632/oncotarget.10048 8. Goodman RH, Smolik S (2000) CBP/p300 in cell growth, transformation, and development. Genes Dev 14(13):1553–1577 9. Evjenth R, Hole K, Ziegler M, Lillehaug JR (2009) Application of reverse-phase HPLC to quantify oligopeptide acetylation eliminates

interference from unspecific acetyl CoA hydrolysis. BMC Proc 3(Suppl 6):S5. https://doi. org/10.1186/1753-6561-3-s6-s5 10. Karanam B, Jiang L, Wang L, Kelleher NL, Cole PA (2006) Kinetic and mass spectrometric analysis of p300 histone acetyltransferase domain autoacetylation. J  Biol Chem 281(52):40292–40301. https://doi. org/10.1074/jbc.M608813200

Chapter 5 Measurement and Analysis of Lysine Acetylation by KAT Complexes In Vitro and In Vivo Anahita Lashgari, Jean-Philippe Lambert, and Jacques Côté Abstract The acetylation of the ε-amine of lysine residues has significant impacts on the cellular functions of proteins. Through the combination of unbiased and targeted analysis of acetylated proteins, biological insights on lysine acetylation are now routinely generated. To help in this endeavor, we describe detailed protocols for the identification of acetylated lysine residues and the preparation of multiple reagents for the characterization of these sites in order to obtain functional insights on this widespread modification. Key words Lysine acetylation, KAT, HAT, TAP, LC-MS/MS

1  Introduction Protein post-translational modifications (PTMs) introduce novel characteristics to the modified proteins through altering molecular structure and geometry, resulting in changes in enzymatic activity, interaction partners, subcellular localization, protein stability, and DNA binding. Among the PTMs, acetylation of the ε-amine of lysine residues has emerged as one of the most abundant which plays an important role in epigenetic processes, as well as the regulation of protein–protein interactions and protein stability [1]. This modification was first discovered on histones associated with gene transcription and, subsequently, the catalyzing enzymes were termed histone acetyltransferases (HATs) [2, 3]. Acetylation neutralizes the lysine’s positive charge and destabilizes the interaction between histones and DNA. Acetylated lysine residues also function as binding sites for epigenetic reader protein modules such as bromodomains, YEATS, and PHD fingers to mediate protein– protein interactions [4]. In addition to histones, acetyltransferases are now known to modify a variety of other proteins, including

Jean-Philippe Lambert and Jacques Côté are both corresponding author. Robert M. Brosh, Jr. (ed.), Protein Acetylation: Methods and Protocols, Methods in Molecular Biology, vol. 1983, https://doi.org/10.1007/978-1-4939-9434-2_5, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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transcription factors and metabolic enzymes. High-resolution mass spectrometry-based proteomics screens, in combination with immunoprecipitation with anti-acetyl-lysine antibodies, have allowed the identification of >10,000 acetylation sites on ~3000 human proteins from Jurkat cells [5] and >21,000 sites on ~5300 proteins from mouse embryonic fibroblast [6]. In fact, as of writing this chapter, the PhosphoSitePlus(R) repository (www.phosphosite.org) reports 21,228 acetylation sites on 6927 distinct human proteins [7]. Lysine acetylation is a reversible modification catalyzed by lysine acetyltransferases (KATs) and removed by lysine deacetylases (KDACs). Acetylation and deacetylation of lysines is a tightly regulated process in living cells, and its misregulation is associated with many diseases including cancer. Furthermore, several KATs, KDACs, and bromodomain-containing proteins are linked to cancer development [8] and have emerged as important therapeutic targets [9–11]. Despite the significant increase in our understanding of protein acetylation by global proteomic analyses and functional studies in recent years, there is still much to be understood about the functions of this abundant PTM. Access to purified native KAT complexes is an invaluable tool to study the precise function of these enzymes in vitro. In this chapter we present a series of protocols to study KAT complexes and acetylated protein. First, we described an efficient protocol to purify native KAT complexes from human cells stably expressing near physiological levels, based on genome editing and tandem affinity purification (TAP). This approach was previously described in a publication by the Côté and Doyon labs [12, 13] and can also be used to purify any human protein or complex to study, for example, their native acetylation status. In the first part, we describe a method to generate isogenic cell lines expressing TAP-tagged cDNAs using CRISPR-mediated targeted integration into a previously described safe harbor, a genomic locus known as AAVS1 [14], for stable transgenic cell line generation in K562 cells (or any human cell line). We then describe preparation of nuclear extracts from large-scale culture of these cells followed by TAP purification of the tagged protein to allow their use in functional/ biochemical and proteomic assays. We further discuss two different approaches that can be combined to study the purified complexes: (a) lysine acetyltransferase assay and (b) a protocol for detection of acetylated lysine residues by mass spectrometry, employing small-­ scale affinity purification of acetylated-lysine peptides, followed by their desalting and concentration using C18-StageTips. In parallel, we also describe a simple protocol to analyze acetylated proteins in a crude mixture/extract, using immunoprecipitation with anti-­ acetyl-­lysine antibodies. To study the impact of specific lysine(s) acetylation on the structure and activity of a protein, we need to obtain a homoge-

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neously acetylated form of the protein. The study of the detailed structure of acetylated proteins is therefore impeded by the limited availability of these homogeneously acetylated proteins (i.e., on a specific lysine residue). Here, we describe a general method to produce homogeneously and site-specifically acetylated recombinant proteins in E. coli by an orthogonal N3-acetyllysyl-tRNA synthetase/ tRNACUA pair, originally described by the Chin lab [15, 16]. See Fig. 1.

Fig. 1 Purification of native lysine acetyltransferase complexes and enzymatic assays. (a) Tandem-affinity purification (TAP) of native human SAGA acetyltransferase complex from engineered K562 cells. Silver-stained SDS–PAGE showing the SAGA complex obtained when expressed from the AAVS1 (SUPT7L-AAVS1) loci (Subheading 3.1). The first and the second eluates (E1 and E2, respectively) of FLAG and Strep elution steps are shown. (b) In vitro liquid histone acetyltransferase (HAT) assays with purified native complex from panel (a) using short oligonucleosomes as substrate. The purified SAGA complexes shows HAT activity toward native nucleosomes purified from HeLa cells. (c) The same liquid HAT assays were processed for fluorography on 15% SDS-PAGE. The gel was first stained with Coomassie as loading control, then treated with EN3HANCE, dried, and exposed on film

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2  Materials 2.1  Preparation of Native KAT Complexes from 3 L Cultures

1. K562 cells (ATCC; CCL-243). 2. AAVS1_Puro_PGK1_3×FLAG_Twin_Strep #68375).

(Addgene

3. AAVS1-targeting CRISPR plasmid (contains both gRNA and Cas9 sequences): eSpCas9(1.1)_No_FLAG_AAVS1_T2 (Addgene #79888). 4. RPMI 1640 medium [+] GlutamaxTM [+] 25 mM Hepes (Gibco). 5. RPMI 1640 medium [+] l Glutamine (Gibco). 6. RPMI 1640, 2× (Wisent). 7. New born calf serum (NBCS) (Wisent). 8. Fetal Bovine Serum (FBS). 9. Opti-MEM (Gibco). 10. Methylcellulose (Sigma). 11. GlutaMAX™ Supplement (Gibco). 12. Pen strep 10,000 U/mL (Gibco). 13. Lipofectamine 2000. 14. Puromycin. 15. 6-well plates. 16. 100 mm plates. 17. 96-well plates. 18. T175 Flasks. 19. Three-liter capacity spinner. 20. 50 mL canonical tubes. 21. 15 mL canonical tubes. 22. 40 mL Dounce homogenizer type B pestle. 23. 15 mL Dounce homogenizer type B pestle. 24. PBS (Phosphate-buffered saline): 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, pH 7.4. 25. Hypotonic buffer: 10 mM Hepes-NaOH pH 7.9, 1.5 mM MgCl2, 10 mM KCl, 1 mM DTT, 1 mM PMSF, 2 μg/mL Leupeptin, 5 μg Aprotinin, 2 μg/mL Pepstatin, 10 mM Na-­ butyrate, 10 mM β-glycerophosphate, 100 μM Na-­ orthovanadate, 5 mM N-Ethylmaleimide, 2 mM Ortho-Phenanthroline. 26. Low-salt buffer: 20 mM Hepes-NaOH pH 7.9, 10% Glycerol, 1.5 mM MgCl2, 20 mM KCl, 0.2 mM EDTA, 1 mM PMSF, 2  μg/mL Leupeptin, 5 μg Aprotinin, 2 μg/mL Pepstatin,

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10 mM Na-butyrate, 10 mM β-glycerophosphate, 100 μM Na-orthovanadate, 5 mM N-Ethylmaleimide, 2 mM Ortho-Phenanthroline. 27. High-salt buffer: 20 mM Hepes-NaOH pH 7.9, 10% Glycerol, 1.5 mM MgCl2, 1.2 M KCl, 0.2 mM EDTA, 1 mM PMSF, 2  μg/mL Leupeptin, 5 μg Aprotinin, 2 μg/mL Pepstatin, 10 mM Na-butyrate, 10 mM β-glycerophosphate, 100 μM Na-orthovanadate, 5 mM N-Ethylmaleimide, 2 mM Ortho-Phenanthroline. 28. Protease inhibitor cocktail (PIC, Sigma). 29. Sepharose CL-6B resin (Sigma). 30. Poly-prep® chromatography column (Bio-Rad). 31. Anti-FLAG® M2 Affinity Gel (Sigma #2220). 32. Glycine-HCl buffer: 100 mM Glycine-HCl pH 3.5. 33. Wash buffer 1: 20 mM Hepes-NaOH pH 7.9, 10% glycerol, 300 mM KCl, 0.1% Tween 20, 1 mM DTT, 1 mM PMSF, 2  μg/mL Leupeptin, 5 μg Aprotinin, 2 μg/mL Pepstatin. 10 mM Na-butyrate, 10 mM β-glycerophosphate, 100 μM Na-orthovanadate, 5 mM N-Ethylmaleimide, 2 mM Ortho-Phenanthroline. 34. Wash buffer 2: 20 mM Hepes-NaOH pH 7.9, 10% glycerol, 150 mM KCl, 0.1% Tween 20, 1 mM DTT, 1 mM PMSF, 2  μg/mL Leupeptin, 5 μg Aprotinin, 2 μg/mL Pepstatin. 10 mM Na-butyrate, 10 mM β-glycerophosphate, 100 μM Na-orthovanadate, 5 mM N-Ethylmaleimide, 2 mM Ortho-Phenanthroline. 35. 3×FLAG peptide. 36. Micro bio-spin™ columns (Bio-Rad). 37. Protein low bond microcentrifuge tube. 38. Strep-Tactin® #2-1201-010).

Sepharose®

50%

suspension

(IBA

39. D-biotin (Thermo Fisher). 40. Ultracentrifuge with 70Ti rotor. 41. Benchtop centrifuge with refrigeration. 42. Centrifuge with JA-10, JA-14, and JA-20 rotors (or equivalent). 2.2  Histone/Lysine Acetyltransferase (HAT) Assay

1. 5× HAT buffer: 250 mM Tris–HCl pH 8.0, 25% glycerol, 0.5 mM EDTA, 5 mM DTT, 5 mM PMSF. 2. Carbonate buffer: 0.5 M Na2CO3-NaHCO3 pH 9.2. 3. Na-butyrate. 4. [3H] Acetyl CoA (4.7 Ci/mmol).

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5. Whatman paper P81. 6. EN3HANCE (PerkinElmer). 7. 4× protein loading buffer: 0.25 M Tris–HCl pH 6.8, 8% SDS, 40% glycerol, 8% β-mercaptoethanol, 0.02% bromophenol blue. 8. Transfer buffer: 20% methanol 20 mM Tris–HCl, 25 mM glycine, 0.375% SDS. 9. Coomassie staining solution: 40% methanol, 7% acetic acid, 0.1% (W/V) Coomassie Brilliant Blue R-250. 10. Coomassie destaining solution: 40% methanol, 7% acetic acid. 11. Gel dryer. 12. Scintillation Counter. 2.3  Detection of Human Acetylated Lysine Peptide by Mass Spectrometry

1. ABC buffer: 50 mM Ammonium BiCarbonate, pH 8.5 (use LC-MS Grade water for all buffers employed for mass spectrometry analysis). 2. 50% formic acid in H2O. 3. LC-MS grade water. 4. 20 mM Tris–HCl pH 8, 2 mM CaCl2. 5. SpeedVac. 6. KAc IP buffer (50 mM MOPS pH 7.4, 10 mM NaPO4, 50 mM NaCl). 7. Anti-KAc beads (ImmuneChem, product #ICP0388). 8. Trifluoroacetic acid (TFA). 9. Desalting Buffer A (0.5% Formic acid in H2O). 10. Desalting Buffer B (0.5% Formic acid in 80% acetonitrile and 19.5% H2O). 11. C18 StageTip (ThermoFisher, catalogue #87782).

3  Method 3.1  Preparation of Native Lysine Acetyltransferase (KAT) Complexes from 3 L Cultures 3.1.1  Generate Isogenic Cell Lines Expressing TAP-Tagged cDNAs

K562 cells stably expressing near physiological levels of 3×FLAG-­ Twin-­ Strep-tagged protein was described previously [12, 13]. cDNA of the gene of interest is cloned into AAVS1_Puro_ PGK1_3×FLAG_Twin_Strep (Addgene #68375). The cassette is integrated at the AAVS1 locus after DSB induction and recombination targeted by co-transfection with a vector expressing the gRNA targeting AAVS1 and the Cas9 nuclease. An AAVS1-­ targeting ZFN has also been previously used [14]. The expression vector was modified to remove the FLAG epitopes.

Analysis of Lysine Acetylation in Vitro and in Vivo

1. Grow K562 cells in RPMI 1640 [−] 10% NBCS and 1% GlutamaxTM.

l

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Glutamine containing

2. Transfect 0.2 × 106 K562 cells with 400 ng CRISPR (gRNA/ Cas9) expression vector and 4 μg of donor constructs using lipofectamine 2000 in 6-well plate. 3. 48 h later transfer the cells into 100 mm plate and select the transfected cells with 0.5 μg/mL puromycin. 4. The next day, harvest the cell by centrifugation at 90 × g for 3 min at room temperature. Remove the media and resuspend the cells in 1 mL RPMI (see Note 1). 5. Inoculate 25 mL 1.3% methylcellulose RPMI (see Note 2) complemented with 0.5 μg/mL puromycin with 200–400 μL cells in 100 mm plate. 4–5 days later, pick several colonies derived from single cell expansion from the methylcellulose-­ RPMI plate and transfer each colony to a well of 96-well plate (see Note 3). Expand the clones and analyze the expression of the protein by classic SDS-PAGE followed by western blot with anti-FLAG antibody. 6. Expand the selected clone of K562 cell expressing the protein of your interest in T175 flask with 90 mL of RPMI media completed with 10% NBCS and 0.5 μg/mL puromycin till you reach 1 million cells/mL (routinely we grow 7× T175 flaks for a total of 600 × 106 cells at this step). 7. Pellet the cells by centrifugation at 90 × g for 3 min at room temperature and pool the pellets. 8. Inoculate 1.5–3 L RPMI 1640 [+] GlutamaxTM [+] 25 mM Hepes media completed with 10% NBCS (w/o puromycin) in a 3-L capacity spinner. Count the cells everyday until it reaches 0.8 × 106 cells/mL. 3.1.2  Preparation of Nuclear Extract (NE)

All the steps of the following protocol should be carried out at 0 to 4 °C, preferably in a cold room, unless otherwise indicated. Use precooled buffers and equipment. Buffers must be prepared fresh with inhibitors on the day or the day before without the inhibitors and add the inhibitors freshly. 1. Collect the cell by transferring the contents of spinner flask into 500 mL tubes, centrifuge at 900 × g (2000 rpm in a JA-10 rotor) for 10 min at 4 °C. 2. Carefully decant the supernatant and Pool the pellets in 50 mL cold PBS. 3. Centrifuge at 1900 × g for 10 min at 4 °C. Measure the pellet volume. 4. Rapidly resuspend the cells in 4× pellet volume of ice-cold hypotonic buffer.

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5. Centrifuge at 1900 × g for 5 min at 4 °C. Remove the supernatant. 6. Resuspend the cells in 2× pellet volume of ice-cold hypotonic buffer, incubate on ice for 10 min. 7. Transfer the lysate to 40 mL homogenizer and grind with 15 strokes using Dounce B pestle. 8. Centrifuge at 3300 × g for 15 min at 4 °C. Transfer the supernatant (cytoplasmic fraction) into a new tube, flash freeze in liquid nitrogen and store at −80 °C until further analysis. 9. Measure the pack nuclear volume (PNV), resuspend the pellet (nuclear fraction) in ½ of the PNV volume with ice-cold low-­ salt buffer. Add ½ PNV high-salt buffer dropwise while vortexing at low speed. 10. Transfer the lysate to a 15 mL homogenizer and grind with 2 strokes using Dounce B pestle. 11. Incubate on ice for 30 min. 12. Transfer the lysate to 50 mL round bottom centrifuge tube and centrifuge at 25,000 × g in a 70Ti rotor (or equivalent) for 1 h at 4 °C. 13. At this step the supernatant is the nuclear extract and the gummy pellet is the chromatin fraction. Transfer the supernatant (nuclear extract) to a new 15 mL tube and proceed directly to TAP purification OR flash freeze the nuclear extract in liquid nitrogen and store it at −80 °C until TAP purification can be performed. 3.1.3  TAP Purification of KAT

All the steps of the following protocol should be carried out at 0 to 4 °C, preferably in a cold room, unless otherwise indicated. Use precooled buffers and equipment. Buffers must be prepared fresh with inhibitors on the day or the day before without the inhibitors and add the inhibitors freshly. The volume of the beads is adjustable (see Note 4). 1. Prechill the rotor, centrifuge tubes, falcon tubes, and buffers. 2. Thaw the nuclear extract on ice. 3. Determine the volume of nuclear extract, add 0.1% tween-20 to it, and mix gently by inverting. 4. Add the protease inhibitor cocktail (PIC, 1:500 dilution) and 10 mM sodium butyrate (a deacetylase inhibitor). Phosphatase inhibitors can also be added in this step if required. 5. Centrifuge at 25,000 × g in a Beckman 70Ti rotor (or equivalent) for 30 min at 4 °C. 6. In order to reduce contamination with lipids at the surface, immediately transfer the nuclear extract to prechilled 15 mL tubes.

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7. Prepare Sepharose CL-6B resin (80% slurry) (see Note 5).

(a) For each sample, add 250 μL Sepharose CL-6B resin (80% slurry) to poly-prep column.



(b) Wash the column with 5 mL PBS. Let the buffer pass through the column by gravity.



(c) Equilibrate with 5 mL low-salt:high-salt (ratio 3:1, respectively). Let the buffer pass through the column by gravity.



(d) Plug the column to prevent drying.



(e) Place the columns at 4 °C.

8. Preclear the nuclear extract on the prepared CL-6B column by gravity at 4 °C. Take an aliquot of the eluate at this step. It is your input. 9. Prepare 250 μL of anti-FLAG M2 beads for 1.5–3 L of cell culture (500 μL of 50% slurry) (see Note 6).

(a) Gently mix the anti-FLAG resin and pipet into 15 mL tube.



(b) Centrifuge at 750 × g for 3 min at 4 °C. Aspirate the supernatant.



(c) To remove the unbound antibodies, wash the anti-FLAG resin with 20 × cv (cv is column volume of 100% slurry beads/purification) Glycine-HCl buffer (100 mM GlycineHCl pH 3.5). Gently resuspend the resin, avoiding bubbles, and incubate at room temperature for 1 min.



(d) Centrifuge at 750 × g for 3 min at 4 °C. Aspirate the supernatant.



(e) Wash with 40 × cv ice-cold PBS. Centrifuge at 750 × g for 3 min at 4 °C. Aspirate the supernatant.



(f) Wash once with 40 × cv low-salt:high-salt buffer (ratio 3:1, respectively). Centrifuge at 750 × g for 3 min at 4 °C. Aspirate the supernatant.

10. Add precleared nuclear extract to the equilibrated anti-FLAG M2 beads. Incubate for 2 h on a rotating wheel at 4 °C. Meanwhile, go to the step 23 and prepare the StrepTactin beads. *At this step you can proceed to acetylated lysine immunoprecipitation protocol (see Subheading 3.3). 11. Transfer the slurry to a new poly-prep column, recuperate the flowthrough by gravity, and pass it again on the same column. Take an aliquot of anti-FLAG resin flowthrough at this step. It is your anti-FLAG flowthrough sample. 12. Wash the FLAG resin as follow:

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(a) 40 × cv low-salt:high-salt buffer (ratio 3:1, respectively).



(b) 40 × cv wash buffer 1.



(c) 40 × cv wash buffer 2.



(d) Plug the column to prevent drying.

13. Add 500 μL wash buffer 2 on the resin and transfer the resin to 1.5 mL tube. 14. Centrifuge at 750 × g for 3 min at 4 °C. Aspirate the supernatant. 15. For the first FLAG elution, add 2.5 × cv FLAG elution buffer (200  μg/mL 3×FLAG peptide in wash buffer 2 containing protease inhibitors). Incubate on a rotating wheel for 1 h at 4 °C. 16. Centrifuge at 750 × g for 5 min at 4 °C. 17. To avoid beads contamination in the elute, pass the supernatant on micro bio-spin column placed into a 2 mL low protein binding tube. 18. Centrifuge at 750 × g for 1 min at 4 °C. 19. Keep the flowthrough. Take an aliquot of the first FLAG eluate. 20. For the second FLAG elution, repeat steps 15–19. Take an aliquot of the second FLAG eluate. 21. Pool the remainder of the two FLAG eluates. 22. Prepare Strep-Tactin beads 200 μL of 50% slurry per IP.

(a) Pipet the beads to 1.5 mL tube and centrifuge them at 750 × g for 1 min at 4 °C. Remove the supernatant.



(b) Wash the beads with 1 mL PBS and centrifuge them at 750 × g for 3 min at 4 °C. Aspirate the supernatant.



(c) Wash the beads with 1 mL of wash buffer 2 and centrifuge at 750 × g for 3 min at 4 °C. Aspirate the supernatant.



(d) Resuspend the beads at 50% slurry in wash buffer 2.

23. Add the FLAG eluates pool to the equilibrated Strep-Tactin beads and incubate on a rotating wheel for 2 h at 4 °C. 24. Centrifuge at 750 × g for 3 min at 4 °C. Collect the supernatant. (This is the Strep flowthrough. Flash freeze in liquid nitrogen and store at −80 °C.) 25. Wash the beads with 10 × cv wash buffer 2. Centrifuge at 750 × g for 3 min at 4 °C. Aspirate the liquid. 26. Elute in 1.25 × cv wash buffer 2 with 5 mM D-biotin (ThermoFisher; catalogue #B20656). 27. Incubate for 1 h at 4 °C on a rotating wheel. 28. As for the FLAG elution, perform a second elution with 5 mM D-biotin (steps 16 and 17) and do not pool the eluates.

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29. To avoid protein denaturation by freeze and thaw, aliquot the eluates in small volumes (20 μL), flash freeze them in liquid nitrogen and store at −80 °C. 30. Fractions are ready to be analyzed by SDS-PAGE, mass spectrometry (see Note 7) or used for enzymatic activity (e.g., HAT assay). 3.2  Lysine Acetyltransferase Assay

To measure the HAT activity of the purified complex, the transfer of radiolabeled acetyl groups from Acetyl-CoA on NH3+ groups of Lysine amino acid residues of histone can be measured. 1. Use 0.5 μg of native chromatin, recombinant nucleosomes, or core histones in 15 μL HAT buffer mix (3 μL of 5× HAT buffer, 1 mM Na butyrate, and 0.125 μCi [3H] Acetyl CoA (4.7 Ci/mmol)). 2. Adjust the total NaCl/KCl concentration final concentration to 50 mM using 100 mM KCl considering all the reaction components salt concentration (including the purified KAT complex itself). Please note that we found that salt concentration above 100 mM have inhibitory effect on acetyltransferase activity. Preincubate the reaction mix on ice for 10 min (see Note 8). 3. Add up to 3 μL of the purified KAT complex. Larger volumes are usually detrimental to the reaction. 4. Incubate the reaction mix at 30 ° C for 30–60 min. 5. To detect the total HAT activity regardless of the substrate specificity, follow the liquid assay (step 6). To detect histoneor protein-specific HAT activity, proceed to SDS-PAGE followed by fluorography (step 7). 6. For liquid HAT assay:

(a) At the end of the incubation time, spot each reaction on P81 phosphocellulose membrane separately and air-dry the membranes.



(b) Wash the membranes three times in 50 mM carbonate buffer (NaHCO3-NaCO3; pH 9.2) for 5 min.



(c) Rinse the membranes with acetone and air-dry.



(d) Place each membrane into a scintillation vial, cover with scintillation cocktail, and measure counts in scintillation counter for 30 min.

7. For histone-specific or other protein substrate HAT activity:

(a) At the end of the incubation time, stop the reaction from step 4 by adding 5 μL of 4 × Laemmli sample buffer and incubate at 95 °C for 5 min.



(b) Load the samples on an 15% SDS-polyacrylamide gel.

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3.3  Detection of Acetylated Lysine Peptides by Mass Spectrometry 3.3.1  Trypsin Digestion of the Purified Complex “On Beads”



(c) Migrate the proteins for 2 h at 160 V in Tris–Glycine buffer.



(d) To verify equivalent loading between the reactions, stain the gel in Coomassie staining buffer for 30 min (or more) ­followed by incubation in destaining solution until histones’ bands become visible.



(e) Incubate the gel in EN3HANCE autoradiography enhancer for 30 min.



(f) Wash the gel in H2O for 30 min.



(g) Dry the gel at 60 °C for 2 h on a gel dryer.



(h) Expose the gel on film for 2–3 days at −80 °C.

Continue from step 10 of TAP purification of KAT section (Subheading 3.1.3). 1. Wash the beads 3 × with 1 mL of 50 mM ABC (adjust the pH of the ABC buffer to 8.0 using NH4OH). 2. Resuspend the beads in 200 μL of 50 mM ABC and add 2 μg of Trypsin (resuspended in 20 mM Tris–HCl pH 8) and incubate O/N on a rotating wheel at 37 °C. 3. Next day, add another 2 μg of Trypsin and incubate for a further 2–4 h. 4. Centrifuge the beads at 1000 × g for 2 min at 4 °C. Transfer the supernatant (peptides) to a new low protein binding tube. 5. Wash the beads 2× in 200 μL of LC-MS grade water. Centrifuge at 1000 × g for 2 min at 4 °C each time. Collect the supernatants and pool with previously collected supernatant. 6. To inhibit the trypsin activity, add formic acid from a 50% stock solution to the peptide solution to a final concentration of 2%. 7. Centrifuge at 10,000 × g for 10 min at 4 °C, transfer supernatant to a new low protein binding tube (be sure not to collect any beads. To do so, leave 5–15 μL in the bottom). 8. Dry the samples in a SpeedVac. 9. Resuspend the peptides in 48 μL of 5% formic acid. Centrifuge at 12,000 × g for 5 min. Transfer 6 μL to a new tube to be able to analyze 5 μL by LC-MS/MS. 10. Dry the remainder of sample using a SpeedVac and store it at −80 °C. 11. The sample is now ready for affinity purification of acetylated-­ lysine peptides.

Analysis of Lysine Acetylation in Vitro and in Vivo 3.3.2  Small-Scale Affinity Purification of Acetylated-Lysine Peptides

69

1. Resuspend the dry peptides in 400 μL of KAc IP buffer by vortexing it for 5–10 s. Store the samples on ice while the affinity beads are washed. 2. For each sample to be processed, use 50 μL of anti-KAc beads. Aliquot the beads in 1.7 mL tube and wash three times with 1 mL of ice-cold KAc IP buffer. 3. Add the peptides to the beads and incubated on a nutator at 4 °C overnight. 4. The next morning, centrifuge the samples at 750 × g for 2 min and transfer the unbound fraction to a fresh tube. This is the flowthrough fraction. 5. Wash the beads as follow:

(a) Wash with 1 mL KAc IP buffer and centrifuge at 750 × g for 2 min at 4 °C. Aspirate the supernatant.



(b) Wash with 1 mL 20 mM Tris–HCl pH 8, 2 mM CaCl2 and centrifuge at 750 × g for 2 min at 4 °C. Aspirate the supernatant.

6. Elute the peptides with 1 mL 0.5% TFA by incubating at room temperature for 20 min on a nutator. 7. Centrifuge the samples at 750 × g for 2 min. Transfer the supernatant to a fresh tube and SpeedVac to dryness. This is the bound fraction. 8. The samples are ready for mass spectrometry analysis once they have been desalted with a C18 StageTip. 3.3.3  Cleanup and Concentration of the Affinity-Purified K-Ac Peptides Using C18-StageTips

1. Conditioning the C18-StageTips:

(a) Wet the disks by passing 20 μL of 100% methanol through the StageTip. Centrifuge at 750 × g for 2 min. (b) Add 20  μL desalting buffer B (0.5% Formic acid in 80% acetonitrile and 19.5% H2O) to the StageTip. Centrifuge at 750 × g for 2 min.





(c) Add 20  μL desalting buffer A (0.5% Formic acid in 99.5% H2O) to the StageTip. Centrifuge at 2250 × g for 4 min.

2. Load the desired acidified sample volume by the addition of sufficient formic acid from a 50% stock solution onto the sample (final formic acid concentration should be 0.5–2% to obtain a pH of 2.5–3). If sample is currently dry, add sufficient buffer A to fully resuspend the sample (e.g., 20 μL). Centrifuge the StageTip at 750 × g for 2 min. 3. Reload flowthrough onto the StageTip to allow complete peptide capture. Centrifuge at 750 × g for 2 min. 4. Wash the StageTip twice by adding 20 μL desalting buffer A and centrifuge at 2250 × g for 4 min (see Note 9).

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5. Elute the desalted samples:

(a) Place Stage Tip in a new tube. Keep all eluate moving forward.



(b) To elute the peptides, apply 20 μL desalting buffer B to the StageTip three times. Collect the eluate by centrifugation at 750 × g for 2 min.



(c) Combine all the eluted fraction in a single fresh Eppendorf tube.

6. Speedvac sample to dry and store at −80 °C until LC–MS/MS analysis. 3.4  Expression and Purification of Recombinant Protein Acetylated at Single Sites

3.4.1  Materials Expression and Purification of PCNA Acetylated at Single Sites (K20)

As example, recombinant yeast PCNA (Pol30) acetylated at single site was expressed in Escherichia coli BL21 Codon-Plus (DE3)-RP with the protocol described by Chin lab [15, 17]. BL21 cells are transformed with pCDF PylT and pBK AcKRS-3 plasmids previously described [15]. The transformed cells were treated with nicotinamide and acetyl-lysine, and protein production was then induced by IPTG. His-tagged/acetylated at single site PCNA is then purified on Ni-NTA resin according to standard procedures. 1. BL21 (DE3) Competent E. coli. 2. pBK AcKRS-3 (KanR) expressing the aminoacyl-tRNA synthetase. 3. pCDF PylT plasmid carrying the ORF for the protein of interest with amber codons at the desired positions. 4. LB agar plate. 5. Kanamycin (50 μg/mL), Spectinomycin (100 μg/mL), and Chloramphenicol (34 μg/mL). 6. 100 mM MgCl2 sterile solution. 7. 100 mM CaCl2 sterile solution. 8. 3000 MWCO dialysis tubing. 9. IGEPAL® CA-630. 10. Lysis Buffer: 20 mM Tris–HCl pH 7.5, 400 mM NaCl, 1 mM PMSF. 11. Wash Buffer 1: 20 mM Tris–HCl pH 7.5, 200 mM NaCl. 12. Wash Buffer 2: 10 mM Tris–HCl pH 7.5, 500 mM NaCl, 0.1% IGEPAL, 20 mM imidazole pH 7. 13. Elution buffer: 10 mM Tris–HCl pH 7.5, 100 mM NaCl, 0.1% IGEPAL, 500 mM imidazole pH 7.0. 14. Dialysis buffer: 10 mM Tris–HCl pH 7.5, 100 mM NaCl, 0.1% IGEPAL, 10% glycerol, 0.5 mM DTT. 15. 1 M Nicotinamide.

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16. 1 M acetyl-lysine. 17. 1 M imidazole pH 7.0. 3.4.2  Methods Expression and Purification of PCNA Acetylated at Single Sites (K20)

1. Transform the BL21(DE3) competent cells with the pBK AcKRS3 to express the aminoacyl-tRNA synthetase [15]. Plate on LB agar supplemented with 50 μg/mL Kanamycin. Grow at 37 °C overnight. 2. Inoculate a single colony in 4 mL LB supplemented with kanamycin (50 μg/mL). Grow at 37 °C in a shaking incubator until the absorbance at 600 nm is 0.4. 3. Incubate the cells on ice for 5 min. 4. Harvest the cell by centrifugation at 4000 × g for 10 min at 4 °C. Remove the supernatant, and resuspend the cell pellet in 500 μL ice-cold 100 mM MgCl2 sterile solution. 5. Centrifuge the cell suspension at 4000 × g for 10 min at 4 °C. Remove the supernatant and resuspend the pellet in 100 μL of ice-cold 100 mM CaCl2 sterile solution. 6. Once the pellet is well resuspended, add 1 mL of ice-cold 100 mM CaCl2 sterile solution. 7. Incubate for 1 h on ice. 8. Centrifuge the cell suspension at 4000 × g for 10 min at 4 °C and resuspend the pellet in 125 μL of ice-cold 100 mM CaCl2 sterile solution. Add 15% sterile glycerol if stored at −80 °C. 9. Use 50 μL of theses transformed cells (BL21 containing pBK AcKS3) to transform with the pCDF PylT pol30 WT and pol30 K20amber, respectively. 10. Plate on LB agar supplemented with 100 μg/mL spectinomycin and 50 μg/mL kanamycin. Grow at 37 °C overnight. 11. Inoculate a single colony in 3 mL LB supplemented with 50  μg/mL Kanamycin, 100 μg/mL Spectinomycin, and 34 μg/mL Chloramphenicol. Incubate at 37 °C in a shaking incubator overnight (see Note 10). 12. Inoculate 3 mL of the overnight culture in pre-warmed 500 mL LB (this volume can be adjusted) with 50 μg/mL Kanamycin, 100 μg/mL Spectinomycin, and 34 μg/mL Chloramphenicol. Incubate at 37 °C until the absorbance at 600 nm is 0.4 (an OD more than 0.6 may affect the induction). 13. Add 20 mM nicotinamide and 10 mM acetyl-lysine. Incubate for 30 min at 37 °C in a shaking incubator. 14. Add 1 mM IPTG to induce the production of Pol30. 15. Incubate the flask at 16 °C in a shaking incubator overnight. 16. Harvest the cells by centrifugation at 4500 × g for 15 min at 4 °C.

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17. Resuspend the cell pellet in 50 mL wash buffer 1. Centrifuge the cell suspension at 4500 × g for 15 min at 4 °C. 18. Resuspend the cell pellet in 30 mL lysis buffer. 19. Add 0.5 mg/mL lysozyme and incubate on a rotator at room temperature for 30 min. 20. Add 0.5% IGEPAL. 21. Sonicate on ice, 10 s ON 30 s OFF with 55% amplitude for four to six times using a Branson sonicator model 450. Mix the cell lysates well between each sonication. 22. Add 2% TritonX-100 and incubate on a rotator at room temperature for 5 min. Centrifuge the lysate at 48,000 × g (20,000 rpm in a JA-20 rotor) for 30 min at 4 °C. 23. Prepare Ni-NTA agarose beads:

(a) Use the 1 mL of Ni-NTA beads for 1 L culture. (The amount of beads can be adapted according to culture volume.)



(b) Wash the beads three times with 10 mL lysis buffer containing detergents.

24. Transfer the Supernatant to a new tube. Add imidazole to the tube to 20 mM final. 25. Add the Ni-NTA agarose beads. Incubate for 1 h at 4 °C with gentle agitation. 26. Wash the beads five times for 5 min in 10 mL wash buffer 2 at 4 °C with gentle agitation. Centrifuge at 750 × g for 2 min in between the washes and discard the supernatant. 27. Add 1 mL elution buffer to the beads. Incubate for 30 min at 4 °C with gentle agitation. Repeat this step and pool the elutes in a single tube. 28. Using 3 kDa cutoff dialysis bags, dialyze the supernatant in 1 L of dialysis buffer at 4 °C for 2 h followed by dialysis O/N in 1 L fresh dialysis buffer at 4 °C to remove imidazole and add glycerol. 29. At the end of dialysis aliquot, flash freeze in liquid nitrogen and store at −80 °C. 30. Analyze the elutes by SDS-PAGE following by western blot. 31. Crystallization and crystal structure determination of acetylated PCNA at lysine 20 were performed following standard procedures [17]. 3.5  Detection of Yeast Acetylated Protein by Immunop-­ recipitation

This protocol was designed to use with yeast strains expression GST-fusion proteins under the inducible GAL promoter (modified from [18]). It can be easily adapted to any protein for which a tag or an antibody is available.

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1. SD-URA (dextrose/raffinose): Yeast nitrogen base w/o amino acids (Difco), Hopkins Synthetic complete supplement (SCSM) mix drop out: -URA (Formedium), Glucose (Sigma) 2% final or Raffinose (Sigma) 2% final. 2. d-(+)-Galactose (Sigma). 3. Lysis Buffer (100 mM Tris–HCl pH 7.5, 50 mM NaCl, 1 mM EDTA, 1 mM EGTA, 10% glycerol, 1% Tween-20, 50 mM NaF, 1 mM DTT, 1 mM PMSF, 5 μM pepstatin A, 1 μM MG132, 15 μM TSA, 50 mM Nicotinamide, 50 mM Na Butyrate, 5 μg/μL Aprotinin, 2 μg/μL Leupeptin). 4. Wash buffer (50 mM HEPES pH 7.0 and 100 mM NaCl). 5. Bead beater with 80 mL chamber (Biospec products). 6. Glass beads (0.5 mm). 7. Acetylated-Lysine Mouse mAb (Ac-K-103) (Cell signaling 9681). 8. Protein-A sepharose. 9. Wash buffer (10 mM Hepes pH 7.5, 350 mM NaCl, 0.1% IGEPAL). 10. 4× protein loading buffer: 0.25 M Tris–HCl pH 6.8, 8% SDS, 40% glycerol, 8% β-mercaptoethanol, 0.02% bromophenol blue. 11. Anti-GST antibody.

3.5.2  Method

1. Inoculate GST-Gal yeast strain in 2 mL SD-URA (dextrose) overnight OR inoculate 50–100 mL SD-URA (dextrose) overnight and the next morning proceed directly to step 3. 2. Inoculate 100 mL SD-URA (dextrose) in the morning and grow at 30 °C until the absorbance at 600 nm is ~2–3. 3. Wash cells in H2O and inoculate a 500 mL culture in SD-URA (2% raffinose) at the absorbance at 600 nm is 0.1. 4. Grow the cells at 30 °C in a shaking incubator until the absorbance at 600 nm is 0.8. 5. Add galactose to final concentration 2% and allow to express the protein for 4 h. 6. Harvest cells by centrifugation at 4500 × g at 4 °C (5000 rpm in a JA-10 rotor), wash in 50 mL of 10 mM Tris–HCl pH 8, 150 mM NaCl and transfer to a falcon tube. Flash freeze in liquid nitrogen and store at −80 °C until ready to proceed to step 7. 7. Resuspend the cell pellet in 8 mL lysis buffer. 8. Disrupt the cells using 80-mL chamber bead beater filled up with 40 mL of glass beads and lysis buffer in the cold room (setting: 20 s on- 45 s off). Confirm breakage by microscope (see Note 11).

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9. Transfer lysate and beads to a falcon tube, spin 1–2 min at 900 × g at 4 °C and keep supernatant. 10. Centrifuge at 10,000 × g for 30 min at 4 °C. 11. Ultracentrifuge 1 h at 40,000 × g at 4 °C. 12. While sample is centrifuging, prepare protein A-sepharose conjugated with anti-KAc antibodies:

(a) Wash 60  μL protein-A sepharose beads (50% slurry) three times in 1 mL lysis buffer. Centrifuge at 750 × g for 2 min at 4 °C and in between the washes, aspirate the supernatant carefully.



(b) Add 6  μL of mouse monoclonal anti-acetyl lysine antibody and incubate on rotating wheel at 4 °C for 1 h.



(c) Wash the antibody/protein A-sepharose beads conjugate mixture twice in 1 mL lysis buffer. Centrifuge at 750 × g for 2 min at 4 °C in between the washes and aspirate the supernatant carefully.

13. Keep 1/50 of lysate volume as input (~200 μL). Add 4× protein loading buffer and boil for 5 min. Store at −20 °C until its analysis. 14. Add the conjugated antibody/protein A-sepharose beads to 1/10 of lysate volume. 15. Incubate on a rotating wheel for 3 h at 4 °C. 16. Wash four times for 10 min on a rotating wheel at 4 °C in 500 μL wash buffer. 17. Add 20 μL Add 4× protein loading buffer to IP beads and boil for 5 min. 18. Run inputs and IP samples on SDS-PAGE followed by Western blot with anti-GST antibody (or for other tags/antibodies).

4  Notes 1. Subculture the rest of cells in a new plate and continue selection with puromycin for 1 week. Freeze 1 × 106 cells per vial in 1 mL 20% NBCS, 10% DMSO in RPMI and keep as the “pool.” 2. To prepare 1.3% methylcellulose-RPMI medium:

(a) Precool 1/2 total volume ddH2O on ice (125 mL for total 250 mL).



(b) Heat 1/2 total volume ddH2O (125 mL) in a 500 mL flask with stir bar agitation until temperature reaches 80 °C. You will see the water starting to evaporate, do not boil.

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(c) Add 6.50 g of methylcellulose gradually to avoid clumping. Mix solution with stir bar until methylcellulose is uniform. Methylcellulose will not dissolve at high temperatures.



(d) Add the 125 mL precooled ddH2O. Mix with stir bar at 4 °C (cold room). Once methylcellulose is completely dissolved, the solution becomes very viscous.



(e) Autoclave with stir bar (the solution usually evaporates to ~215 mL after autoclaving).



(f) Incubate in cold room at 4 °C overnight while stirring slowly.



(g) Add the 250 mL RPMI 1640 2×, FBS 70 mL, Glutamax 5 mL, Pen Strep 5.5 mL.



(h) Shake vigorously to mix. Let the medium stand at 4 °C to allow the bubbles to rise to the top.



(i) Slowly pipet 27.5 mL of methylcellulose media into 50 mL falcon tubes (this should yield about 25 mL of solution since some media will remain in the pipet). Store at −20 °C.



(j) Thaw at 4 °C the day before use.

3. To pick the colonies, put an inverted light microscope under cell culture hood and carefully peak a single colony with a 20–200 μL pipette. In advance, prepare a 96-well plate with 200 μL/well RPMI supplemented with 0.5 μg/mL puromycin and incubate at 37 °C till use. 4. TAP can be carried out with 500 mL to 6 L cell culture volume depending on the protein expression abundancy and desired analysis (e.g., analytical enzymatic assay or detection of PTMs). The aim is to purify the tagged-protein complex with minimal background contaminants. For low abundance proteins, using too much anti-Flag resin can lead to more contaminants, so cutting the amount of beads two- to fivefold may give better results in term of concentration and purity. Empirical determination of the best ratio of beads to cell number/extracts is necessary. 5. Mix the slurry well and use pipette tips with the end cut off to prevent damage to the beads or use 5 mL pipets to take the Sepharose CL-6B resin. 6. Mix the beads slurry well. Cut pipette tip 5 mm from the end. Pipette carefully as beads tend to stick to the sides of the tip. See also Note 4 for volume of beads. 7. To send for MS analysis: use sterile material, be clean. Use fresh gloves. Wash the tubes with acetonitrile. Prepare your buffers and stock inhibitors with a lot of care to minimize keratin contamination.

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8. It is important to preincubate the purified HAT complex with chromatin (or other protein) substrates before adding Acetyl-CoA. 9. The peptide-loaded, washed C18 StageTips can be stored in buffer A in the fridge or at −20 or −80 °C. We have done so in our laboratories for several weeks without compromising sensitivity of subsequent MS analysis. 10. It is important to test several colonies for the expression on a small scale before proceeding to large-scale preparation. Pick several colonies and inoculate 3 mL LB supplemented with 50  μg/mL Kanamycin, 100 μg/mL Spectinomycin, and 34 μg/mL Chloramphenicol. Incubate at 37 °C in a shaking incubator overnight. Add 20 mM nicotinamide and 10 mM acetyl-lysine. Incubate for 30 min at 37 °C in a shaking incubator. Add 1 mM IPTG to induce protein production. Incubate the flask at 16 °C in a shaking incubator overnight. Monitor the protein expression of different clones by western blot. 11. To verify the level of yeast cells disruption, take an aliquot after every two cycles and observe under the light microscope (400× magnification). Disruption level over 80% leads to protein degradation and contamination of purified protein at the end of purification.

Acknowledgments We are grateful to Yannick Doyon, Karine Jacquet, Pierre Billon, and Xue Cheng for sharing their protocols. Research in the Côté and Lambert laboratories is funded by a Discovery Grants from the Natural Sciences and Engineering Research Council of Canada (NSERC) (1304616-2017 to J.-P.L.) and a Foundation Grant from the Canadian Institutes of Health Research (CIHR) (FDN-­ 143314 to J.C.). J.-P.L. is supported by a Junior 1 salary award from the Fonds de Recherche du Québec-Santé (FRQ-S; 251747), by an Operating Grant from the Cancer Research Society (22779) and a Leader’s Opportunity Funds from the Canada Foundation for Innovation (37454). J.C. holds a Canada Research Chair (Tier 1) in Chromatin Biology and Molecular Epigenetics. References 1. Kouzarides T (2000) Acetylation: a regulatory modification to rival phosphorylation? EMBO J 19(6):1176–1179 2. Allfrey VG, Mirsky AE (1964) Structural modifications of histones and their possible role in the regulation of RNA synthesis. Science 144(3618):559

3. Pogo BG, Allfrey VG, Mirsky AE (1966) RNA synthesis and histone acetylation during the course of gene activation in lymphocytes. Proc Natl Acad Sci U S A 55(4):805–812 4. Bannister AJ, Kouzarides T (2011) Regulation of chromatin by histone modifications. Cell Res 21(3):381–395

Analysis of Lysine Acetylation in Vitro and in Vivo 5. Svinkina T et al (2015) Deep, quantitative coverage of the lysine acetylome using novel antiacetyl-lysine antibodies and an optimized proteomic workflow. Mol Cell Proteomics 14(9):2429–2440 6. Weinert BT et al (2018) Time-resolved analysis reveals rapid dynamics and broad scope of the CBP/p300 acetylome. Cell 174(1):231–244, e12 7. Hornbeck PV et al (2015) PhosphoSitePlus, 2014: mutations, PTMs and recalibrations. Nucleic Acids Res 43(Database issue): D512–D520 8. Gil J, Ramirez-Torres A, Encarnacion-Guevara S (2017) Lysine acetylation and cancer: a proteomics perspective. J Proteome 150: 297–309 9. Jones PA, Issa JP, Baylin S (2016) Targeting the cancer epigenome for therapy. Nat Rev Genet 17(10):630–641 10. Lasko LM et al (2017) Discovery of a selective catalytic p300/CBP inhibitor that targets lineagespecific tumours. Nature 550(7674):128–132 11. Fujisawa T, Filippakopoulos P (2017) Functions of bromodomain-containing proteins and their roles in homeostasis and cancer. Nat Rev Mol Cell Biol 18(4):246–262

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12. Dalvai M et al (2015) A scalable genome-­ editing-based approach for mapping multiprotein complexes in human cells. Cell Rep 13(3): 621–633 13. Doyon Y, Cote J (2016) Preparation and analysis of native chromatin-modifying complexes. Methods Enzymol 573:303–318 14. Hockemeyer D et al (2009) Efficient targeting of expressed and silent genes in human ESCs and iPSCs using zinc-finger nucleases. Nat Biotechnol 27(9):851–857 15. Neumann H et al (2009) A method for genetically installing site-specific acetylation in recombinant histones defines the effects of H3 K56 acetylation. Mol Cell 36(1):153–163 16. Neumann H, Peak-Chew SY, Chin JW (2008) Genetically encoding N(epsilon)-acetyllysine in recombinant proteins. Nat Chem Biol 4(4): 232–234 17. Billon P et al (2017) Acetylation of PCNA sliding surface by Eco1 promotes genome stability through homologous recombination. Mol Cell 65(1):78–90 18. Lin YY et al (2009) Protein acetylation microarray reveals that NuA4 controls key metabolic target regulating gluconeogenesis. Cell 136(6): 1073–1084

Chapter 6 Site-Specific Lysine Acetylation Stoichiometry Across Subcellular Compartments Anastasia J. Lindahl, Alexis J. Lawton, Josue Baeza, James A. Dowell, and John M. Denu Abstract Posttranslational modifications of proteins control many complex biological processes, including genome expression, chromatin dynamics, metabolism, and cell division through a language of chemical modifications. Improvements in mass spectrometry-based proteomics have demonstrated protein acetylation is a widespread and dynamic modification in the cell; however, many questions remain on the regulation and downstream effects, and an assessment of the overall acetylation stoichiometry is needed. In this chapter, we describe the determination of acetylation stoichiometry using data-independent acquisition mass spectrometry to expand the number of acetylation sites quantified. However, the increased depth of data-­independent acquisition is limited by the spectral library used to deconvolute fragmentation spectra. We describe a powerful approach of subcellular fractionation in conjunction with offline prefractionation to increase the depth of the spectral library. This deep interrogation of subcellular compartments provides essential insights into the compartment-specific regulation and downstream functions of protein acetylation. Key words Protein acetylation, Acetylation, Mitochondria, Chromatin, Stoichiometry, Posttranslational modification, Mass spectrometry, Proteomics

1  Introduction Posttranslational modification of proteins is a mechanism of controlling many complex biological processes, including genome expression, chromatin dynamics, metabolism, and cell division. Reversible phosphorylation was the first modification to be well studied and demonstrated to have widespread regulatory effects [1–3]. Many studies have contributed to the identification of other posttranslational modifications, including protein acetylation and more recently other protein acylations [4–16]. Protein acetylation was first characterized on histone proteins and has been well studied in the regulation of gene expression and chromatin dynamics [17–20]. More recent studies have found acetylation sites on

Robert M. Brosh, Jr. (ed.), Protein Acetylation: Methods and Protocols, Methods in Molecular Biology, vol. 1983, https://doi.org/10.1007/978-1-4939-9434-2_6, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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­ roteins in compartments throughout the cell, investigating the p functional outcomes of protein acetylation [15]. Advancements in mass spectrometry-based proteomics have led the widespread identification of protein acetylation. However for the majority of the acetylation sites identified thus far, their biological effect and their mechanism of regulation of these acetylation events has remained unknown [4–16, 21–34]. Protein acetylation is a reversible enzymatic reaction, with the addition of an acetyl group on the ε-amino group of lysine residues catalyzed by a lysine acetyltransferase and the removal catalyzed by a lysine deacetylase [35–37]. Lysine acetyltransferases (KATs) contain two families of proteins, the MYST and GNAT families as well as the metazoan-specific CBP/p300 histone acetyltransferases, which utilize acetyl-CoA as the acetyl donor [38–42]. Lysine deacetylases have four classes, separated by different mechanisms and co-substrates to perform deacetylation [43–45]. Class I, II, and IV are canonical lysine deacetylases and utilize a Zinc ion and coordinated water to hydrolyze their acetyl-lysine substrates, whereas class III are NAD+-dependent deacetylases called sirtuins, which couple the deacetylation reaction with hydrolysis of the NAD+ cofactor [44]. Studies have shown lysine acetyltransferases and deacetylases may be regulated by alterations in cellular metabolism. These enzymes are sensitive to changes in concentration of acetyl-CoA and NAD+, the cellular metabolites that are used as co-substrates, and enzymatic activity may be regulated by altering the availability of co-substrates [9, 10, 46–55]. Studies investigating reversible protein acetylation have explored protein acetylation in the context of metabolism and the response to changes in concentration of the acetyl donor acetyl-­ CoA [55–57]. Particularly, the metabolic enzyme acetyl-CoA synthetase in both bacteria and mammals was found to be acetylated on a single conserved lysine residue within its active site, rendering the enzyme inactive. The enzyme is restored to full activity through deacetylation by the sirtuin proteins, a family of conserved proteins which coordinate many pathways within metabolism [58–60]. Acetylation also plays an important regulatory role in the nucleus, mitochondria, and endoplasmic reticulum, influencing protein-­ DNA interactions of histones and DNA to promote transcription, metabolic activity and flux within mitochondria, and protein folding and secretion in the endoplasmic reticulum [17, 21, 25, 61, 62]. Proteins within the nucleus and the cytosol have been characterized as substrates of histone acetyltransferases (HATs), particularly the undiscriminating enzymes CBP and p300, which themselves are regulated by reversible acetylation [63–67]. Similarly to kinase signaling cascades regulated by phosphorylation, the posttranslational regulation, including the auto-­ acetylation, of histone-modifying complexes regulates their activity on histones creating signal amplification in the nucleus [68, 69].

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Alternatively, within the mitochondria, no lysine acetyltransferase has been identified, with active investigation into the roles of nonenzymatic acetylation [70–74]. Currently, acetylation of proteins has been shown to regulate protein-protein interactions, protein stability, localization, and enzymatic activity. The most well-known example of protein acetylation is histone acetylation, regulating gene expression and chromatin dynamics and considered to be an epigenetic mark [17]. Histone acetylation can occur on the same histone tail, different tails within the same nucleosome, or adjacent nucleosomes, giving rise to a highly complex set of substrates for recognition by histone-­ modifying enzymes and other chromatin-associated proteins [75– 78]. Histone acetylation regulates gene expression through a variety of mechanisms including increased specificity, affinity, localization, or release of inhibition, and leads to upregulation of gene expression [76, 79–82]. Beyond histone acetylation, the function of lysine acetylation has been characterized for transcription factors, metabolic enzymes, chaperone proteins, and cytoskeletal proteins [9, 10, 83–88]. However, recent studies enabled by advances in mass spectrometry-based proteomics indicate lysine acetylation occurs broadly within cells, raising questions about function and regulatory mechanisms controlling lysine acetylation throughout the cell [4–16]. Initial mass spectrometry-based proteomics results increased the number of identified acetylation sites; however, such studies fell short of being able to comprehensively quantify stoichiometry of acetylation in an entire proteome. Rather than reporting relative fold change, we recently reported a data-dependent acquisition (DDA) method to quantify acetylation stoichiometries, which provides additional information for interpreting biological significance [24]. An example of the biological importance of understanding stoichiometry comes from the case of mitochondrial acetylation. Relative fold-change of acetylation stoichiometry characterized in response to alterations in diet were found to be large, resulting in expectations of high acetylation stoichiometry [21, 22]. However, stoichiometry measurements suggested the majority of acetylation sites within the mitochondria had very low stoichiometry with a median occupancy of ≤2% stoichiometry, with a small subset of sites having high stoichiometry [74, 89]. This more in-depth information given by stoichiometry assists with interpretation of biological function, particularly the differences between high and low stoichiometry when considering how acetylation results in loss-of-function or gain-of-function. Furthermore, the observation of low stoichiometry broadly in the mitochondria raised the question whether acetylation acts independently or acts in an additive fashion across many proteins in a pathway [89]. Data-dependent methods used in mass spectrometry-based proteomics are stochastic in nature. The selection of a peptide for MS/MS

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fragmentation is determined by its overall abundance during an MS run [90]. In contrast, data-independent acquisition (DIA) is a method that continuously collects fragmentation spectra from all peptides using predefined sequential mass-to-charge windows, thus increasing the quantification of light and heavy acetyl containing peptides captured in our analysis [91, 92]. This analysis addresses the limitation of missing data stemming from stochastic data collection when using DDA. The DIA approach results in the increase in the identification and quantitation of lysine stoichiometry by increasing the sample depth to quantifying both non-­isotopically and isotopically labeled peptides. Performing DIA analysis requires a spectral library of MS2 fragmentation spectra from DDA analysis to deconvolute the DIA MS2 spectra for identification and quantification. To increase depth of peptide coverage within the spectral library, we generated libraries utilizing orthogonal separation approaches, previously shown to increase sampling depth of DDA analysis [93–95]. In this chapter, we describe methods of subcellular organelle enrichment and chemical labeling of lysine residues coupled with dataindependent acquisition mass spectrometry to quantify site-specific lysine acetylation stoichiometry.

2  Methods 2.1  Methods for Sample Preparation and Subcellular Fractionation 2.1.1  General Experimental Design

2.2  Methods for Cell Culture and Serum Stimulation

Our DIA approach results in the increase in the identification and quantitation of lysine stoichiometry over other shotgun-based methods of quantifying acetylation stoichiometry. Performing DIA analysis requires a spectral library of MS2 fragmentation spectra from DDA analysis to deconvolute MS2 spectra. Here we detail generation of spectral libraries from analysis of a non-isotopically labeled DDA analysis with in silico generation of the heavy acetyl-­ lysine label. This approach creates a library with all possible isotopically labeled peptide combinations, many of which may be too low in abundance and to generate experimental peptide fragmentation spectra. Utilizing the experimentally produced library with in silico generation of heavy deuterated label, we quantify both the light endogenous non-isotopic acetylation and the heavy deuterium-­ labeled acetylation, derived from the in vitro chemical labeling reaction, to calculate site-specific stoichiometry across the proteome. Protein posttranslational modifications, such as acetylation, provide cells with an ability to rapidly modulate the activities of existing proteins within the cells, rather than through the slower processes of protein synthesis and degradation [96, 97]. To adequately capture the relatively rapid process of dynamic acetylation changes in response to stimuli in cell culture, time points such as 0, 1, 2, and 4 h will be examined. These times are longer than the

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rapid metabolic response; however, they were shorter than previously experimentally determined average protein half-life, ideally capturing dynamic changes of protein acetylation through active acetylation and deacetylation rather than passive changes through protein synthesis or turnover [98]. To examine the dynamic changes in acetylation stoichiometry, we utilized a time course to quantify the alterations of acetylation over time in response to growth factor stimulation of MCF7 cells in tissue culture. Generation of samples for the spectral library requires at least 50 μg of protein for offline prefractionation of labeled and digested peptides and subsequent LC-MS/MS analysis. As demonstrated in Fig.  1, utilizing subcellular fractionation, focusing on chromatin protein (shown in green) and offline prefractionation (shown in light blue) to create a spectral library increased the depth of protein, peptide, and acetyl site coverage as shown by the increased identifications observed in each of these categories. Furthermore, generating a spectral library with subcellular fractionation in combination with offline prefractionation (shown in dark blue) resulted in increased depth of coverage of subcellular organelles, beyond the depth of coverage of offline prefractionation alone, with a particular increase in depth on the peptide level (Fig. 2b).

Fig. 1 Data-dependent acquisition spectral library quality and depth analysis. An analysis of the spectral libraries generated from a Single Injection Whole Cell Lysate (WCL), a Single Chromatin Isolation (utilizing subcellular fractionation without offline prefractionation), a Fractionated WCL utilizing offline prefractionation, Fractionated Chromatin Isolation utilizing offline prefractionation on a subcellular fractionated chromatin isolation, a Fractionated WCL utilizing offline prefractionation with a single chromatin isolation (utilizing subcellular fractionation without offline prefractionation), and the Fractionated Chromatin Isolation and Fractionated WCL together utilizing offline prefractionation, on both whole cell lysate and a subcellular fractionated chromatin sample to examine the effects of library preparation on the counts of (a) Acetyl Sites, (b) Acetyl Peptides, (c) Acetyl Proteins, (d) MS2 Fragments, (e) Total peptides, and (f) Total proteins

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Fig. 2 Subcellular localization depth and quality analysis of data-dependent acquisition spectral libraries. An analysis of the spectral libraries generated from a Single Injection Whole Cell Lysate (WCL), a Single Chromatin Isolation (utilizing subcellular fractionation without offline prefractionation), a Fractionated WCL utilizing offline prefractionation, Fractionated Chromatin Isolation utilizing offline prefractionation on a subcellular fractionated chromatin isolation, a Fractionated WCL utilizing offline prefractionation with a single chromatin isolation (utilizing subcellular fractionation without offline prefractionation), and the Fractionated Chromatin Isolation and Fractionated WCL together utilizing offline prefractionation, on both whole cell lysate and a subcellular fractionated chromatin sample to examine the effects of library preparation on the counts of (a) Proteins and (b) Peptides in a subcellular localization specific manner

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To perform whole cell lysate analysis, a single 10 cm plate provides sufficient material for analysis. Alternatively, to perform analysis on subcellular compartments more starting material was required, utilizing a 70–80% confluent 15 cm plate. This protocol has been readily adapted to performing analysis on alternate cell types, as well as mouse tissue such as brain and liver. For tissue utilize approximately 10–15  mg of starting material for whole cell lysate and 40–50 mg for subcellular fractionation to obtain adequate yields. 2.3  Preparing Whole Cell Lysate (Without Subcellular Fractionation)

1. Resuspend cell pellet in 200 mM Ammonium Bicarbonate with protease and HDAC inhibitors: 10  μg/mL leupeptin, 10 μg/mL aprotinin, 100 μM phenylmethanesulfonyl fluoride (PMSF), 1 mM sodium butyrate, 4 μM trichostatin A, 10 mM nicotinamide, and 1 mM DTT. 2. Lyse cells using mechanical force via sonication. Using a tip sonicator, sonicate the sample on 5 s on-off pulse for 30 s at 20% amplitude. Avoid chemical lysis when possible for mass spectrometry sample preparation. If chemical lysis is needed for a specific sample, follow-up protein isolation with a precipitation protocol such as acetone or chloroform methanol precipitation to minimize detergent contamination of downstream analyses. For tissue samples, first homogenize the tissue sample previous to tip sonication to disrupt subcellular organelles using a dounce homogenizer with 20 strokes loose pestle and 20 strokes tight pestle. Let the samples stand on ice for 10 min to complete hypotonic lysis. For tissue analysis, filter the homogenate through a cell strainer to remove debris. 3. Measure protein concentration using a Bicinchoninic Acid assay (BCA) (Invitrogen).

2.4  Isolating Subcellular Fractions from Cell Culture or Tissue Samples

1. Performing subcellular fractionation will increase coverage of acetyl sites within a compartment of interest (Fig.  2). Resuspend cell pellet in 800 μL of hypotonic lysis buffer (10 mM Tris–HCl, 10 mM NaCl, 3 mM MgCl2 at pH 7.4) in a 1 mL dounce homogenizer. Mechanically lyse cells by homogenizing samples with 20 strokes loose pestle and 20 strokes tight pestle. Let the samples stand on ice for 10 min to complete hypotonic lysis. For tissue analysis, filter the homogenate through a cell strainer to remove debris. 2. Centrifuge homogenate at 800 × g for 5 min at 4 °C to pellet crude nuclear fraction. Supernatant contains cytoplasm and mitochondria. To isolate mitochondria, centrifuge supernatant at 10,000 × g for 10 min at 4 °C which pellets the mitochondria, while the supernatant contains cytoplasmic proteins. Resuspend pellet with 500 μL buffer and repeat centrifugation.

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3. To isolate chromatin proteins from the crude nuclear fraction, wash crude nuclear fraction with 2 mL cold lysis buffer and centrifuge at 800  ×  g for 5  min at 4 °C.  Repeat wash steps again with lysis buffer and then PBS. 4. Resuspend nuclear pellet in 500 μL of nuclear lysis buffer (10 mM Tris–HCl 10 mM NaCl 3 mM MgCl2 0.2 mM EDTA at pH 7.4) with protease and histone deacetylase inhibitors added immediately before use. Incubate for 15  min on ice with vortexing every 5 min. 5. Centrifuge at 16,000 × g for 5 min. Pellet is chromatin with histones. Resuspend pellet with lysis buffer and centrifuge at 3000 × g and keep pellet. 6. To isolate chromatin proteins from the highly abundant histones proteins which dominate the proteins observed from the nucleus, resuspend chromatin pellet in 300 μL of 150 mM salt in the lysis buffer (10 mM Tris–HCl 3 mM MgCl2 pH 7.4) with protease and histone deacetylase inhibitors added immediately before use. Let stand on ice for 15 min with vortexing every 5 min. Centrifuge at 3000 × g for 5 min and keep pellet for further extraction. Supernatant can be kept for gel electrophoresis if you wish. 7. Resuspend in 300 μL 450 mM salt in the lysis buffer (10 mM Tris–HCl 3 mM MgCl2 pH 7.4) with protease and histone deacetylase inhibitors added immediately before use. Let stand on ice for 15 min with vortexing every 5 min. Centrifuge at 3000 × g for 5 min. Supernatant contains chromatin bound proteins. Keep pellet for further extraction of histones if desired, following the previously described acid extraction of histone and subsequent mass spectrometry histone posttranslational modification profiling strategies [99, 100]. 8. Perform a Bicinchoninic Acid assay (BCA) (Invitrogen) on each of the subcellular compartments of interest for downstream analysis to measure protein concentration.

3  Sample Analysis 3.1  Chemical Acetylation

Our labeling approach leads to the acetylation of all lysines, whether non-isotopically acetylation from in vivo sources or deuterium-­labeled acetylation from our in vitro labeling reaction. Due to the chemical labeling approach, all lysines are modified with an acetyl group, either a non-isotopic acetyl group which is generated in vivo or deuteriumlabeled acetylation generated from the in vitro chemical labeling reaction. Proteolytic digestion by trypsin generates chemically identical light and heavy peptides which can be directly compared to determine the level of acetylation stoichiometry. Furthermore, the use of chemical labeling over modification enrichment strategies minimizes the

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amount of bias introduced to the sample preparation. It furthers the understanding of the dynamics of acetylation changes in comparison to the state of all lysines across the whole proteome, rather than only quantifying the ratio of acetylated lysines. 3.1.1  Acetic Anhydride Labeling and Digestion of Samples for Spectral Library

1. To prepare samples for the DDA analysis to generate the peptide spectra for the spectral library, pool protein from all of the samples for analysis. For example, when analyzing dynamic acetylation over time after serum stimulation in MCF7 cells, combine equal protein amounts of each condition (time point samples) up to 50 μg. The pooled samples should total to at least 50 μg for subsequent offline prefractionation and downstream LC-MS/MS analysis of the fractions. 2. To start with the same concentration in each of the mixed samples, dry down to 10 μL using a speedvac. 3. Resuspend sample to 50 μL total volume using 8 M urea, 100 mM ammonium bicarbonate pH at 8.5 with 5 mM DTT in a 1.5 mL tube. Prepare the buffers and urea fresh for each experiment. For subsequent labeling steps avoid buffers with free amine groups that would react with the acetic anhydride, affecting the labeling efficiency. 4. To denature the proteins place samples on Thermomixer C (Eppendorf), which has a heated lid to avoid condensation at 60 °C while mixing at 1000 RPM for 30 min. 5. Add 20 mM iodoacetamide to alkylate cysteine residues and incubate at 60 °C while mixing at 1000 RPM for 30 min on the Thermomixer C (Eppendorf). 6. To chemically acetylate unmodified lysine residues, add 2 μL of non-isotopic acetic anhydride to the samples, vortex, and incubate for 30 min, 60 °C 1000 RPM on the Thermomixer C (Eppendorf). For the spectral library samples, use non-­ isotopic acetic anhydride to chemically acetylate all lysine residues with the same label. In downstream processing, heavy acetylated peptide pairs will be generated in silico. This approach creates a spectral library with heavy and light pairs of all lysine containing peptides. 7. Add ~10 μL NH4OH to bring the pH back up to ~8.0. Check the pH with a 0.5 μL spot on litmus paper and add more NH4OH as necessary. 8. For complete labeling repeat the chemical labeling from steps 6 and 7, again using non-isotopically labeled acetic anhydride. 9. After the second round of chemical labeling, incubate samples at pH 8 for 20 min at 60 °C with 1000 RPM mixing on the Thermomixer C (Eppendorf) to hydrolyze any O-acetyl esters formed during the reaction.

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10. Add 100 mM ammonium bicarbonate to bring the volume up to 200 μL to dilute the urea to a final concentration of 2 M. Trypsin digestion efficiency is reduced with urea levels above 2 M. 11. Add sequencing grade trypsin to a 1:100 ratio (trypsin:protein) and incubate from 4 h to overnight at 37 °C mixing at 350 RPM on Thermomixer C (Eppendorf). 12. Add 100 mM ammonium bicarbonate to bring the volume up to 400 μL and the final concentration down to 1 M for the subsequent gluC digestion. GluC digestion efficiency is reduced with urea levels above 1 M. 13. Add gluC in a 1:100 (gluC:protein) ratio and incubate from 4 h to overnight at 37 °C mixing at 350 RPM on Thermomixer C (Eppendorf). Additional digestion with GluC is necessary to digest labeled peptides to lengths amendable for LC-MS/ MS analysis. 14. Quench the reaction by adding ~40 μL of acetic acid. 3.1.2  Prefractionation with High pH Reverse Phase Chromatography

A major challenge to performing stoichiometric acetyl lysine analysis is the lack of an enrichment step for lysines acetylated in vivo. Antibody-based techniques increase acetyl lysine coverage by enriching for acetyl-peptides, thereby simplifying the complex mixture and increasing the sampling depth. Since our strategy does not involve an enrichment step, offline fractionation is used to reduce the proteomic complexity in order to increase the depth of coverage of acetyl lysine containing peptides. The advantage of this strategy is highlighted in Figs. 1 and 2. To evaluate the increase in the depth of coverage within a spectral library, we examined the number of proteins, peptides, and fragment ions identified using a single sample whole cell lysate (whole cell lysate without offline prefractionation), a single sample chromatin isolation (utilizing subcellular fractionation without offline prefractionation), fractionated whole cell lysate utilizing offline prefractionation, and fractionated chromatin isolation utilizing offline prefractionation on a subcellular fractionated chromatin sample. These approaches in combination, displayed in Fig. 1, demonstrate the clear advantage in the depth of the spectral library generated using subcellular fractionation and offline prefractionation in conjunction. 1. To perform the offline prefractionation, we use high pH reverse phase (HPRP) chromatography for peptide separation. Resuspend the labeled and digested peptides in ~2  mL of HPRP solvent A (100 mM Ammonium Formate pH = 10) and separate by high pH reverse phase chromatography on a Waters-C18 column (5  μm, 130  Å, 250 × 2.1  mm) with a Shimadzu LC-­20AT HPLC system with a 60 min gradient of two solvents, solvent A is 100 mM Ammonium formate, pH 10 and solvent B is ACN. The gradient is a 60 min gradient,

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starting at 2% solvent B for 30 min, and then a linear gradient from 2% to 90% solvent B with a flow rate of 400 μL/min. Fractions of 0.7 mL are collected from 30 to 72 min in the gradient, with 24 collected in total. Peptides are detected by a UV detector at 280 nm. 2. Samples are dried down using the speedvac concentrator, to concentrate the fractions until they reach a small enough volume to be combined. The following HPRP fractions are combined using a concatenated strategy to form six final fractions: (a) 1, 7, 13, 19 (b) 2, 8, 14, 20 (c) 3, 9, 15, 21 (d) 4, 10, 16, 22 (e) 5, 11, 17, 23 (f) 6, 12, 18, 24 3. These recombined 6 fractions are dried down completely and then resuspend in 100 μL of 10% acetic acid for subsequent desalting and cleanup. 3.1.3  Desalting Prior to MS Analysis

Sample preparation for MS analysis requires desalting of peptide samples as well as removal of hydrophilic molecules. We generate our own desalting tips for our peptide cleanup based on a previously published protocol [101]. The desalting tips we make have the added benefit of being cost efficient as well as having minimal sample loss. The desalting tips are made with C-18 disks (Empore) in p200 pipette tips and act as spin columns in 1.5 mL microcentrifuge tubes. 1. The desalting tip must first be activated. To activate the C-18 disks, add ~50 μL of methanol to the desalting tip and centrifuge at 1000 × g for 1 min. 2. To condition the desalting tip, add ~50 μL of the elution buffer, 0.5% acetic acid in 80% ACN in LC-MS grade H2O, to the desalting tip and centrifuge at 1000 × g for 1 min. 3. To equilibrate the desalting tip, add ~50 μL of the wash buffer, 0.5% acetic acid in LC-MS grade H2O, to the desalting tip and centrifuge at 1000 × g for 1 min. 4. Load the sample in the desalting tip and centrifuge at 1000 × g until the sample volume has run completely through the ­desalting tip. The peptides are now bound to the C-18 disks within the desalting tip. 5. Wash the sample by adding ~50 μL of the wash buffer, 0.5% acetic acid in LC-MS grade H2O, to the desalting tip and centrifuge at 1000 × g for 1 min.

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6. Switch the desalting tip to a new tube and elute the peptides by adding ~50 μL of the elution buffer, 0.5% acetic acid in 80% ACN in LC-MS grade H2O, to the desalting tip and centrifuge at 1000 × g for 1 min. 7. Dry down the sample using the speedvac. 8. Prepare the samples for MS analysis by resuspending in the sample diluent of 2% ACN, 0.1% formic acid in LC-MS grade H2O. For analysis using Spectronaut software, add the internal retention time standards required, the Biognosys iRT kit, as instructed [102]. 3.1.4  LC-MS/MS Data-Dependent Acquisition (DDA) Analysis of Spectral Library Samples

Our method to analyze site-specific lysine acetylation by mass spectrometry-­ based proteomics utilizes a Thermo Q-Exactive Orbitrap mass spectrometer coupled to a Dionex Ultimate 3000 RSLC nano UPLC with a Waters Atlantis reverse phase column (100 μm × 150 mm). The mobile phases contain 0.1% formic acid in HPLC grade H2O as solvent A and 0.1% formic acid in HPLC grade ACN as solvent B. The sample is eluted over a linear gradient of 2–40% B at a flow rate of 700 nL/min over a 60 min gradient. The peptides are introduced into the mass spectrometer by nanoelectrospray ionization. The mass spectrometer performs in positive mode with a survey scan with a 70,000 resolution, AGC of 1E6, max fill time 250 ms, and a scan range of 350–2000 m/z. The data-dependent MS/MS analysis is performed with a resolution of 17,500, AGC of 1E5, max fill time 100 ms, isolation window of 2.0 m/z and a loop count of 10. The voltage of the source is set at 2.3 kV and the capillary temperature is 250 °C [103].

3.2  Analyzing Site-Specific Acetylation Stoichiometry Using Isotopic Labeling and Data-Independent Acquisition Analysis

Utilizing chemical acetylation labeling strategy for stoichiometry determination of acetyl lysine requires heavy labeling from an in vitro labeling reaction, as detailed in this chapter. The isotopic label used should have a mass shift that can be easily observed in the MS2 fragmentation pattern, ideally outside of the isotopic envelope accurate determination of acetylation stoichiometry. We choose to use deuterated acetic anhydride (CD3CO2)O, for our in vitro labeling reaction, which results in the 3 Da mass shift at the acetylation site [103]. Furthermore, to account for the overestimation of stoichiometries at a high abundance with low heavy labeling, we corrected for the natural isotopic abundance of the light acetyl peptide by subtracting the natural isotopic abundance M+3 isotopic peak from the heavy deuterated acetyl M+0 isotopic peak based upon calculations of natural isotopic abundance distribution of the fragment ion. Utilizing deuterated acetic anhydride for labeling to generate a 3  Da mass shift in conjunction with MS2 fragment based in conjunction with MS2 fragment ion-based quantification of acetyl lysine stoichiometry significantly reduces the interference between the light and heavy acetylated fragment ions, increasing the accuracy and reducing the overestimation of

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stoichiometry determination [103]. These problems were discussed and tackled by moving toward other forms of quantification of the lysine stoichiometry using the MS2 fragments. This latest step forward in methodology to quantify stoichiometry of lysine acetylation has been described here as well as by others, utilizing data-independent acquisition to increase the sampling depth of a sample and the number of acetylation sites quantified. 3.2.1  Deuterated Acetic Anhydride Labeling and Digestion of Samples

1. To start with the same concentration in each of the samples dry down using a speedvac. 2. Resuspend sample to 50 μL total volume using 8 M urea, 100 mM ammonium bicarbonate with 5 mM DTT, with the pH at 8.5 in a 1.5 mL tube. Prepare the buffers fresh for each experiment. For subsequent labeling steps avoid buffers with free amine groups that would react with the acetic anhydride. 3. To denature the proteins place samples on Thermomixer C (Eppendorf), which has a heated lid to avoid condensation at 60 °C while mixing at 1000 RPM for 30 min. 4. Add 40 mM iodoacetamide to alkylate cysteine residues and incubate at 60 °C while mixing at 1000 RPM for 30 min on the Thermomixer C (Eppendorf). 5. To chemically acetylate unmodified lysine residues, add 2 μL of deuterated acetic anhydride to the samples, vortex, and incubate for 30 min, 60 °C, 1000 RPM on the Thermomixer C (Eppendorf). 6. Add ~10 μL NH4OH to bring the pH back up to ~8.0. Check the pH with a 0.5 μL spot on litmus paper and add more NH4OH as necessary. 7. For complete labeling repeat the chemical labeling from steps 6 and 7, again using deuterated acetic anhydride. 8. After the second round of chemical labeling incubate sample at pH 8 for 20 min at 60 °C with 1000 RPM mixing on the Thermomixer C (Eppendorf) to hydrolyze any O-acetyl esters formed during the reaction. 9. Add 100 mM ammonium bicarbonate to bring the volume up to 100 μL and the final concentration down to 2 M. 10. Add sequencing grade trypsin to a 1:100 ratio (trypsin:protein) and incubate from 4 h to overnight at 37 °C mixing at 350 RPM on Thermomixer C (Eppendorf). 11. Add 100 mM ammonium bicarbonate to bring the volume up to 200 uL and the final concentration down to 1 M for the subsequent gluC digestion. GluC digestion efficiency is reduced with urea levels above 1 M. 12. Add gluC in a 1:40 (gluC:protein) ratio and incubate from 4 h at 37 °C mixing at 350 RPM on Thermomixer C (Eppendorf).

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Additional digestion with GluC is necessary to digest labeled peptides to lengths amendable for LC-MS/MS analysis. 13. Quench the reaction by adding ~40 μL of acetic acid. 3.2.2  Desalting Prior to MS Analysis

Perform peptide desalting as detailed in Subheading 3.1.3, following steps 1–8 as written.

3.2.3  LC-MS/MS DIA Data Analysis

Our method to quantify acetylation stoichiometry uses a Thermo Q-Exactive Orbitrap mass spectrometer coupled to a Dionex Ultimate 3000 RSLC nano HPLC with a Waters Atlantis reverse phase column (100 μm × 150  mm). The mobile phases contain 0.1% formic acid in HPLC grade H2O as solvent A and 0.1% formic acid in HPLC grade ACN as solvent B. The sample is eluted over a linear gradient of 2–40% B at a flow rate of 700 nL/min over a 60 min gradient. The peptides are introduced into the mass spectrometer by nanoelectrospray ionization. The mass spectrometer performs in positive mode with a survey scan with a 70,000 resolution, AGC of 1E6, max fill time 100 ms, and a scan range of 400– 1000 m/z. The survey scan was followed 30 DIA scans in profile mode with a resolution of 35,000, AGC 1E6, 20 m/z window, and NCE of 25 to balance the frequency of b-ions, y-ions, and the number of PSMs to gain higher fragmentation coverage for MS2 quantitation, particularly important for site-specific quantitation of multiple lysine containing peptides [103]. The voltage of the source is set at 2.3 kV and the capillary temperature is 250 °C.

4  Data Analysis and Quantification of Acetylation Stoichiometry 4.1  Generating a Spectral Library

Analyzing acetylation stoichiometry using DIA requires a spectral library containing all combinations of light and heavy acetyl fragment ions. In this study, we use MaxQuant to perform a database search of the completely light acetylated samples to generate a spectral library followed by the subsequent in silico generation of the heavy labeled fragment ions [103]. To perform DDA data analysis, we use the openly available MaxQuant software package to perform our database searches and generate our library within the Spectronaut Professionals software (Biognosys) [104–108]. 1. Perform a database search in MaxQuant to identify peptides present in the DDA samples analyzed. Add the modifications for the non-isotopically labeled “Acetyl (K)” and the heavy deuterated “Acetyl D3 (K)” to the database search. The “Acetyl (K)” modification should be specified as a modification on lysines (K) be located anywhere except the C terminus of the peptide, and contains 2 carbons, 2 hydrogens, and 1 oxygen, which has a mass of 42  Da. The “Acetyl D3 (K)” modification should be specified as a modification on lysines (K) be located anywhere except the C terminus of the peptide,

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and contains 2 carbons, 3 deuterium, 1 oxygen, and the loss of 1 hydrogen, which has mass of 45  Da. Assign “Acetyl (K)” and “Oxidation (M)” modifications as variable modification through the sample with “Carbamidomethyl (C)” as a fixed modification [104–108]. Allow for up to 5 modifications and 5 missed cleavages by Trypsin and GluC due to the acetyl labeling blocking all lysine digestion sites within the peptide. 2. For the FASTA file used, you will need to configure a file that contains the internal retention time standard peptides if utilizing the Biognosys iRT kit by adding the Biognosys iRT sequence to the proteome of the organism of interest. In our case, we added the Biognosys iRT sequence to the human SwissProt sequences FASTA file [102]. 3. To prepare a library in Spectronaut (Biognosys), generate the library from the MaxQuant output utilizing the same modified FASTA file as a protein database. This will generate the spectral library file of the non-isotopically labeled acetyl peptide spectral library. 4. To add the deuterated heavy acetyl label in silico, export the library file for subsequent analysis. 5. Load the library file into the Spectronaut Library Modifier, and select the modification you would like to add, in this case “Acetyl D3 (K)” and which modification it pairs with, the “Acetyl (K)”, from the existing library. Export the in silico inflated spectral library file [103]. 6. Import the in silico inflated library file into Spectronaut (Biognosys) to utilize the inflated library for downstream analysis. 7. Alternatively to using Spectronaut (Biognosys), the open source Skyline proteomics software can be used to generate a spectral library and analyze DIA data. In the case of Skyline, the heavy label can be generated within the modification settings, by ­adding the isotope label of deuterium to the “Acetyl (K)” modification settings [109]. 4.2  Analyzing DIA Mass Spectrometry Data 4.2.1  Analysis of the Raw DIA Sample Analysis

1. Add raw files of the deuterated acetic anhydride labeled samples acquired with the DIA sample analysis. 2. Pair the samples with the inflated spectral library to be used for the search, identification, and quantification of the peptides. 3. Set the identification Q-value cutoff at 0.1 and utilize the earlier generated FASTA file, containing the iRT sequence if using Spectronaut (Biognosys). 4. Export the report, including the modified peptide sequence, the unmodified peptide sequence, the fragment ion sequence, and the peak area of the fragment ions to quantify the acetyl

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stoichiometry for lysine containing peptides. Also include data quality measures, such as the peptide identification Q value for later filtering by data quality. 4.2.2  Quantification of Site-Specific Acetyl Lysine Stoichiometry

The quantification of site-specific lysine acetyl stoichiometry DIA analysis quantifies stoichiometry of MS2 fragments, rather than a MS1-based approach. This allows for site specificity within a peptide containing multiple lysines; calculating the stoichiometry of fragments from each lysine independently, rather than an overall average of the peptide [103]. MS2 fragment quantification overcomes interference and bias with MS1 quantification that can result in an overestimation of the acetyl stoichiometry at many low abundant and acetyl occupancy sites. We perform our stoichiometry analysis utilizing R version 3.3.3 (2017-03-06) “Another Canoe” and R studio version 1.0.1316. The analysis described here can be performed in many data analysis programs or software available. 1. Filter the data by MS2 fragments containing a single lysine for site-specific quantification. Fragments containing multiple lysines will not provide site-specific information on lysine acetylation label stoichiometry. 2. Filter the data by the presence of both the heavy and light label to have both values for subsequent stoichiometry calculations. For the analysis of offline prefractionated DIA data, filter by the presence of both the heavy and light label before summing the values across any fractions of a single biological sample. 3. Filter by peptide identification quality, we require a Q score of the peptide identification for one of the two labeled forms, heavy or light, of the acetyl lysine containing peptide. 4. Perform isotopic envelope corrections on the heavy labeled peak by using the BRAIN package in R [110]. This package calculates the M+3 peak of the light peak using the fragment ion sequence and natural isotopic distributions, which then is subtracted from the heavy M+0 peak to determine the abundance of the heavy peak. This calculation corrects for the overlap between the natural abundance of peptides containing three naturally occurring heavy atoms and the peak area assigned to the M+0 peak from the labeled heavy acetylated peptide. The isotopic correction is particularly important for reducing the overestimation of high stoichiometry lysine acetylation sites [103]. 5. Sum the area of the MS2 peaks of the y and b fragment ions of each lysine from the light and heavy labels separately to determine the total peak area of the heavy and light acetylated lysine. 6. Calculate the stoichiometry of the summed MS2 peaks for each lysine by dividing the total non-isotopically labeled peak area by the total peak area of both labels of the fragments.

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7. Using the FASTA file, assign the residue number of the lysine site within the fragments. This is important for multiply acetylated peptides which have unique sets of MS2 fragment for the C and N proximal lysine residues. 4.2.3  Statistics and Determining Significant Changes in Acetyl Stoichiometries

Calculating statistical significance of acetyl stoichiometry changes in dynamic systems poses a challenge to interpreting acetyl stoichiometry alterations due to the nature of correcting for multiple testing across samples as well as the dynamics along the time course. We go about tackling this issue by performing ANOVA analysis instead of single pairwise interactions between two time points. 1. Calculate the mean stoichiometry for each lysine within the proteome observed. 2. Calculate the linear model to perform one way repeated measures analysis of variance (ANOVA), which performs the comparison of three or more means within-subject variables. If performing comparisons between multiple groups rather than a time course, a t-test for two samples or sets of paired samples would be appropriate with a multiple testing correction depending on the experimental design. If performing between the means of more than two groups, a one way ANOVA or a mixed ANOVA may be more appropriate.

4.2.4  Analyzing Global Trends in Site-Specific Acetylation Stoichiometry Proteomics Data

In this chapter we have used subcellular fractionation with chemical labeling of lysine residues coupled to data-independent acquisition mass spectrometry-based proteomics to quantify lysine acetylation stoichiometry. The subcellular localization distribution within the spectral libraries demonstrated the increase in coverage provided by both the subcellular isolation and offline prefractionation, as well as the two methods in conjunction (Fig. 2). Offline prefractionation (light blue) resulted in approximately double the numbers of proteins identified throughout the compartment as compared to a single whole cell lysate DDA spectral library (gray), and a similar increase in the number of peptides identified. The subcellular isolation followed a similar pattern, with the spectral library of the offline prefractionated chromatin isolation (green) resulting in double the protein and peptide identifications of the single sample of the chromatin isolation (orange) (Fig. 1). The two separation techniques used in conjunction expanded the spectral library even further, increasing protein and peptides identifications threefold above a single whole cell lysate DDA spectral library. For analysis of subcellular compartment isolation and its impact on increasing the coverage of lysine sites, we analyzed subcellular localization based upon evidence in annotated databases, such as UniProt (Figs. 3 and 4a). The greatest increase in compartment-­ specific identifications in the spectral library was seen in the chromatin isolation in combination with offline prefractionation, increasing the percent of nuclear proteins, peptides, and acetyl

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Fig. 3 Subcellular localization identification and abundances of spectral libraries and DIA sample analysis. (a) Percent of Proteins identified in spectral libraries by subcellular localization and (b) and percent of peptides identified in spectral libraries by subcellular localization of the spectral libraries generated from a Single Injection Whole Cell Lysate (WCL), a Single Chromatin Isolation (utilizing subcellular fractionation without offline prefractionation), a Fractionated WCL utilizing offline prefractionation, Fractionated Chromatin Isolation utilizing offline prefractionation on a subcellular fractionated chromatin isolation, a Fractionated WCL utilizing offline prefractionation with a single chromatin isolation (utilizing subcellular fractionation without offline prefractionation), and the Fractionated Chromatin Isolation and Fractionated WCL together utilizing offline prefractionation, on both whole cell lysate and a subcellular fractionated chromatin sample. (c) Percent of acetylated peptides identified from analysis of DIA samples from an offline prefractionated WCL, a single injection WCL sample, a mitochondrial isolation, and a chromatin isolation by subcellular compartment. (d) Percent of acetylated peptide abundance of DIA samples from an offline prefractionated WCL, a single injection WCL sample, a mitochondrial isolation, and a chromatin isolation by subcellular compartment

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Fig. 4 Analysis of overlapping and unique features of subcellular localization in spectral libraries. (a) The overlap of the annotated subcellular localization of proteins as obtained by UniProt demonstrating the overlap of proteins across compartments and the existence of a single protein across compartments. (b) An analysis of the overlap of peptide identifications of spectral libraries generated from a Single Injection Whole Cell Lysate (WCL), a Single Chromatin Isolation (utilizing subcellular fractionation without offline prefractionation), a Fractionated WCL utilizing offline prefractionation, and Fractionated Chromatin Isolation utilizing offline prefractionation on a subcellular fractionated chromatin sample. (c) DAVID analysis of the unique KEGG pathway enrichment and significance of the enrichment of peptides from subcellular chromatin isolation in comparison to the offline prefractionation WCL and fractionated chromatin isolation spectral library

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peptides identified to approximately 50% of the library (Fig. 3a, b). The combination of subcellular compartment isolation and offline prefractionation significantly increases spectral library coverage and enables deep interrogation of compartment-specific acetylation stoichiometry as shown in Fig. 4, which highlights the subcellular functions captured in our chromatin isolation and offline prefractionation approach (Fig. 4a, c). Increasing depth of the spectral library results in increased, peptides, acetyl peptides, and both forms of the isotopic acetyllysine label deconvoluted from the DIA MS2 spectra, as shown in Fig. 5. Analysis of the acetylation stoichiometry of a single injection of whole cell lysate without offline prefractionation resulted in 25% coverage of the acetylation site stoichiometry when compared to offline prefractionation DIA analysis. However, depth of coverage in a single DIA sample was greatly improved by subcellular compartment isolation, with the depth of coverage within isolated chromatin being comparable to chromatin acetylation sites quantified in an offline prefractionation of a whole cell lysate (Fig. 3c, d). Furthermore, the increased depth of the spectral library in conjunction with subcellular isolation prior to DIA analysis resulted in the increase in the number of acetylation site stoichiometries quantified in the compartment of interest. In addition, the reduction in sample complexity by subcellular isolation increased the likelihood of observing both the non-isotopic and isotopically labeled acetyl fragment ions to calculate acetylation stoichiometry.

Fig. 5 Subcellular fractionation increases depth similar to offline prefractionation with lower peptide abundance. (a) The difference in unique peptides identified, unique lysine (K) containing peptides, and unique peptides with quantified stoichiometry in the DIA analysis of Offline Prefractionated Whole Cell Lysate (WCL, red), Subcellular Mitochondrial Isolation (green), and Subcellular Nuclear Isolation (blue) using the Fractionated WCL Fractionation spectral library and the Subcellular Fractionation with Offline WCL Fractionation spectral library demonstrating similar levels of acetylation stoichiometry coverage. (b) The distribution of the log10 peak area of MS2 fragments from acetyl-lysine containing peptides with determined stoichiometry showed the mean peak area of peptides from the single DIA analysis of subcellular isolation quantifies a similar depth of lysine acetylation sites captured with offline prefractionation

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The global distribution of acetylation stoichiometry within a whole cell lysate measurements yielded a median of approximately 2% (Fig.  6a). Our method of subcellular compartment isolation demonstrates important differences, with resolution capable of revealing the higher acetylation stoichiometry proteins occurring within the isolated mitochondria and nucleus [24, 111] (Fig. 6b). Calculation of site-specific stoichiometry from a whole cell lysate reflect the combination of stoichiometries of a protein across subcellular compartments. However, our subcellular fractionation method captures the dynamic stoichiometries of a protein across different subcellular compartments, which may be independently regulated (Fig. 7).

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Fig. 6 Analysis of global lysine acetylation stoichiometry by spectral library. (a) Density plot of the relative distribution of lysine acetylation stoichiometry of DIA samples from an offline prefractionated whole cell lysate (WCL, pink), a single injection WCL (green), a mitochondrial isolation (blue), and a chromatin isolation (purple). The median stoichiometry in these samples is 2% and the distribution was not highly variable, area under the curve normalized across samples. (b) Histogram of the absolute distribution of mean lysine site acetylation stoichiometry of DIA analysis from an offline prefractionated WCL, a Single Injection WCL sample, and a chromatin isolation across a time course experiment demonstrates the similar distribution of acetylation stoichiometries. (c) Density plot of the relative distribution of lysine acetylation stoichiometry by subcellular localization of DIA samples from an offline prefractionated WCL, a single injection WCL sample, a mitochondrial isolation, and a chromatin isolation shows unique subcellular patterns appear in the subcellular isolation DIA samples not seen in the whole cell lysate due to whole cell averages in stoichiometry

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Fig. 7 DIA analysis of subcellular fractions reveals subcellular compartment-specific patterns of lysine acetylation stoichiometry. (a) Density plot of the relative distribution of lysine acetylation stoichiometry of nuclear localized lysine acetylation sites from DIA samples from an offline prefractionated whole cell lysate (WCL, pink), a single injection WCL sample (green), and a chromatin isolation (blue). (b) Density plot of the relative distribution of lysine acetylation stoichiometry of mitochondria localized lysine acetylation sites from DIA samples from an offline prefractionated WCL (pink), a single injection WCL sample (green), and a mitochondrial isolation (blue). (c) Venn diagram of overlap and unique nuclear lysine acetylation sites from DIA samples from an offline prefractionated whole cell lysate (brown), a single injection WCL sample (purple), and a chromatin isolation (green). (d) Venn diagram of overlap and unique mitochondria lysine acetylation sites from DIA samples from an offline prefractionated WCL (brown), a single injection WCL sample (green), and a mitochondrial isolation (purple). (e) Density plot of the relative distribution of lysine acetylation stoichiometry of unique nuclear lysine acetylation sites from DIA samples from an offline prefractionated WCL (pink), a single injection WCL sample (green), and a chromatin isolation (blue). (f) Density plot of the relative distribution of lysine acetylation stoichiometry of unique mitochondria localized acetylation sites from DIA samples from an offline prefractionated WCL (pink), a single injection WCL sample (green), and a mitochondrial isolation (blue)

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5  Discussion and Perspective We have discussed methods to utilize chemical labeling of lysine residues coupled with data-independent acquisition mass spectrometry-­based proteomics to quantify lysine acetylation stoichiometry, increasing the sampling depth and the number of acetylation sites captured. Furthermore, the methods describing subcellular organelle enrichment and its impact on increasing the coverage of lysine sites of a specific organelle, provided a useful tool for in-depth studies to investigate the impacts of different ­factors on lysine acetylation dynamics in specific organelles. This approach provides a useful tool for examining the differences in the regulation of lysine acetylation within cellular organelles, in particular the differences between enzymatic and nonenzymatic acetylation in the nucleus, cytoplasm, and mitochondria. Our approach has further demonstrated that subcellular fractionation gives an increase in sampling depth of a specific organelle over orthogonal separation approaches of a whole cell lysate alone. Finally, we have demonstrated the utility for a deep spectral library for DIA sample analysis. With a deep spectral library, offline prefractionation of the experimental samples was not necessary when performing final DIA analysis. The main bottleneck in the depth of peptide identification is the generation of the spectral library. Furthermore, the technical error has been decreased from the reduced handling of samples without the offline prefractionation on DIA sample analysis. Mass spectrometry-based proteomics approaches for identifying and quantifying lysine acetylation have undergone many improvements within the last decade, now enabling comprehensively quantification of acetylation stoichiometry in an entire proteome [24]. Initial proteomics studies used antibody-based enrichment strategies, which largely expanded the acetylation sites identified [4, 12]. Quantification of these sites was undertaken by combining enrichment-based methods with SILAC labeling for relative quantification of acetylation [13, 14, 28]. These studies began to profile the dynamic nature of lysine acetylation; however, they were limited to systems where SILAC was readily usable. Furthermore, questions remained about the stoichiometry of lysine acetylation, as relative quantification can indicate how much acetylation of a site has increased or decreased. Because enrichment strategies removed non-acetylated peptides from the sample analyzed, these ratios do not reflect the total peptide abundance. The first undertaking of quantifying lysine acetylation with tandem mass tags (TMTs) revealed more information about the complex dynamics of mitochondrial protein acetylation in response to diet [21, 25, 112]. Recently, we and others have reported a data-­ dependent acquisition (DDA) method to determine stoichiometric

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quantification rather than relative fold changes; however, this method had limits based upon the use of MS1 quantification, and lowly abundant modification was more susceptible to signal interference and bias [24, 74, 113, 114]. These problems were discussed and addressed by using the MS2 fragments. The latest step forward, described here as well as by others, utilizes data-­ independent acquisition to increase the peptides identified and the number of acetylation sites quantified [74, 110]. In this chapter, we have expanded the coverage of this method to subcellular organelles, resulting in considerably deeper coverage of the acetyl stoichiometry of these organelles, allowing for more detailed investigations of the dynamics and regulation of acetyl lysine stoichiometry, which is functionally compartmentalized within the cell. This method may be more broadly applied to other lysine acyl modifications [110]. Recent studies have used DIA approaches to quantify lysine succinylation across the proteome, demonstrating how this method can be expanded to examine the biological regulation and function of acyl modifications on lysine residues [110].

Acknowledgments We would like to thank Ian Lienert, Tejas Gandhi, Oliver Bernhardt, Lukas Reiter from Biognosys for the development of the software to generate the in silico labeled spectral library and analyze DIA MS data. This work was supported by GM65386 to J.M.D. and by NIH National Research Service Award T32 GM007215 to (A.L. and J.B.) References 1. Olsen JV, Mann M (2013) Status of large-­ scale analysis of post-translational modifications by mass spectrometry. Mol Cell Proteomics 12:3444–3452 2. Olsen JV, Vermeulen M, Santamaria A et  al (2010) Quantitative phosphoproteomics reveals widespread full phosphorylation site occupancy during mitosis. Sci Signal 3:ra3 3. Wu R, Haas W, Dephoure N et al (2011) A large-scale method to measure absolute protein phosphorylation stoichiometries. Nat Methods 8:677–683 4. Kim SC, Sprung R, Chen Y et  al (2006) Substrate and functional diversity of lysine acetylation revealed by a proteomics survey. Mol Cell 23:607–618 5. Yu BJ, Kim JA, Moon JH et  al (2008) The diversity of lysine-acetylated proteins in Escherichia coli. J  Microbiol Biotechnol 18:1529–1536 6. Zhang J, Sprung R, Pei J  et  al (2009) Lysine acetylation is a highly abundant and

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56. Carrer A, Parris JLD, Trefely S et  al (2017) Impact of a high-fat diet on tissue acyl-CoA and histone acetylation levels. J  Biol Chem 292:3312–3322 57. Lee JV, Carrer A, Shah S et  al (2014) Akt-­ dependent metabolic reprogramming regulates tumor cell histone acetylation. Cell Metab 20:306–319 58. Hallows WC, Lee S, Denu JM (2006) Sirtuins deacetylate and activate mammalian acetyl-­ CoA synthetases. Proc Natl Acad Sci U S A 103:10230–10235 59. Schwer B, Bunkenborg J, Verdin RO et  al (2006) Reversible lysine acetylation controls the activity of the mitochondrial enzyme acetyl-­CoA synthetase 2. Proc Natl Acad Sci U S A 103:10224–10229 60. Starai VJ (2002) Sir2-dependent activation of acetyl-CoA synthetase by deacetylation of active lysine. Science 298:2390–2392 61. Pehar M, Puglielli L (2013) Lysine acetylation in the lumen of the ER: a novel and essential function under the control of the UPR. Biochim Biophys Acta 1833:686–697 62. Pehar M, Lehnus M, Karst A et  al (2012) Proteomic assessment shows that many endoplasmic reticulum (ER)-resident proteins are targeted by N(epsilon)-lysine acetylation in the lumen of the organelle and predicts broad biological impact. J  Biol Chem 287:22436–22440 63. Ogryzko VV, Schiltz RL, Russanova V et  al (1996) The transcriptional coactivators p300 and CBP are histone acetyltransferases. Cell 87:953–959 64. Schiltz RL, Mizzen CA, Vassilev A et  al (1999) Overlapping but distinct patterns of histone acetylation by the human coactivators p300 and PCAF within nucleosomal substrates. J Biol Chem 274:1189–1192 65. Kundu TK, Palhan VB, Wang Z et al (2000) Activator-dependent transcription from chromatin in vitro involving targeted histone acetylation by p300. Mol Cell 6:551–561 66. McManus KJ, Hendzel MJ (2003) Quantitative analysis of CBP- and P300-­ induced histone acetylations in vivo using native chromatin. Mol Cell Biol 23:7611–7627 67. Yang Y-Y, Yu-Ying Y, Grammel M et al (2011) Identification of lysine acetyltransferase p300 substrates using 4-pentynoyl-coenzyme A and bioorthogonal proteomics. Bioorg Med Chem Lett 21:4976–4979 68. Karanam B, Jiang L, Wang L et  al (2006) Kinetic and mass spectrometric analysis of p300 histone acetyltransferase domain autoacetylation. J Biol Chem 281:40292–40301 69. Stiehl DP, Fath DM, Liang D et  al (2007) Histone deacetylase inhibitors synergize p300 autoacetylation that regulates its transactiva-

Site-Specific Lysine Acetylation Stoichiometry Across Subcellular Compartments tion activity and complex formation. Cancer Res 67:2256–2264 70. Wagner GR, Payne RM (2013) Widespread and enzyme-independent Nε-acetylation and Nε-succinylation of proteins in the chemical conditions of the mitochondrial matrix. J Biol Chem 288:29036–29045 71. Ghanta S, Grossmann RE, Brenner C (2013) Mitochondrial protein acetylation as a cell-­ intrinsic, evolutionary driver of fat storage: chemical and metabolic logic of acetyl-lysine modifications. Crit Rev Biochem Mol Biol 48:561–574 72. Wagner GR, Hirschey MD (2014) Nonenzymatic protein acylation as a carbon stress regulated by sirtuin deacylases. Mol Cell 54:5–16 73. Paik WK, Pearson D, Lee HW et  al (1970) Nonenzymatic acetylation of histones with acetyl-CoA.  Biochim Biophys Acta 213:513–522 74. Baeza J, Smallegan MJ, Denu JM (2015) Site-specific reactivity of nonenzymatic lysine acetylation. ACS Chem Biol 10:122–128 75. Hansen JC, Tse C, Wolffe AP (1998) Structure and function of the core histone N-termini: more than meets the eye. Biochemistry 37:17637–17641 76. López-Rodas G, Brosch G, Georgieva EI et al (1993) Histone deacetylase. A key enzyme for the binding of regulatory proteins to chromatin. FEBS Lett 317:175–180 77. Daujat S, Bauer U-M, Shah V et  al (2002) Crosstalk between CARM1 methylation and CBP acetylation on histone H3. Curr Biol 12:2090–2097 78. Hecht A, Laroche T, Strahl-Bolsinger S et al (1995) Histone H3 and H4 N-termini interact with SIR3 and SIR4 proteins: a molecular model for the formation of heterochromatin in yeast. Cell 80:583–592 79. Yu Y, Song C, Zhang Q et al (2012) Histone H3 lysine 56 methylation regulates DNA replication through its interaction with PCNA. Mol Cell 46:7–17 80. Struhl K (1998) Histone acetylation and transcriptional regulatory mechanisms. Genes Dev 12:599–606 81. Clark D, Reitman M, Studitsky V et al (1993) Chromatin structure of transcriptionally active genes. Cold Spring Harb Symp Quant Biol 58:1–6 82. Dhalluin C, Carlson JE, Zeng L, et al (2002) Structure and ligand of a histone acetyltransferase bromodomain. https://doi. org/10.2210/pdb1n72/pdb 83. Soutoglou E, Katrakili N, Talianidis I (2000) Acetylation regulates transcription factor activity at multiple levels. Mol Cell 5: 745–751

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Chapter 7 Lysine Acetylation of Proteins and Its Characterization in Human Systems David K. Orren and Amrita Machwe Abstract Posttranslational acetylation modifications of proteins have important consequences for cell biology, including effects on protein trafficking and cellular localization as well as on the interactions of acetylated proteins with other proteins and macromolecules such as DNA. Experiments to uncover and characterize protein acetylation events have historically been more challenging than investigating another common posttranslational modification, protein phosphorylation. More recently, high-quality antibodies that recognize acetylated lysine residues present in acetylated proteins and improved proteomic methodologies have facilitated the discovery that acetylation occurs on numerous cellular proteins and allowed characterization of the dynamics and functional effects of many acetylation events. This article summarizes some established biochemical information about how protein acetylation takes place and is regulated, in order to lay the foundation for subsequent descriptions of strategies used by our lab and others either to directly study acetylation of an individual factor or to identify groups of proteins targeted for acetylation that can then be examined in isolation. Key words Posttranslational modifications, Acetyltransferases, Histone deacetylases, Sirtuins, Immunodetection, Proteomics, Bromodomain

1  Introduction Research over at least the last three decades has revealed that posttranslational protein modifications are ubiquitous and essential for many cellular functions. Numerous types of protein modifications exist, from the covalent additions of small moieties such as phosphate, methyl, and acetyl groups or much larger modifications including farnesyl, ubiquitin (and small ubiquitin-like modifiers [SUMO]), and ADP-ribose groups. These modifying groups are generally linked to characteristic amino acid side chains within proteins, although linkage of acetyl groups to N-terminal amine groups is also possible. Importantly, posttranslational modifications serve to alter protein function without the need for new transcription and translation; in many cases, these modifications

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allow cells to more quickly respond to circumstances inside and outside the cell or its sub-compartments. At the molecular level, these modifications can mediate alterations in a protein’s catalytic activity, its interactions with other proteins and other macromolecules, its cellular trafficking and localization, and its stability. The array and timing of possible modifications on individual proteins can be highly complex and serve to fine-tune these alterations in function. For example, the p53 (TP53) tumor suppressor protein is subject to acetylation, ubiquitination, sumoylation, and multiple phosphorylation events that regulate its abundance and primary function as a stress-inducible transcription factor. Therefore, in addition to assessing the rates of gene transcription and mRNA translation, one must consider the possibility and type of posttranslational modifications to an individual protein when characterizing its function and effects on broader intracellular pathways.

2  Protein Acetylation Biochemistry 2.1  Basics of Lysine Acetylation of Proteins

Because of their key roles in cell signaling pathways governing cell proliferation and survival, it is appropriate that phosphorylation and dephosphorylation events have been investigated extensively. However, acetylation events are also very important to study, as they have major roles in intracellular metabolism and other pathways. Recent studies using high-resolution mass spectrometry identified at least 1750 different acetylated proteins in human cell lines, many having multiple acetylation sites [1, 2]. Notably, the Uniprot and Phosida databases are valuable bioinformatics tools to search for acetylated proteins and sites across organisms. Acetylation of proteins occurs overwhelmingly on the ε-amine group present on the side chain of lysine residues, also known as Nε-acetylation (Fig. 1), although this modification can occasionally occur on the N-terminal amine group (designated Nα-acetylation) of proteins. Acetylation masks the basic and positively charged character of

Fig. 1 Lysine acetylation and deacetylation reactions. Substrates, enzymes, and products involved in the acetylation of lysine residues and the reverse reaction (deacetylation) are depicted. KAT lysine acetyltransferase, HAT histone acetyltransferase, HDAC histone deacetylase

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these amine groups, generally serving to reduce or eliminate ­electrostatic interactions with acidic or negatively charged moieties. Among other possibilities, lysine acetylation can affect intramolecular protein structure or affect interactions with other proteins, DNA, and other macromolecules. In perhaps the most well-known example, acetylation of lysines on the positively charged tails of histones loosens the association of nucleosomes with the negatively charged phosphodiester backbone of DNA, thus opening the local chromatin structure to facilitate access to DNA for transcription or other DNA metabolic processes. Also in regard to chromatin structure and gene regulation, acetylation of transcription factors (often within complexes containing acetyltransferases) plays a role in transcriptional status. Acetylation of histones and transcription factors generally favors transcriptional activation, although this paradigm is not absolute. More recently, it has been recognized that acetylation of histone and nonhistone proteins plays a key role in many other processes, including DNA repair, sister chromatid cohesion, cytoskeletal dynamics, and cellular metabolic state [3]. These investigations have expanded our appreciation of acetylation as a key posttranslational modification that is critical to many cellular processes. Many acetylated proteins seem to be present as parts of large protein complexes, suggesting that acetylation may help mediate or stabilize protein-protein interactions. It is noteworthy that so-­ called bromodomains, identified within at least 40 human proteins, appear specific in mediating albeit relatively weak interactions with acetylated lysine residues; however, this binding affinity can be synergized in proteins with multiple bromodomains [4]. In regard to histone tails containing acetylated lysines, these can help specify the binding of proteins containing bromodomains and contribute to the (usually activated) transcriptional state of that chromatin region. Intriguingly, bromodomains are actually present in several acetyltransferase enzymes. In such cases, the interaction of this domain with acetylated lysines may facilitate subsequent modification of additional lysines in the bound protein, as has been suggested for the possibly ordered acetylation of K16, K12, K8, and K5 in histone H4 [4]. In many if not most cases, the acetylation state of a protein is determined by a balance between addition and removal of acetyl groups carried out by acetyltransferases and deacetylases, respectively. The interplay between these opposing actions can mediate several mechanistically distinct events. One possibility is that acetylation of a target lysine can directly block other potential modifications (ubiquitination, sumoylation, methylation, proprionylation, butyrylation, or neddylation) at that specific residue, while deacetylation would permit other modifications to occur. For example, acetylation of lysines also targeted by ubiquitination has been shown, in some cases, to stabilize proteins by reducing turnover

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normally carried out by ubiquitin-mediated proteosomal ­degradation [5, 6]. A second mechanistic scenario is that acetylation (or deacetylation) may control modification of a nearby residue in the target protein. Notably, the catalytic activities for these non-­acetylation modifications may be contained in the acetyltransferase or deacetylase itself, although it is more likely that other modifying activities will be present in associated factors. Another mechanistic possibility is that acetylation state will determine localization or trafficking of the target protein via interactions with other proteins or macromolecules. Assembly and action of transcription-related complexes at discrete promoter regions depending upon histone acetylation status is a well-known example; another is the role of acetylation of the Werner syndrome (WRN) protein in its trafficking between the nucleolus and nuclear foci associated with DNA replication or repair [7–9]. Interestingly, in some circumstances noncoding RNAs may interact with acetylases or deacetylases to help target their action and regulate acetylation of key targets [10]. In relation to characterizing and understanding the dynamics of these posttranslational modifications, it is important that the acetylation status of proteins is linked to cellular energy status in two ways: (1) through production of acetyl-CoA needed for all acetylation reactions and (2) and through levels of intracellular NAD+, the necessary cofactor for sirtuin deacetylases, with nutrient deprivation conditions favoring (higher NAD+ and) deacetylation [4]. Notably, use of deacetylase inhibitors (listed in Table 3) is a general strategy that decreases deacetylation of target proteins, thereby shifting this balance toward acetylation and enhancing both detection of this modification and its potential molecular and cellular effects. Conversely, acetyltransferase inhibitors would shift this balance toward deacetylation and its concomitant effects. It should be emphasized that use of these inhibitors (which are often not specific to a single acetyltransferase or deacetylase) in cellular or clinical settings likely impacts the modification state of multiple proteins simultaneously, having pleiotrophic cellular and molecular effects that are often difficult to attribute to changes in the modification of a single protein. Investigators should be aware of the specificity of inhibitors to be considered and their appropriateness for the design of each experiment. In cases where multiple acetyltransferases or deacetylases can act on a specific protein target, inhibitors that block multiple enzymes might even be preferable. Irregardless, deacetylase and acetyltransferase inhibitors can be useful for circumstances beyond enriching for the acetylated or unmodified form of the target protein, for instance to test whether acetylation state might impact the subcellular localization of a target protein or its association or co-localization with another factor. Similarly, experiments employing overexpression of acetyltransferases or deacetylases can alter this balance toward acetylation or

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Table 3 Selected histone deacetylase inhibitorsa Inhibitor name

Targeted HDACs

Chemical type

Trichostatin A

Class I–IV

Hydroxamic acid

SAHA/Vorinostatb

Class I–IV

Hydroxamic acid

Belinostat

Class I–IV

Hydroxamic acid

Class I–IV

Hydroxamic acid

Givinostat

Class I–IV

Hydroxamic acid

Resminostat

Class I–IV

Hydroxamic acid

Abexinostat

Class I–IV

Hydroxamic acid

Quisinostat

Class I–IV

Hydroxamic acid

Rocilinostat

Class II

Hydroxamic acid

Practinostat

Class I, II, and IV

Hydroxamic acid

CHR-3996

Class I

Hydroxamic acid

Valproic acidb

Class I and IIa

Short chain fatty acid

Butyric acid

Class I and II

Short chain fatty acid

Phenylbutyric acid

Class I and II

Short chain fatty acid

Chidamide

HDAC1, HDAC2, HDAC3, HDAC10

Benzamide

Entinostat

Class I

Benzamide

Tacedinaline

Class I

Benzamide

4SC202

Class I

Benzamide

Mocetinostat

Class I and IV

Benzamide

Romidepsinb

Class I

Cyclic peptide

Apicidin

Class I

Cyclic peptide

Nicotinamide

Class III

Sirtuin inhibitor

Sirtinol

SIRT1, SIRT2

Sirtuin inhibitor

Cambinol

SIRT1, SIRT2

Sirtuin inhibitor

EX-527

SIRT1, SIRT2

Sirtuin inhibitor

b

Panabiostat

b

Derived from refs. 16, 23, 40 Indicates FDA-approved drug

a

b

deacetylation, respectively, of (target) proteins, and vice versa for studies using silencing of acetyltransferases or deacetylases. 2.2  Acetyltransferases

Lysine acetyltransferases or KATs, often also called histone acetyltransferases (HATs) based on their most well-known but not exclusive targets, transfer acetyl groups from their acetyl coA cofactors

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to the aforementioned (epsilon) amine groups of specific lysines in proteins. There is also a separate group of N-terminal acetyltransferases (NATs) that transfer acetyl groups to the N-terminal (alpha) amine group of proteins, although they will not be described further here [for review, see [11]]. Most acetyltransferases act through ordered binding and formation of a ternary complex between enzyme, acetyl coA, and target protein, although p300 does not appear to form a stable ternary complex [4]. Numerous enzymes with lysine acetyltransferase activity have been identified in human cells (Table 1), including the reasonably well-studied p300, CBP, HAT1, GCN5, PCAF, TAF1, TIP60/PLIP, MYST1/HMOF, MYST2/HBO1, MYST3/MOZ, MYST4/MORF, ELP3, TFIIIC90, SRC1, ACTR, P160, and CLOCK enzymes [12–14]. Lysine acetyltransferases have been categorized into three groups, the (1) GCN5/PCAF (or GNAT), (2) MYST, and (3) p300/CBP families [10, 15]. These families and individual protein names have survived in the literature, even though revised nomenclature was proposed in 2007 to designate all known lysine acetyltransferases as KATs (see Table 1), with protein designations given the same number with a unique letter being most highly homologous to one another [12]. Lysine acetyltransferases have also been differently categorized into two categories based on subcellular localization, with Type A lysine acetyltransferases being nuclear and Type B being cytoplasmic. Relevant to the functional significance of this classification, Type B lysine acetyltransferases are responsible for acetylating newly synthesized histones, while Type A enzymes are involved the acetylation dynamics of histones and other nuclear proteins [16]. Acetylation of histones by lysine acetyltransferases not only promotes transcription but also helps facilitate DNA replication and repair [13]. Moreover, acetylation has been demonstrated to affect other factors involved in DNA replication and DNA damage responses, including H2AX, p53, PCNA, TDG, OGG1, NEIL2, APE-1, pol beta, FEN-1, WRN, FANCJ (BACH1), XPA, and XPG [17]. Among the many acetyltransferases in cells from higher eukaryotes, CBP (CREB binding protein) and p300 are not as discriminating in their protein targets [4] and have been more extensively investigated than others. p300 and CBP are downstream effectors in cAMP, Notch, NFkappaB, and estrogen receptor signaling and in the cellular responses to stress including DNA damage and hypoxia [reviewed in [4]]. Much of the research on CBP and p300 [reviewed in [17]] has focused on their roles as transcriptional co-­ activators, via their ability to acetylate the lysine-rich N-terminal tails of histones H2A (K5 and K9), H2B (K5, K12, K15, and K20), H3 (K9, K14, K18, and K23), and H4 (K5, K8, K12, and K16), although most other acetyltransferases have also been demonstrated to acetylate histones [11]. These modifications partially neutralize positive charges on these histone tails, reducing their

Common name/other designations

HAT1

GCN5/GCN5L2

PCAF

CBP

p300

TAF/TAFII250

TIP60/PLIP

MYST3/MOZ

MYST4/MORF

MYST2/HBO1

MYST1/HMOF

ELP3

TFIIIC90/GTF3C4

SRC1/NCOA1

SRC3/NCOA3/ACTR/TRAM1

SRC2/NCOA2/TIFS/GRIP1/p160

CLOCK

ATF-2/CREB2/CREBP1

KAT designation

KAT1

KAT2A

KAT2B

KAT3A

KAT3B

KAT4

KAT5

KAT6A

KAT6B

KAT7

KAT8

KAT9

KAT12

KAT13A

KAT13B

KAT13C

KAT13D

Unassigned

Table 1 Human lysine acetyltransferasesa

n/a

n/a

n/a

n/a

n/a

n/a

GNAT/GCN5

MYST

MYST

MYST

MYST

MYST

n/a

CBP/p300

CBP/p300

GNAT/GCN5

GNAT/GCN5

GNAT/GCN5

Family designation(s)

Nuclear/cytoplasmic

Nuclear/cytoplasmic

Nuclear

Nuclear/cytoplasmic

Nuclear

Nuclear

Nuclear/cytoplasmic

Nuclear

Nuclear

Nuclear

Nuclear

Nuclear/cytoplasmic

Nuclear

Nuclear/cytoplasmic

Nuclear/cytoplasmic

Nuclear

Nuclear

Nuclear

Subcellular localization

P15336

O15516

Q15596

Q9Y6Q9

Q15788

Q9UKN8

Q9H9T3

Q9H7Z6

Q95251

Q8WYB5

Q92794

Q92993

P21675

Q09472

Q92793

Q92831

Q92830

O14929

(continued)

Accession numberb

Characterization of Protein Acetylation 113

ATAT1

ACAT1

NAT10

GCN5L1/BLOC1S1

Unassigned

Unassigned

Unassigned

Unassigned

n/a  not applicable (not assigned to existing families) a Derived from refs. 10, 11, 13 b Swiss-Prot database accession number

Common name/other designations

KAT designation

Table 1 (continued)

n/a

n/a

n/a

GNAT/GCN5

Family designation(s)

Mitochondrial

Nuclear

Mitochondrial

Cytoplasmic

Subcellular localization

P78537

Q9H0A0

P24752

Q5SQ10

Accession numberb

114 David K. Orren and Amrita Machwe

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interactions with other histones and particularly with the negatively charged phosphodiester backbone of DNA, the latter helping to mediate access of other proteins necessary to carry out transcription (or other DNA metabolic processes). CBP and p300 not only have highly homologous acetyltransferase catalytic domains, both also contain homologous bromodomain, cysteineand histidine-rich (CH) regions, and so-called KIX and SID domains that mediate interactions with other transcription activators and basal transcription factors [17]. Thus, CBP and p300 have two functions that promote transcription: (1) they serve as scaffold proteins that help form functional transcription complexes, and (2) contain histone acetyltranferase activity that modifies chromatin structure in promoter regions. In regard to their key roles in transcription of many genes, CBP and/or p300 serve as co-activators for at least 40 unique transcription factors and regulate transcription of at least 615 genes [4, 18, 19]. Moreover, more than 60 transcription factors have been demonstrated to be themselves acetylated [10]. CBP and p300 have been shown act redundantly in many circumstances (particularly in in vitro biochemical assays and in cellular studies using overexpression), but in other scenarios their functions do not overlap and each has some unique protein substrates and/or preferentially targeted lysines [17]. At least partially separate functions have been confirmed in experiments with transgenic mice [20–22]. An important consideration is that these and other lysine acetyltransferases can have both common and unique targets. This should be kept in mind in experiments to determine the acetyltransferase(s) responsible for acetylating any target protein as well as in experiments in which RNA interference-­ mediated or chemical inhibition of acetyltransferases is performed. Other investigations have shown that the relative abundance of CBP and p300 is involved in regulating cellular responses including proliferation and differentiation, likely due to limited amounts of these acetyltransferases being transferred between competing complexes [reviewed in [4, 17]]. Thus, unanticipated cellular consequences should not be surprising when inhibiting/silencing or overexpressing these (or other) lysine acetyltransferases in experimental settings (see below). There are some pathological circumstances for which inhibition of lysine alkyltransferases might be desirable, including heart disease, thrombocytopenia, diabetes, obesity, fatty liver disease, retroviral infections, and certain cancers with hyperactive lysine acetyltransferase function [4]. Furthermore, inhibitors of these enzymes would very useful in experimental settings to understand the functions and dynamics of acetylation modifications. Therefore, some effort has gone into identifying small molecule inhibitors of lysine acetyltransferases, although these compounds are not as far along in development and implementation as HDAC inhibitors (see below). A number of natural product acetyltransferase

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i­nhibitors have been discovered, including anacardic acid, curcumin, garcinol (and its derivative LTK-14), γ-butyrolactone analogs including MB-3, plumbagin, epigallocatechin-3-gallate (EGCG), quercetin, ochratoxin A, and Δ12-PG-J2 prostaglandin [4]. Some of these have preferential potency toward discrete lysine acetyltransferases but, unfortunately, most of these have modest potency at best and/or have concerning off-target effects. For example, EGCG also inhibits topoisomerases. Among potential synthetic inhibitors of acetyltransferases are analogs of isothiazolones, thiazoles, and quinolones [4]. Furthermore, some bisubstrate molecules mimicking conjugated substrate-product analogs (including Lys-CoA, Lys-CoA-Tat, H3-CoA20, H4K16-CoA, and H3K14-­CoA) have shown promise in inhibiting CBP, p300, or PCAF lysine acetyltransferases [4]. Unfortunately, these bisubstrate molecules in their current form are poorly taken up by cells and not amenable to clinical uses. However, this class of inhibitors has been utilized to advance mechanistic and crystallographic studies of various acetyltransferases. In their entirety, current lysine acetyltransferase inhibitors are generally either not potent enough or otherwise not suitable for clinical use. However, some of these inhibitors may have utility in experimental settings to decrease levels of the acetylated forms of target proteins. In such cases, lysine acetyltransferase inhibitors may have differential effects when compared to acetyltransferase knockdown strategies, particularly in stable knockdown models where compensation by upregulation of another acetyltransferase can ameliorate effects [4]. 2.3  Deacetylases or HDACs

Obviously, deacetylase enzymes (most often called histone deacetylases or HDACs) remove acetyl groups from lysine residues of histones and other proteins, reversing the action of lysine acetyltransferases. There are two groups of HDACs in human cells, the classical and sirtuin families that are segregated by homology and different cofactor usage. All classical members require Zn2+ for deacetylase activity, while sirtuins utilize NAD+. Deacetylases have also been assigned to four classes (Table 2), with class I consisting of HDAC1, HDAC2, HDAC3, and HDAC8, class II consisting of HDAC4 (IIa), HDAC5 (IIa), HDAC6 (IIb), HDAC7 (IIa), HDAC9 (IIa), and HDAC10 (IIb), class III consisting of sirtuins SIRT1–7, and with HDAC 11 being the sole member of class IV [10, 16]. From the discussion above, it is clear that the transcriptional status of genes is partly determined by the acetylation state of histones in their promoter regions. Histone acetylation by lysine acetyltransferases is critical for mediating transcriptional activation of genes, while HDAC-mediated histone deacetylation generally results in transcriptional repression. Because of their ability to deacetylate histones and other transcription-related target proteins, HDACs have become important drug targets, and a number of nonspecific and more specific inhibitors are now available

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Table 2 Human deacetylase enzymesa HDAC class (family)

Common name (sub-class)

Localization

Accession no.b

Class I (classical)

HDAC1 HDAC2 HDAC3 HDAC8

Nuclear Nuclear Nuclear Nuclear/cytoplasmic

Q13547 Q92769 O15379 Q9BY41

Class IIb (classical)

HDAC4 (IIa) HDAC5 (IIa) HDAC6 (IIb) HDAC7 (IIa) HDAC9 (IIa) HDAC10 (IIb)

Nuclear/cytoplasmicc Nuclear/cytoplasmic Cytoplasmic Nuclear/cytoplasmic Nuclear/cytoplasmic Cytoplasmic

P56524 Q9UQL6 Q9UBN7 Q8WUI4 Q9UKV0 Q969S8

Class III (sirtuin)

SIRT1 SIRT2 SIRt3 SIRT4 SIRT5 SIRT6 SIRt7

Nuclear Cytoplasmic Mitochondrial Mitochondrial Mitochondrial Nuclear Nuclear

Q96EB6 Q8IXI6 Q9NTG7 Q9Y6E7 Q9NXA8 Q8N6T7 Q9NRC8

Class IV (classical)

HDAC11

Nuclear

Q96DB2

Derived from refs. 11, 13 Class II HDACs are often expressed in a cell-specific manner c Some class II HDACs can move partly to nucleus, dependent on conditions a

b

(Table 3), with a few even approved for clinical uses. Since expression of tumor suppressor genes is often downregulated during carcinogenesis, HDAC inhibitors have recently gained some traction as anticancer agents [reviewed in [23]]. While HDAC inhibitors are often grouped by the classes of enzymes they target, these drugs also can be classified according to their chemical properties (Table  3). In relation to experimental use, HDAC inhibitors applied to cellular settings are often used to enhance acetylation of target proteins (see below). The most frequently used HDAC inhibitors in the lab are trichostatin A, which can act on all HDACs, and nicotinamide, which inhibits all sirtuin family members. Another salient biochemical feature of lysine acetyltransferases and acetylation modifications (as well as their reversal by HDACs) is illustrated by comparison to kinases and phosphorylation modifications. Kinases phosphorylate specific sites based on consensus sequences present in the target proteins; for example, the PI3K kinase family members phosphorylate threonine or serine in TQ or SQ sequences, respectively. By contrast, different lysine acetyltransferases do not recognize specific amino acid sequences beyond the lysine to be acetylated. Also, acetylation sites tend to be located in structurally ordered regions of proteins, unlike phosphorylation sites

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which are often in unstructured loops [1]. Lysine acetyltransferases apparently rely on structural interactions with their specific protein targets, and these interactions are often mediated by other proteins that form complexes with the core lysine acetyltransferase subunit. Much the same can be said as to how HDACs recognize their specific protein targets. Thus, lysine acetyltransferases and HDACs often exist in complexes that specify their protein targets, and individual acetyltransferases and deacetylases may be present in multiple complexes that determine their different substrates [10]. This is an important concept to consider and adds substantial complexity to investigations into acetylation or deacetylation events in vitro and in cellular systems. For instance, a purified lysine acetyltranferase may modify another protein in solution, but that does not guarantee that the same acetyltransferase performs that specific acetylation in cells. In cellular systems, one needs to consider the possibility that individual lysine acetyltransferases or HDACs might act independently and/or within one or more complexes that could be affected by expression levels or other experimental variables.

3  Methodologies to Study Protein Acetylation Historically, detection and characterization of protein acetylation has been somewhat challenging, especially when compared to some other common posttranslational modifications. For example, ubiquitination modifications add sufficient mass to a protein to change its migration on SDS-PAGE, which is readily monitored by Western blotting with an appropriate antibody to the target protein or ubiquitin moiety. Phosphorylation also often changes the migration of a protein sufficiently such that this modification can again be followed by SDS-PAGE and Western blotting; moreover, use of [32P]-ATP in phosphorylation reactions is a highly sensitive way to monitor these modifications. In contrast, acetylation modifications do not alter protein characteristics sufficiently enough to be detected by changes in migration on standard SDS-­PAGE. Although labeling strategies have been developed to track protein acetylation (see below), issues with sensitivity hindered progress in the field. As mentioned above, broad and overlapping protein substrate specificity of many lysine acetyltransferases and HDACs sometimes makes it difficult not only to investigate often dynamic protein acetylation events but also to specify the r­esponsible enzymes. While these issues must be considered when investigating protein acetylation, appropriate use of antibody-based detection and isolation methods, gene transfection strategies, and HDAC inhibitors has greatly facilitated detection and characterization of individual and pancellular acetylation modifications. Below are brief descriptions of established and relatively new methods to examine these important posttranslational modifications.

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3.1  Detection of Acetylation Events

Conceptually, the easiest way to determine if a protein (or a peptide within a protein) can be acetylated by a candidate acetyltransferase enzyme is by performing acetylation reactions with the required components—i.e., an acetyltransferase (or cell lysate or partially purified fraction containing acetyltransferase activity) with its substrates, target peptide/protein and acetyl-coA (Fig. 1). Notably, catalytically active GCN5 and PCAF acetyltransferases and p300 acetyltransferase fragment as well as a complete histone acetyltransferase kit (containing a fluorescent compound that reacts with CoA-SH product to detect the reaction but does not tag the peptide or protein) are now available commercially (Active Motif, Enzo Life Sciences) for use in these types of reactions and testing of acetyltransferase inhibitors. Determining whether the acetylated peptide or protein has indeed been formed is a detection problem that has been addressed mainly using different labeling strategies or, alternatively, immunodetection methods using antibody specific for acetylated lysine. However, investigators should be aware that biochemical reactions with pure or partly pure reagents can but do not always identify the physiologically relevant lysine acetyltransferase or even the appropriate acetylation sites in the target protein. These details need to be confirmed using experimental strategies carried out in cellular systems and optimally under nonartificial conditions.

3.1.1  Radiolabeling of Target Proteins

An early strategy to detect acetylation of peptides or proteins has been to use radiolabeled acetyl-CoA for biochemical reactions or radiolabeled acetate for experiments performed with cells or isolated nuclei. In fact, incorporation of radiolabeled acetate into histones in isolated nuclei was originally detected in the early 1960s [24, 25]. A typical biochemical reaction is carried out with an acetyltransferase (or lysates or partial purification fractions), and a peptide or protein of interest (in early testing often histones), and initiated by addition of [3H]-acetyl-CoA or [14C]-acetyl-CoA. It is notable that nonenzymatic chemical acetylation of peptides or proteins is also possible, and these products can be used as reference standards [26]. In the standard assay [27], protein from these reactions is adsorbed on phosphocellulose filter paper, with unincorporated acetyl-coA washed away. Incorporation of radiolabel into peptide or protein can be assessed by liquid scintillation counting or autoradiography, although the latter suffers from long exposure times due to the low energy and slow decay of these isotopes. It is noteworthy that an application of this type of labeling reaction in a gel matrix can allow acetylation and labeling of a protein substrate as well as help to identify candidate acetyltransferases [28]. Although these methods are strategically comparable to radioactive labeling of phosphorylated peptides and proteins by kinases, the ability to use much more highly energetic [32P]-ATP as a substrate makes phosphorylation products readily visualized by auto-

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radiography and phosphorimaging and therefore much less time consuming and more convenient. Therefore, alternative labeling strategies to study acetylation events have now been developed that use modified, reactive versions of the CoA substrate to covalently tag protein or peptide targets. 3.1.2  Azide-Alkyne-­ Mediated Identification of Alkylation Sites

In updated methodologies to directly track acetylated proteins, bioorthogonal chemistry methods have been used to attach various chemical moieties to lysines in peptides or proteins in a manner that reflects acetylation [reviewed in [29, 30]]. Notably, recently developed protocols can be performed in vitro or in cellular contexts and allow for nonradioactive fluorescent tagging of targeted lysines in peptides and proteins, while also being compatible with separation and characterization strategies [30]. These strategies replace acetyl-CoA substrate (or acetate precursors of acetyl-CoA) in the acetylation reaction with a modified CoA substrate (or modified precursors) containing a longer alkyl chain with a reactive alkyne moiety at the distal end. Briefly, 4-pentynoyl-CoA (or less efficiently, 5-hexynoyl-CoA) can be incubated with an acetyltransferase and a peptide (or target protein) of interest containing lysine to determine whether it can be an acetylation target for this enzyme. In cellular settings, a similar reaction can be accomplished using alkynyl-acetate analogs (3-butynoate, 4-pentynoate and 5-hexynoate). The covalently linked alkynes (in solution or cell lysates) can then be incubated with fluorescent azides (in the presence of catalyst Cu[I], in so-called copper-catalyzed azide–alkyne 1,3-dipolar cycloaddition reactions) such as azido-rhodamine to generate fluorescently tagged peptides or proteins for subsequent imaging after separation (for example, by SDS-PAGE). This method is not only capable of labeling proteins at potential acetylation sites in biochemical or cellular experiments, but can also be combined with biotin-tagging or immunoprecipitation methods for the protein of interest and/or MS analyses to identify modification sites. A limitation to applying these protocols in living cells is Cu-mediated cytotoxicity.

3.1.3  Immunodetection and Immunoprecipitation

Difficulties with detection and specific binding of acetylated forms of proteins have been somewhat relieved by the availability of antibodies that specifically recognize acetylated lysine. Such antibodies are independently produced and commercially sold by a number of different biotech companies. It should be apparent that although these “pan” acetyl-lysine antibodies (ac-K-Ab) will recognize available epitopes within a wide variety of proteins, it is important to confirm that a specific antibody will interact with the acetylated form of your protein of interest. This can be accomplished by creating a reference using an in vitro reaction with individual components as outlined above. If you already have purified (or partly purified) protein of interest, such reactions can be used

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to generate acetylated versions of proteins that can then be detected using the ac-K-Ab followed by standard immunodetection methodology—i.e., compatible secondary antibody conjugated with an enzyme that converts a substrate to a product emitting chemiluminescence which can be measured using autoradiography or imaging technologies. Depending upon the quality and amount of ac-K-Ab used, this strategy can be quite sensitive for detecting acetylated forms of proteins or peptides. In cases where the acetylated proteins need to be separated from unreacted proteins, immobilization or immunoprecipitation (pull-down) methods employing an ac-K-Ab are useful. When an ac-K-Ab is used as bait (either prior to or after immobilization on an affinity resin bead), only acetylated forms of proteins are bound, leaving unacetylated proteins unbound. Separation techniques such as centrifugation or gel filtration can then be applied to remove unacetylated proteins from bound acetylated proteins that can subsequently be liberated for further analysis. Importantly, such strategies to isolate acetylated proteins can be utilized with purified proteins or complex mixtures containing many potential acetylation targets. Immunoprecipitation of cell or nuclear lysates (or mitochondrial fractions) using a ac-K-Ab followed by SDS-­ PAGE separation and Western blotting of acetylated proteins has permitted proteomic/MS analyses (see below for examples) of excised bands to identify multiple acetylated proteins as well as their specific acetylation sites. While antibodies that bind to acetylated lysines across a broad protein spectrum are very useful, study of acetylation at a defined site in an individual protein would greatly benefit from having an antibody specifically recognizing that acetylated form of the protein. A number of such antibodies are sold commercially, including products that recognize acetylated forms of histones H2A, H2B, H3, and H4, p53 tumor suppressor protein, α-tubulin, Ku70, Foxo1, and Tau. Notably, antibodies specific for acetylated versions of histones have been used to do chromatin immunoprecipitation (ChIP) experiments to identify active gene regions and other associated proteins [31, 32]. If an antibody to a specific acetylation site within your protein of interest is not commercially available, such antibodies can be produced using an appropriate acetylated peptide. However, the peptide must be carefully chosen and synthesized so as not to include nontarget epitopes; once the antibody is raised, it must be purified by adsorption to non-­modified peptide to remove any nonspecific antibodies. While antibodies produced in this manner can be highly specific, one can end up with an antibody having specificity only for acetylated lysine itself, being no more specific than available commercial antibodies that recognize acetylated lysine residues within the broad array of proteins.

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3.2  Strategies to Enrich for Acetylated Proteins in Cellular Systems

When investigating whether intracellular acetylation of protein targets and the possible roles for these modifications, it is often helpful to enrich for the levels of the acetylated proteins to enhance their detection. Since a protein’s acetylation state is determined by a balance between the activities of acetyltransferases and deacetylases, a conceptually straightforward experimental strategy is to tip that balance more toward their acetylated state. The primary strategy to increase the levels of proteins in their acetylated state is to inhibit protein deacetylation. One way to do this is to add deacetylase (HDAC) inhibitors (Table 3) to the media for cell culture experiments. When starting to assess acetylation of one or more protein targets, it is often prudent to use both TSA (~10 μM) and nicotinamide (~5 mM) that act on all HDACs and sirtuin family members, respectively, and thus maximize inhibition of deacetylase activity, leading to accumulation and persistence of the acetylated state of the target protein [6, 9, 33]. Alternatively, if the responsible deacetylase(s) are known, more specific inhibitors can be used that will perhaps reduce unwanted pleiotropic effects on cells. Knowledge of the responsible deacetylase(s) also permits the use of RNA interference methods (siRNA, shRNA) to specifically knockdown the abundance of the responsible deacetylase instead of inhibiting its activity. This alternative strategy of targeting deacetylases again shifts the balance toward higher amounts of the acetylated state of the target protein. Another alternative (or complementary) method to increase the amount of a protein in its acetylated state in a cellular setting is to increase the amount of acetyltransferase activity. This is nearly always done by transfection of an expression vector containing the gene/cDNA encoding the acetyltransferase responsible for the acetylation of the specific protein of interest, although microinjection or electroporation of purified acetyltransferase might also be considered. If the responsible acetyltransferase is not known, researchers often introduce either p300 or CBP acetyltransferase because of the broad spectrum of proteins that they already have been shown to act on. In transfection assays, cell lines (such as HEK293 human embryonic kidney cells) that are more readily transfected are usually used, and often an N- or C-terminal protein tag is included in the expression vector so that the acetyltransferase can be affinity purified or immunoprecipitated to also examine possible association with the target protein or perform other cell-­free assays.

3.3  Proteomics and Mass Spectrometry

Proteomic studies suggest that up to 10% of mammalian proteins can be acetylated [3]. Proteomic strategies encompassing the spectrum of acetylated proteins or focusing on individual proteins have been used to identify acetylated proteins and acetylation sites within these proteins. Identification of acetylation sites by these strategies is critical for mutagenesis-based experiments that can

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further define and confirm the function and importance of individual or multiple acetylation modifications within a target protein. Below we only touch upon common unbiased strategies that are capable of analyzing multiple acetylated proteins simultaneously. However, similar strategies can be used to focus on acetylation of a single or small cohort of acetylated proteins. 3.3.1  Unbiased Proteomic/MS Approaches

A broad strategy initially published by Iwabata and colleagues [34] involves separation of total cellular proteins or purified subcellular fractions by (one- or two-dimensional) SDS-PAGE followed by protein transfer to membranes, with acetylated proteins detected by immunoblotting using an anti-acetylated lysine antibody. Provided sufficient protein is present in bands or spots detected by this method, individual bands can be excised, extracted, digested with proteases, and subsequently subjected to MS analysis. Such a strategy could potentially yield, for each band or spot analyzed, peptides with acetylated residues. The peptide sequences can be derived from MS data and then the acetylated peptide and protein identified by comparison to known protein sequence databases. In cases where the acetylated peptide contains only one lysine, the specific residue that is acetylated can be specified. Another differently staged strategy has been employed by other groups [35, 36] to identify acetylated peptides corresponding to different proteins from mammalian cell systems. Notably, the majority of acetylated proteins identified by this approach were present in mitochondrial fractions. In this approach, cell lysates or subcellular fractions are initially digested with trypsin (or potentially another protease) to yield peptides. Acetylated peptides are then immunoprecipitated using anti-acetylated lysine antibody. This pool of acetylated peptides can then subjected to analysis by tandem chromatography-including nano-HPLC or standard LC-MS technologies, which separate individual peptides and obtain mass fragmentation patterns for each. As above, this information can be compared to protein sequence databases to identify peptides and their respective proteins, as well as specify the acetylated lysine residue in many instances. Other investigators have recently coupled stable-isotope labeling with amino acids in cell culture (SILAC) followed by separating acetylated peptides by isoelectric focusing techniques (instead of SDS-PAGE) with nanoLC-(high-resolution, high accuracy) MS/MS to further expand the ability to detect and identify acetylated proteins [1]. Incorporation of stable isotopic (13C, 14N, 15N, and 2H) versions of lysine and/or arginine into proteins allowed better discrimination during MS, while isoelectric focusing was used after protease digestion and immunoprecipitation with anti-­acetylated lysine antibodies to separated acetylated peptides into 12 separate fractions for subsequent MS analysis. Amazingly, this approach identified 1750 different acetylated proteins with 3600 acetylated sites in human cell lines.

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3.4  Manipulation of Acetylation Sites

Once acetylation sites in a specific protein are established, alteration of the relevant lysine residues should be considered as a means to investigate the effects of acetylation. Mutagenesis strategies are most often used, and are particularly valuable for examining the effects of acetylation of a protein in a cellular or organismal context, even though they are also useful for generating expressed proteins for biochemical studies. Although changes to other amino acids could be considered, mutation of lysine to arginine is almost always preferred to prevent acetylation at specific sites, since arginine cannot be modified by acetyltransferases but retains the positive charge/basic character and is thus more likely to preserve protein structure. Conversely, to closely mimic (constitutive) acetylation at specific sites, mutation of lysine to glutamine is generally performed; this change adds an acetyl group to a linear carbon chain, similar to the structural and electrostatic effect of an acetylated lysine residue. It should be noted that mutations of target lysines to either arginine or glutamine do not always phenocopy the effects of unacetylated or acetylated lysine, respectively, so caution should be exercised in interpreting results from such studies. In the case of multiple possible acetylation sites, different single mutants or multiple site mutant combinations may need to be constructed to fully understand the functions and relevance of each site. Once constructed, these mutants can be introduced into cells (preferably null for the protein to be introduced) by transfection of expression vectors. Again, tags can be introduced N- or C-terminal to the mutant gene to facilitate separation and/or visualization procedures specific for the mutant protein. Acetylation mutants can even be introduced into mouse ES cells to create transgenic mouse models. Using these mutagenesis strategies, investigations into the relevance of acetylation (or deacetylation) at one or more specific sites in a protein has been greatly advanced. In one interesting example, knock-in mice have been created with their p53 gene mutated at multiple lysine residues that are known acetylation sites [37]. As possible alternatives to mutagenesis strategies, other methods (nonsense-suppression and protein ligation or native chemical ligation strategies) can be used to incorporate acetyl-lysine into proteins, although each of these strategies has limitations [4]. Cysteine alkylation chemistry performed on pure protein targets can generate an acetyl-lysine, although this method also will modify cysteine residues. Alternatively, peptides synthesized to specifically include acetyl-lysine can be conjugated to the N- or C-terminus of other peptides or proteins, but this has obvious limitations for examining acetylation sites in the middle of full-length proteins or even large domains. Another possible strategy is purification of target proteins from cells lacking acetylases or deacetylases, respectively, to generate hypo- or hyperacetylated forms of proteins; it is worthwhile to note that this strategy will usually affect multiple

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proteins and perhaps multiple residues within a protein. Alternatively, reactions of purified target proteins with pure acetyltransferases and acetylCoA can be performed, but target proteins are frequently not completely acetylated and/or the relevant in vivo site specificity can be lost. 3.5  Tracking Acetylation in Live Cells

While immunoprecipitation and Western blotting are suitable to monitor collective protein acetylation after cell lysis, these methods are not capable of tracking acetylation in living cells. To address this issue, Minoru Yoshida and colleagues have developed fluorescence resonance energy transfer (FRET) assays that allow visualization and measurement of acetylation in live cells [38, 39]. Their strategy is based on the aforementioned binding affinity between acetylated lysines and bromodomains present in a variety of proteins. Briefly, they designed a reporter construct containing gene segments encoding, from N- to C-terminus, Venus (a color mutant of GFP emitting around 535 nm), a bromodomain, a flexible linker region, the region of histone H4 subject to acetylation, and CFP (a different color mutant of GFP emitting about 480 nm). Theoretically, acetylation of lysines in the histone H4 segment would mediate binding of the bromodomain via an intramolecular conformation change that changes the distance between the fluorescent reporter regions, generating a detectable and quantifiable FRET signal. Upon transfection of the construct, the Yoshida lab was able to determine proper chromatin localization of the construct, monitor generation of FRET signal after incubation of cells with the HDAC inhibitor TSA, and confirm that the FRET signal was specifically mediated through binding between acetylated lysines and the bromodomain using appropriate mutants in those regions [38]. Importantly, this strategy can be used for in vivo testing of acetyltransferases, HDAC inhibitors, and of compounds that interfere with binding of bromodomains and their acetylated lysine targets [38, 39]. On this note, inhibitors that block the acetyl binding pocket of bromodomains are now in cancer clinical trials [4]. This strategy (either using unimolecular or bimolecular constructs) may be adaptable to other acetylation reactions, provided that these events mediate close interactions with specific ­bromodomains or other entities that can be introduced into cells and directed to their site of action.

4  Structured Plan to Initiate Investigation of Acetylation of a Specific Protein With the information above in mind, a plan can be formed to investigate intracellular acetylation (and deacetylation) of a specific protein target. Importantly, this plan is dependent upon having a specific and reasonably sensitive antibody to the target protein of interest which binds both the unmodified and acetylated form(s).

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Strategies may also need to be modified based on the abundance of the target protein and lysine acetyltransferase(s) responsible for its acetylation. First, it is prudent to carry out experiments with your chosen cell line after preincubation with the HDAC inhibitors TSA and nicotinamide at concentrations sufficient to completely inhibit all deacetylase activity, thereby maximizing acetylation of your protein target to improve your chances of detecting the acetylated form. After one or more intervals of deacetylase inhibition, cell lysates can be prepared by methods that maintain the solubility of the protein of interest. Since acetylation state could affect the localization of the protein, it is best to use whole cell lysates initially, although comparing subcellular fractions could be considered at this stage or later to identify the compartment in which the acetylated protein resides. Immunoprecipitation with antibody specific for your protein is performed on the cell lysate, with immunoprecipitated (and supernatant) fractions run in separate lanes on SDS-­ PAGE along with a measured amount of the original cell lysate (input), appropriate molecular weight markers and, if available, a purified (unacetylated) protein reference. In initial experiments, relatively high amounts of total protein should be loaded for the immunoprecipitated and supernatant fraction lanes to maximize detection. The proteins are then transferred to a support membrane for standard Western blotting techniques using an ac-K-Ab that will hopefully detect the acetylated protein of interest as well as perhaps other acetylated proteins that might have co-­ immunoprecipitated with your target protein and thus be present on the membrane. Detection of a band associated with the immunoprecipitated fraction at the molecular weight of the protein of interest strongly suggests that some of the target protein was acetylated under the conditions of this experiment. If a band is present in the supernatant fraction at the same molecular weight, caution is urged as there are several explanations: (1) the band represents a different acetylated protein of similar molecular weight that would not be immunoprecipitated by the antibody, (2) the antibody used for immunoprecipitation did not work on the acetylated form of the target protein, or (3) was of insufficient concentration to (wholly) precipitate the targeted protein. After the initial Western is analyzed, the membrane can be stripped and reprobed with antibody against the (non-acetylated) form of the target protein. Bands should clearly be present in the reference, input, and immunoprecipitated fraction; a band in the supernatant indicates that the immunoprecipitation antibody was insufficient to pull down all of the target protein. Comparison of these blots should determine whether the cellular conditions yielded an acetylated form of the protein of interest and the immunoprecipitation methods used were successful in pulling down this modified protein. Notably, although this procedure could be reversed, using the ac-K-Ab for

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immunoprecipitation with Western blotting with the antibody specific to your protein, it is subject to a potential misleading result— i.e., a band detected at the proper molecular weight could be the unmodified target protein that co-immunoprecipitated with a different acetylated protein pulled down by the ac-K-Ab. Importantly, this possibility can be eliminated using stringent immunoprecipitation conditions that abrogate protein-protein associations. Once it is clear that the protein of interest is indeed acetylated under cellular conditions favoring acetylation and using optimized immunoprecipitation methods, other questions pertaining to these acetylation events can be addressed, including: (1) using various HDAC inhibitors to determine which deacetylase (s) are responsible for removing acetyl groups from the protein, (2) using acetylase inhibitors and/or transfection strategies to identify the acetyltransferases responsible for protein acetylation, (3) exploring other cellular conditions that impact acetylation state of the protein, (4) determining how acetylation and deacetylation affect trafficking and localization of the protein, (5) determining how acetylation state influences macromolecular interactions of the target protein, and (6) scaling up ac-K-Ab immunoprecipitation reactions to acquire sufficient amounts of acetylated protein for proteomic-MS analysis to identify acetylation sites. In instances where acetylation of the target protein is still suspected but cannot be detected readily under the conditions outlined above, it may be necessary to increase expression of the target protein and/or lysine acetyltransferases using transfection strategies. Again it may be useful to include, in these expression vectors, tags N- or C-terminal to the genes for these factors to facilitate visualization and/or affinity purification strategies that assist in further characterization. If these overexpression strategies are used initially to help determine whether your protein is acetylated and/ or to identify acetylation sites by proteomic methods, it is best to confirm that your endogenous target protein is acetylated on the identical sites by the same lysine alkyltransferase. Confirmation of the responsible lysine acetyltransferase is best accomplished using RNA interference techniques, although use of specific acetyltransferase inhibitors if available might be considered. Ideally, ­confirmation of the authenticity of specific acetylation sites identified by overexpression strategies needs to be accomplished by mutagenizing the sites and reintroducing these constructs to determine their effect on protein function in an appropriate cell background (null or knocked down for the unmutated protein) that is otherwise relatively non-perturbed. Although other strategies and specific experiments to analyze protein acetylation can be envisioned, this plan provides a general guide for investigators to determine whether their protein is acetylated as well as the specific acetylation sites and potentially generates tools or reagents to characterize acetylation events in cells and in biochemical tests.

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5  Summary Acetylation of histone and nonhistone proteins is an important posttranslational modification that impacts numerous cellular events. These modifications most prominently affect chromatin structure and the epigenetic and transcriptional status of the genome, but also influence, among other processes, DNA replication and repair, sister chromatid cohesion, cellular metabolism, and cytoskeletal dynamics. Acetylation of specific lysines in target proteins is carried out by various lysine acetyltransferases (Table 1), while deacetylation is performed by HDACs (Table 2). The acetylation state of an individual protein is determined by the balance between its acetylation and deacetylation, which can be influenced by using readily available HDAC inhibitors (Table 3) for experimental purposes to enhance the acetylated state to facilitate identification and further characterization. The existence of HDAC inhibitors, specific antibodies to acetylated lysine moieties and increasingly to acetylated forms of specific proteins, and proteomic/MS technologies has augmented the number of tools and strategies available to study these modifications and facilitated detection of acetylation events across biological systems. As a consequence, appreciation of the number of proteins subject to acetylation (and the roles of these modifications) in humans and other organisms is steadily increasing, ensuring that acetylation modifications of proteins will continue to be a subject of vigorous investigation.

Acknowledgment This work was supported by NIH grants R01 AG027258 and R01 AG026534 as well as by the Department of Toxicology and Cancer Biology of the University of Kentucky College of Medicine. References 1. Choudhary C, Kumar C, Gnad F, Nielsen ML, Rehman M, Walther TC, Olsen JV, Mann M (2009) Lysine acetylation targets protein complexes and co-regulates major cellular functions. Science 325:834–840 2. Zhou L, Zeng Y, Li H, Li Y, Shi J, An W, Hancock SM, He F, Qin L, Chin J, Yang P, Chen X, Lei Q, Xiong Y, Guan KL (2010) Regulation of cellular metabolism by protein lysine acetylation. Science 327:1000–1004 3. Kim GW, Yang XJ (2011) Comprehensive lysine acetylomes emerging from bacteria to humans. Trends Biochem Sci 36:211–220

4. Dancy BM, Cole PA (2015) Protein lysine acetylation by p300/CBP. Chem Rev 115:2419–2432 5. Sadoul K, Boyault C, Pabion M, Khochbin S (2008) Regulation of protein turnover by acetyltransferases and deacetylases. Biochimie 90:306–312 6. Li K, Wang R, Lozada E, Fan W, Orren DK, Luo J (2010) Acetylation of WRN regulates it stability by inhibiting ubiquitination. PLoS One 5:e10341 7. Blander G, Zalle N, Daniely Y, Taplick J, Gray MD, Oren M (2002) DNA damage-induced

Characterization of Protein Acetylation translocation of the Werner helicase is regulated by acetylation. J Biol Chem 277:50934–50940 8. Karmakar P, Bohr VA (2005) Cellular dynamics and modulation of WRN protein is DNA damage specific. Mech Ageing Dev 126:1146–1158 9. Li K, Casta A, Wang R, Lozada E, Fan W, Kane S, Ge Q, Orren D, Luo J (2008) Regulation of WRN protein cellular localization and enzymatic activities by SIRT1-mediated deacetylation. J Biol Chem 283:7590–7598 10. Yang XJ, Seto E (2007) HATs and HDACs: from structure, function and regulation to novel strategies for therapy and prevention. Oncogene 26:5310–5318 11. Drazic A, Myklebust LM, Ree R, Arnesen T (2016) The world of protein acetylation. Biochim Biophys Acta 1864:1372–1401 12. Allis CD, Berger SL, Cote J, Dent S, Jenuwein T, Kouzarides T, Pillus L, Reinberg D, Shi Y, Shiekhattar R, Shilatifard A, Workman J, Zhang Y (2007) New nomenclature for chromatin-­ modifying enzymes. Cell 131:633–636 13. Gong F, Miller KM (2013) Mammalian DNA repair: HATs and HDACs make their mark through histone acetylation. Mutat Res 750:23–30 14. Yang XJ (2015) MOZ and MORF acetyltransferases: molecular interaction, animal development and human disease. Biochim Biophys Acta 1853:1818–1826 15. Lee KK, Workman JL (2007) Histone acetyltransferase complexes: one size doesn’t fit all. Nat Rev Mol Cell Biol 8:284–295 16. Liu N, Li S, Wu N, Cho KS (2017) Acetylation and deacetylation in cancer stem-like cells. Oncotarget 8:89315–89325 17. Kalkhoven E (2004) CBP and p300: HATs for different occasions. Biochem Pharmacol 68:1145–1155 18. Goodman RH, Smolik S (2000) CBP/p300 in cell growth, transformation, and development. Genes Dev 14:1553–1577 19. Vo N, Goodman RH (2001) CREB-binding protein and p300 in transcriptional regulation. J Biol Chem 276:13505–13508 20. Yao TP, Oh SP, Fuchs M, Zhou ND, Chang LE, Newsome D et al (1998) Gene dosage-­ dependent embryonic development and proliferation defects in mice lacking the transcriptional integrator p300. Cell 93:361–372 21. Rebel VI, Kung AL, Tanner EA, Yang H, Bronson RT, Livingston DM (2002) Distinct roles for CREB-binding protein and p300 in

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mice by use of anti-acetyllysine and -methyllysine mouse monoclonal antibodies. Proteomics 5:4653–4664 35. Kim SC, Sprung R, Chen Y, Xu Y, Ball H, Pei J, Cheng T, Kho Y, Xiao H, Xiao L, Grishin NV, White M, Yang XJ, Zhao Y (2006) Substrate and functional diversity of lysine acetylation revealed by a proteomics survey. Mol Cell 23:607–618 36. Zhao S, Xu W, Jiang W, Yu W, Lin Y, Zhang T, Yao J, Zhou L, Zeng Y, Li H, Li Y, Shi J, An W, Hancock SM, He F, Qin L, Chin J, Yang P, Chen X, Lei Q, Xiong Y, Guan KL (2010) Regulation of cellular metabolism by protein lysine acetylation. Science 327: 1000–1004 37. Li T, Kon N, Jiang L, Tan M, Ludwig T, Zhao Y, Baer R, Gu W (2006) Tumor suppression

in the absence of p53-mediated cell-­ cycle arrest, apoptosis, and senescence. Cell 149: 1269–1283 38. Sasaki K, Ito T, Nishino N, Khochbin S, Yoshida M (2009) Real-time imaging of histone H$ hyperacetylation in living cells. Proc Natl Acad Sci U S A 106:16257–16262 39. Ito T, Umehara T, Sasaki K, Nakamura Y, Nishino N, Terada T, Shirouzu M, Padmanabhan B, Yokoyama S, Ito A, Yoshida M (2011) Real-time imaging of histone H4K12-specific acetylation determines the modes of action of histone deacetylase and bromodomain inhibitors. Chem Biol 18: 495–507 40. West AC, Johnstone RW (2014) New and emerging HDAC inhibitors for cancer treatment. J Clin Invest 124:30–39

Part III Sirtuin Targets of Acetylation

Chapter 8 Molecular and Cellular Characterization of SIRT1 Allosteric Activators Michael B. Schultz, Conrad Rinaldi, Yuancheng Lu, João A. Amorim, and David A. Sinclair Abstract SIRT1 is an NAD+-dependent lysine deacetylase that promotes healthy aging and longevity in diverse organisms. Small molecule allosteric activators of SIRT1 such as resveratrol and SRT2104 directly bind to the N-terminus of SIRT1 and lower the Km for the protein substrate. In rodents, sirtuin-activating compounds (STACs) protect from age-related diseases and extend life span. In human clinical trials, STACs have a high safety profile and anti-inflammatory activities. Here, we describe methods for identifying and characterizing STACs, including production of recombinant protein, in vitro assays with recombinant protein, and cellular assays based on mitochondrial dynamics. The methods described in this chapter will facilitate this discovery of improved STACs, natural and synthetic, in the pursuit of interventions to treat age-related diseases. Key words Sirtuin, SIRT1, Histone deacetylase (HDAC), Deacylase, Deacetylase, NAD+, Nicotinamide, ADP-ribose, Aging, Longevity, Metabolism, Epigenetics, Histone, p53, Mitochondria, Membrane potential, Reactive oxygen species (ROS), Sirtuin-activating compound (STAC), Resveratrol, Allosteric activator, Recombinant protein

1  Introduction Since the discovery of the yeast longevity gene Silent Information Regulator 2 (SIR2), much attention has been given to its seven mammalian homologs, the sirtuins [1]. Of these, SIRT1 is the closest homolog to its yeast counterpart and, along with SIRT6, is one of only two sirtuins shown to extend life span in mammals [2]. SIRT1, an NAD+-dependent lysine deacetylase, serves as an NAD+sensor that promotes efficient energy utilization and cellular defenses in response to changes in the environment such a decline in nutrient availability. SIRT1 has numerous deacetylation targets involved in key biological processes, including histones [3], transcription factors like p53 [4, 5], PGC-1α [6], and NF-κB [7], signaling proteins such as the Notch intracellular domain Robert M. Brosh, Jr. (ed.), Protein Acetylation: Methods and Protocols, Methods in Molecular Biology, vol. 1983, https://doi.org/10.1007/978-1-4939-9434-2_8, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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(NICD) [8], and insulin receptor substrate-2 (IRS2) [9], and enzymes such as LKB1 [10]. The net effect of increased SIRT1 activity is to send a cell into “survival mode,” with improved DNA repair, epigenetic stability, and metabolic efficiency. SIRT1 promotes healthy aging and longevity, while its dysregulation accelerates many age-related diseases such as Alzheimer’s disease, cancer, cardiovascular disease, and diabetes [11]. Because of SIRT1’s salutary effects, it is an enticing target for pharmacological activation. In comparison to enzyme inhibition, discovering an activator is a rarity; it is easier to throw a wrench in the works than to enhance a machine’s function. Luckily, SIRT1 is the rare enzyme with activation potential. It possesses a large, loosely structured N-terminus called the STAC binding domain (SBD), the only mammalian sirtuin with such a domain. While not required for the enzyme’s activity, the SBD increase SIRT1 activity by stabilizing the interaction of the domain with its acetylated protein targets. STACs allosterically bind to this N-terminus, further stabilizing the interaction between SIRT1 and its targets beyond normal levels, in effect lowering the apparent Km of the enzyme [12]. It is not known if this opportunity for allosteric activation is an accident of biochemistry or a product of natural selection. If an endogenous small molecule SIRT1 activator exists in mammals, it has yet to be discovered. The first discovery of SIRT1 allosteric activators came in 2003 with the identification of resveratrol and similarly structured polyphenols [13]. The first SIRT1-mediated activities described for these molecules were increased mitochondrial function and protection from the negative effects of a high fat diet [10, 14]. Since 2003, synthetic molecules have been discovered with much higher affinity for SIRT1. Two such molecules, SRT1720 and SRT2104, extend healthspan and longevity in mice [15, 16]. In humans, over 50 clinical trials have been carried out with resveratrol or SRT2104, with a focus on neuropathies, cardiovascular disease, inflammation, and diabetes. Considering resveratrol’s less-than ideal pharmacokinetic properties and the variability of human patients, clinical results have been mixed. A recent meta-analysis concluded that resveratrol supplementation has positive effects on multiple health parameters in type 2 diabetes patients, with no adverse effects over placebo [17]. Similarly, SRT2104 has been shown to reduce disease severity in a subset of psoriasis patients when compared to placebo in a randomized double-blind trial [18]. New SIRT1 activating compounds with improved pharmacokinetic and pharmacodynamic profiles are an area of intense research interest. Here, we describe methods for the discovery and characterization of STACs. First, we describe a method for producing recombinant SIRT1 enzyme for biochemical assays [19]. In addition to producing the wild-type enzyme, it is also possible to produce mutant versions of SIRT1. For example, SIRT1-H363Y is

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e­ nzymatically dead and SIRT1-E230K is resistant to allosteric activation [12]. This protocol is also used to produce recombinant yPNC1, which is required for a subsequent assay. Next, we describe the Fluor-de-Lys assay for measuring SIRT1 activity in vitro [20]. SIRT1 is incubated with an acetylated substrate, NAD+, and the test STAC. The substrate has a quenched fluorophore proximal to its acetylated lysine such that the removal of the acetyl group by SIRT1 allows the lysine to be cleaved by trypsin in the subsequent development stage, thereby freeing and un-quenching the fluorophore (Fig. 1). Fluorescence is therefore proportional to the amount of deacetylated substrate due to SIRT1 activity. This assay has been a focus of controversy, as some have argued that the activity of SIRT1 on a fluorophore-tagged substrate does not reflect its activity on endogenous substrates [21]. However, further investigation has shown that the hydrophobic fluorophore in this assay mimics natural, bulky, hydrophobic residues such as tryptophan and phenylalanine that are present in endogenous substrates [12]. STACs identified in this assay have proved robust in other in vitro and in vivo contexts. Another assay for measuring SIRT1 activity in vitro is the PNC1-OPT assay [22] (Fig. 2). This assay is designed for using a native peptide (without a fluorophore) as the SIRT1 substrate. It measures the amount of nicotinamide, another product produced in the SIRT1 deacetylation reaction. SIRT1 activators will increase the production of nicotinamide in step (1). yPnc1, an enzyme from budding yeast, will then convert the nicotinamide into nicotinic acid and NH4+ (free ammonia) in step (2). Finally in step (3), the ammonia will react with ortho-phthalaldehyde (OPT) and dithiothreitol (DTT) to generate a fluorescent product—1-­ alkylthio-­substituted isoindoles, which can emit fluorescence near 476 nm once excited. Therefore, the fluorescence detected is proportional to the amount of nicotinamide produced and is thereby representative of SIRT1 activity. Finally, we describe methods for measuring STACs’ effects on mitochondrial activity in cells. Resveratrol, SRT1720, and SRT2104 have all been shown to promote mitochondrial biogenesis and function and to suppress the production of reactive oxygen species (ROS) [10, 23, 24]. As mitochondrial content and activity vary with diet, exercise, aging, and disease status, these methods are a strategic starting place for assessing novel STACs’ efficacy in vivo. Here, we present relatively simple methods for quantifying mitochondrial mass, membrane potential, and ROS production based on the use of fluorescent dyes and flow cytometry. Four dyes are utilized in this assay: mitotracker deep red (MitoTracker® Deep Red FM) and nonyl-acridine orange (NAO) for measuring mitochondrial mass; tetramethylrhodamine methyl ester (TMRM) for measuring mitochondrial membrane potential; and dihydroethidium (DHE) measuring ROS.

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Fig. 1 Schematic of the Fluor-de-Lys assay

Fig. 2 Schematic of the PNC1-OPT assay. A potential SIRT1 activator is added in step (1) and its effect on the SIRT1 deacetylation reaction will be revealed in step (3)

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2  Materials 2.1  Expression and Purification of Recombinant SIRT1

1. pET-based His-tagged SIRT1 plasmid. 2. BL21 pLysS(DE3), Rosetta, or other competent bacteria for protein expression. 3. LB media and LB-agar plates with antibiotic. 4. Bacterial incubator. 5. Centrifuge for spinning down large volumes of bacterial cultures. 6. Protease inhibitors (EDTA free). 7. Isopropyl beta-d-1-thiogalactopyranoside (IPTG). 8. Sonicator. 9. Ni-NTA agarose beads. 10. Lysis buffer

(a) 1% Triton X-100.



(b) 50 mM Tris pH 8.0.



(c) 150 mM NaCl.



(d) 20 mM imidazole.



(e) 3 mM beta-mercaptoethanol (BME).

11. Wash buffer

(a) 1% Triton X-100.



(b) 50 mM Tris pH 8.0.



(c) 300 mM NaCl.



(d) 20 mM imidazole.



(e) 3 mM BME.

12. Elution buffer

(a) 50 mM Tris pH 8.0.



(b) 250 mM imidazole.



(c) 3 mM BME.

13. Spectrophotometer for measuring absorbance. 14. Optional: Bio-Rad Poly-Prep® columns. 15. Optional: Millipore Microcon® dialysis columns. 2.2  Fluor de Lys Assay for SIRT1 Activators In Vitro

All of the following materials are available in the SIRT1 Fluorometric Drug Discovery Kit (Enzo® Life Sciences) 1. SIRT1 (Sirtuin 1, hSir2SIRT1) (human, recombinant) in 25 mM Tris, pH 7.5, 100 mM sodium chloride, 5 mM dithiothreitol, and 10% glycerol. Store at −80 °C.

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2. Fluro de Lys® SIRT1, Deacetylase Substrate (100 μL; 5 mM solution in 50 mM Tris/Cl, pH 8.0, 137 mM sodium ­chloride, 2.7 mM potassium chloride, 1 mM magnesium chloride). Store at −80 °C. 3. Fluor-de-Lys® Developer II Concentrate (5×) (5 × 250 μL; 5× Stock Solution; Dilute in Assay Buffer before use). Store at −80 °C. 4. NAD+ (Sirtuin Substrate) (500 μL; 50 mM β-Nicotinamide adenine dinucleotide (oxidized form) in 50 mM Tris/Cl, pH 8.0, 137 mM sodium chloride, 2.7 mM potassium chloride, 1 mM magnesium chloride). Store at −80 °C. 5. Nicotinamide (Sirtuin Inhibitor) (500 μL; 50 mM Nicotinamide in 50 mM Tris/Cl, pH 8.0, 137 mM sodium chloride, 2.7 mM potassium chloride, 1 mM magnesium chloride). Store at −80 °C. 6. Resveratrol (Sirtuin Activator) (10 mg; Solid MW: 228.2, soluble in DMSO or 100% ethanol (to 100 mM)). Store at −80 °C. 7. Suramin sodium (Sirtuin Inhibitor) (10 mg; Solid MW: 1429.2, soluble in water or assay buffer (to 25 mM)). Store at −80 °C. 8. Fluor-de-Lys® Deacetylated Standard (30 μL; 10 mM in DMSO). Store at −80 °C. 9. Sirtuin Assay Buffer (50 mM Tris/Cl, pH 8.0, 137 mM sodium chloride, 2.7 mM potassium chloride, 1 mM magnesium chloride, 1 mg/mL bovine serum albumin) (20 mL). Store at −80 °C. 10. 96-well white ½ area plates for fluorometry. 11. Spectrophotometer with fluorescence capabilities and appropriate filters (excitation 350–380 nm and emission 450–480 nm). 2.3  PNC1-OPT Assay for SIRT1 Activators In Vitro

1. Purified recombinant SIRT1 and yPnc1 enzymes should be expressed by using the protocol in the same chapter [18]. 2. SIRT1 requires acetylated peptide substrates, the peptide length is usually 5–15 amino acids and it has acetylated lysine near the middle of the sequence (e.g., Ac-TARK(ac)STG-­NH2) (see Note 1). Peptides with hydrophobic groups adjacent to the acetyl-lysine are recommended for STAC-mediated SIRT1 activation [12]. 3. Reaction Buffer: PBS (pH 7.4) or Tris buffer (pH = 8.0) supplemented with 1 mM DTT (Thermo Fisher, R0861). 4. OPT Developer Reagent: To make this 30% ethanol/70% PBS (pH 7.4) solution supplemented with 10 mM OPT and

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10 mM DTT, first dissolve OPT (Sigma, P0657) in pure ethanol (33.3 mM OPT) and DTT in PBS (14.28 mM DTT), respectively, then mix 3 volumes of ethanol (with OPT) to 7 volumes of PBS (with DTT). Protect from light and store at −20 °C until use. 5. NAD+ (Sigma, N7004) should be dissolved in distilled water to make 100 mM aliquots and stored at −20 °C. Avoid freeze-­ thaw cycles. 6. Nicotinamide (Sigma, 72340) is used to make standard curve, prepare its 100 mM stock solution in distilled water, aliquoted and stored at −20 °C. 7. 96-well black polystyrene plates (Corning, 3915) suitable for fluorometry. 8. Plate reader capable Em = 413/476 nm).

of

reading

fluorescence

(Ex/

9. Aluminum foil. 10. 37 °C incubator. 11. Orbital shaker. 2.4  Mitochondrial Assays for SIRT1 Activators in Cells

1. Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum, 2 mM l-glutamine, and 1% antibiotic: penicillin/streptomycin. 2. Phosphate-buffered saline (PBS) (GIBCO). 3. Dimethyl sulfoxide (DMSO). 4. 6-Well Cell Culture plate (2 mL volume per well). 5. Mouse Embryonic Fibroblasts (MEFs) and/or C2C12 cells. 6. 12 × 75 mm round bottom polystyrene tubes. 7. Tetramethylrhodamine methyl ester (TMRM perchlorate, Invitrogen T-668), a light-sensitive cationic, mitochondrial membrane-permeable fluorescent dye, dissolved in DMSO at 100–500 times greater concentration than the final concentration used in the assay. The concentration used in the assay varies depending on the mode used (see Note 2). Cells are equilibrated with the dye in the incubation media for 45–60 min. TMRM displays excitation/emission spectra of 552/575 nm. 8. Dihydroethidium (DHE, Sigma D7008), a cell-permeable fluorescent dye, is extensively used in tissue culture experiments to evaluate ROS. DHE is dissolved in DMSO at 100– 500 times greater concentration than the final concentration used in the assay, protected from the light. Cells are equilibrated with the dye in the incubation media for 30–60 min. It displays excitation/emission spectra of 490/590 nm.

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9. Nonyl acridine orange (NAO, Acridine Orange 10-Nonyl Bromide, Invitrogen A1372), a light-sensitive metachromatic dye which binds to cardiolipin, a phospholipid specifically present on the mitochondrial membrane, should be dissolved in DMSO at 100–500 times greater concentration than the working concentration (see Note 3). Cells are equilibrated with the dye in the incubation media for 30–60 min and its excitation/ emission spectra of 495/538 nm. 10. MitoTracker® Deep Red FM (Invitrogen M22426), a light-­ sensitive dye that passively diffuses across membranes and accumulates in active mitochondria, and is well retained after aldehyde fixation or after permeabilization with detergents in subsequent steps. This dye, which is not easily washed out of cells once the mitochondria experience a loss in membrane potential, is dissolved in DMSO at 100–500 times greater concentration than the final concentration used in the assay. Cells are equilibrated with the dye in the incubation media for 30–60 min. It displays excitation/emission spectra of 644/665 nm. 11. Carbonyl cyanide p-(trifluoromethoxy) phenylhydrazone (FCCP) (Sigma C2920), dissolved in ethanol, 1 mM stock solution. 12. Antimycin A (Sigma A8674), dissolved in ethanol, 1 mM stock solution. 13. Hydrogen peroxide (Sigma H1009), 30% w/w.

3  Methods 3.1  Expression and Purification of Recombinant SIRT1

1. Day 1: Transform competent bacterial cells with the His-­tagged SIRT1 plasmid, according to manufacturer’s instructions. 2. Grow cells on LB-agar plates with appropriate antibiotic overnight at 37 °C. 3. Day 2: Pick a colony from the plate and inoculate a tube with 5 mL of LB media with antibiotic. Grow overnight at 37 °C shaking at 225 rpm. 4. Day 3: Add the 5 mL culture to 2 L of LB media with antibiotic. Grow at 37 °C shaking at 225 rpm, taking optical density (OD) reading occasionally using the spectrophotometer, until an OD600 of 0.6 is reached. 5. Add IPTG to a final concentration of 1 mM to induce recombinant protein expression. 6. Shake culture at 16 °C at 225 rpm overnight. 7. Day 4: Spin down bacterial culture in a centrifuge (e.g., 4633 × g for 20 min at 4 °C), and pour off the supernatant.

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8. Add protease inhibitors (without EDTA) to lysis and wash buffers (see Note 4). Keep the solutions on ice. 9. Add 50–200 mL of lysis buffer to the cell pellet. Resuspend the pellet thorough by pipetting up and down. 10. Let the solution sit on ice for about 30 min until it becomes viscous, which can be monitored by pipetting a small amount of the mixture. 11. Sonicate the solution on ice for 30 s, 5 times, with 1-min cool-­ downs on ice in between sonication steps. 12. Centrifuge the samples at 30,000 × g (16,000 rpm using a Sorval SS34 rotor) at 4 °C for 30 min to pellet the cell debris. 13. Meanwhile, prepare the Ni-NTA agarose beads. Use 1.5 mL of slurry (50% packed beads) per liter of bacteria, for approximately 3 mL slurry. Briefly spin down the slurry at a low speed (~100 × g) and aspirate off the supernatant. Wash the beads 2× with 10 mL cold lysis buffer, briefly spinning and removing the supernatant between washes. 14. Add the sample supernatant to the washed beads. Rotate at 4 °C for 1–2 h. 15. Centrifuge the sample at 100 × g for 1 min to pellet the beads. Aspirate and discard the supernatant. 16. Add twice the bead volume of cold wash buffer. Resuspend, spin, and remove the supernatant. Repeat this for a total of at least five wash steps. 17. Add 1.5 times the bead volume of elution buffer. Resuspend and rotate for 1 h at 4 °C. 18. Remove the beads by spinning at (100 × g). Keep the supernatant. 19. Optional: Transfer the supernatant to a Polyprep column. Filter the supernatant by gravity through the column into a collector tube, according to manufacturer’s instructions. 20. Optional: Dialysis with a Millipore Microcon column may be performed according to manufacturer’s instructions to remove imidazole. 21. Dilute the protein 1:1 with glycerol, aliquot, and freeze at −20 °C. 3.2  Fluor de-Lys Assay for SIRT1 Activators In Vitro 3.2.1  Preparing Reagents

1. All kit components, as well as all dilutions of components, should be defrosted on ice until use. Aliquot the volume of needed reagent so the reagents and plates can be pre-warmed to 37 °C prior to the first incubation (see Note 5). 2. Include an extra number of wells when calculating the volume of reagents needed to account for pipetting error or bubbles. You will need a minimum 10 μL of each reagent per well.

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3. A typical experiment will have each reagent with a working concentration at 3× since the wells will contain three separate parts (SIRT1, compound of interest and NAD+/Substrate) at equal volumes (30 μL total). 4. Dilute the SIRT1 in assay buffer to 3× (0.06 U/μL). 5. Prepare 500 μL dilutions in assay buffer of 3× (300 μM) resveratrol and suramin and aliquot five 100 μL tubes to be used for up to three freeze-thaw cycles each. 6. NAD+/Substrate should be combined into 1 master mix of assay buffer at 3× (75 μM) each for ease of measuring and pipetting (see Note 6). 7. Final concentrations of water, DMSO, or ethanol >1% can affect the SIRT1 activity, so all compounds should be resuspended in assay buffer, when possible. 8. Within 15 min of use, dilute the 5× developer into assay buffer with nicotinamide. The 1 mL of working concentration developer will contain 760 μL assay buffer, 200 μL 5× developer, and 40 μL of 50 mM (stock concentration) nicotinamide. You will need 30 μL of developer per well. 1. Table 1 is a scheme of what a typical experiment will look like with the Fluor de Lys assay.

3.2.2  Performing the Assay

2. Prepare the solutions as described above and put on ice. 3. Place the plate and solutions of SIRT1, resveratrol, suramin, NAD+/Substrate, and test compounds in a 37 °C incubator to pre-warm for 10 min (see Note 7). 4. Add 10–20 μL of assay buffer to the appropriate wells. 5. Add 10 μL of the 3× resveratrol, suramin, or the test compounds to their respective wells. 6. Add 10 μL of the 3× SIRT1 to all of the wells, except for the No Enzyme wells (see Note 8). Table 1 Example of assay mixtures (per well volume)

Sample

Assay buffer (μL)

SIRT1 (3×) (μL)

Amount of sample (3×) (μL)

NAD+/substrate (3×) (μL)

No enzyme

20

0

0

10

Control

10

10

0

10

Resveratrol

0

10

10

10

Suramin

0

10

10

10

Test compound

0

10

10

10

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7. At this step, you can either immediately move on, or wait 5–10 min for the compounds to interact with the SIRT1. 8. Add 10 μL of the 3× NAD+/Substrate to all wells. This must be done last, as it will start the reaction. 9. Cover the plate with foil and incubate at 37 °C for the desired length of time (30–60 min). 10. Once there is ~10 min in the incubation left, quickly thaw the 5× developer and nicotinamide and make the 1× developer mixture. 11. Add 30 μL of the 1× developer to all wells (see Note 9). 12. Recover the plate with foil and leave at RT for at least 15 min before reading. (30–60 min will have the reaction fully stopped and at max signal.) 13. Read the plate in a top-reading microplate fluorometer with an excitation of 360 nm and emission reading of 460 nm (see Note 10). 3.3  PNC1-OPT Assay for SIRT1 Activators In Vitro 3.3.1  Setup and Reaction (Involves Steps 1 and 2)

1. Dilute 100 mM nicotinamide stock solution to make 0, 5, 10, 20, 30, 40, and 50 μM standards. Pipette 1 μL of each standard into its respective well of a 96-well plate. (The final nicotinamide concentrations will be 100-fold further diluted.) 2. Add 1 μg yPNC1 with 100 μL of the reaction buffer to each standard well and mix by pipetting. 3. Prepare a master-mix on ice. Since an extra set of reactions will be used to account for background fluorescence, the matermix volume should be enough for assaying two times the number of samples in triplicates. Each reaction needs the following: 10–30  μM acetylated peptide substrate, ~1–2 μg yPnc1 enzyme, ~1–2 μg SIRT1 enzyme (see Note 11) and reaction buffer (100 μL minus the volume of other components). Mix by gentle vortexing. 4. Split the master-mix into two tubes. Add 75–200 μM final concentration of NAD+ to one tube and add an equal volume of water to another tube (see Note 12). Mix again by gentle vortexing. 5. For each compound sample to be tested, apply 100 μL of master-­mix without NAD+ to three wells, and 100 μL master-­ mix with NAD+ to three wells (all performed in triplicate). Then the same amount of test compound, such as a potential SIRT1 activator, can be added into each well. Test compounds must not alter yPnc1 activity or diminish the fluorescence signal. This can be discerned by incubating compound with a nicotinamide standard curve in this assay. Add equal volume of solvent (e.g. DMSO, the solvent for resveratrol) into control wells (see Note 13). Mix by pipetting.

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6. Gently tap the plate to mix the solutions, and cover with a plate sealer. Incubate at 37 °C on a plate shaker with gentle agitation for 1 h. Longer incubation times may be needed if: (1) the enzyme has lower activity; (2) the activity on the assayed substrate is weak; or (3) the nicotinamide standard curve doses are very high. 3.3.2  Development (Step 3)

1. OPT Developer Reagent should be warm at 37 or 42 °C for 15 min (avoid light) before developing. Vortex if precipitation of DTT is observed. 2. Following the incubation period, add 100 μL of OPT Developer Reagent into each well quickly under dimly lit or dark conditions with a multichannel pipette. Gently tap to mix the solutions, then use plate sealer to cover the 96-wells and wrap it with aluminum foil. Put the plate on a plate shaker in room temperature for 1 h with gentle agitation. 3. Read the fluorescence on a spectrophotometer under dim light. A 0.1–1 s read time is recommended. Filters set to excitation ~420 nm (±10), and emission ~460 nm (±20).

3.3.3  Analysis

1. Use the nicotinamide standards to plot a standard curve of fluorescence intensity versus nicotinamide concentration. 2. Calculate the net fluorescence for each reaction condition by subtracting the mean fluorescence of the background control reactions (no NAD+) from the experimental reaction (with NAD+), F-corrected = F + NAD–F − NAD control (mean value). If high dose of NAD+ is used, performing parallel reactions with and without SIRT1 enzyme will be more appropriate. 3. Using the linear equation generated from the standard curve, net fluorescence can be converted into amounts of nicotinamide. More nicotinamide produced during the reaction represents higher enzyme activity.

3.4  Mitochondrial Assays for SIRT1 Activators in Cells 3.4.1  Method Introduction

While for many years, isolated mitochondria were the preferred choice to investigate the role of the organelle in bioenergetic tissues, several techniques have emerged to analyze mitochondria within the cell. As with any technique, these methods have advantages and disadvantages. As the cellular environment of the mitochondria are maintained, in situ methods may be more physiologically relevant. However, many substrates and reagents that work on isolated mitochondria cannot be used within intact cells due the restricted permeability of the plasma membrane. While it is generally difficult to culture primary cells from adult animals for in situ comparisons, isolation of mitochondria can be used to compare animals of different ages, genotypes, or treatments.

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Table 2 Mitochondrial selective dyes

FACS channel

Reference

552/575

PE

Invitrogen T-668

Dye

Conc.

TMRM

2–100 nM Mitochondrial membrane potential

DHE

10 μM

Reactive oxygen species 490/590

PE

Sigma D7008

10 nM

Mitochondrial mass

495/538

GFP

Invitrogen A1372

10 nM

Mitochondrial mass

644/665

APC

Invitrogen M22426

NAO MitoTracker Deep Red FM ®

Measurement

Wavelength excitation/ emission

Finally, while we frequently analyze mitochondrial function to study cellular bioenergetics, it is important to remember that although the primary function of mitochondria is to fuel ATP to the cell, additional processes are related to mitochondria, such as removal of ROS, ion transport, calcium buffering, and metabolism of fatty acids, amino acids, and nucleotides. Mitochondrial dysfunction can be observed when a failure in any one of these processes is present. Conclusions must take into account that altered bioenergetic behavior may be a cause or an effect of cellular dysfunction. In this section of the chapter, we describe methodology for the investigation of in situ mitochondrial bioenergetics in response to SIRT1 activators using fluorescent dyes (Table 2). 3.4.2  Performing the Assay

1. Plate MEFs and/or C2C12 cells at 60–70% cell confluence (see Note 14). 2. On the following day treat cell with STACs (see Note 15). 3. Remove the media and wash the cell with PBS. 4. Incubate cells with new fresh media supplemented with either 10 nM TMRM for 45–60 min, 10 μM DHE for 30–60 min, 10 nM MitoTracker® Deep Red FM for 30–60 min, or 10 nM NAO for 30–60 min at 37 °C in a 5% CO2 incubator (see Note 16). NAO and MitoTracker® Deep Red FM may be incubated and analyzed together, as their spectra do not interfere with one another. 5. Remove the media and wash the cell with PBS. 6. Add trypsin to the cells and wait for them to detach. 7. Collect media and cells in a 15 mL conical tube. 8. Pellet cells by centrifugation at 300 × g for 3 min.

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9. Remove the supernatant and add 300 μL of PBS without the probe and transfer all cells to a FACS tube. 10. Keep cells on ice. 11. Read in the fluorescence-activated cell sorting (FACS) machine (10,000 or more events) using appropriate channel for the fluorescence intensity of the dye. Analyze by taking median or mean fluorescence of a sample.

4  Notes 1. To reduce background fluorescence, shorter peptides or peptides with fewer aromatic groups are preferred. 2. In situ experiments on any type of cells loaded with membrane potential dyes require a careful interpretation [25, 26]. Typically, as we select the best dye to use in our experiments, it is necessary to consider if the dye can be used in either quenching or non-quenching modes [26, 27]. Quenching mode approaches are a sensitive way to monitor real-time effects on mitochondrial membrane potential (∆ψm) in response to acute application of experimental treatment strategies (i.e., pharmacologic or toxic treatments) [25, 26, 28]. They require higher dye concentrations (50–100 nM to several micromolar), so that the dye accumulates within mitochondrial matrix in a sufficient concentration to form aggregates, thus quenching some fluorescent emissions of the aggregated dye (a phenomenon called autoquenching). Consequently, quenching mode is a nonlinear event [27], which does not detect preexisting differences in ∆ψm between two different populations (i.e., two different cell lines). On the other hand, in non-quenching mode, a lower dye concentration is used (2–30 nM), thus circumventing dye aggregation and quenching in the mitochondrial matrix. Experiments involving dye loading after experimental treatments affecting ∆ψm (i.e., STACs treatment before the dye loading to compare preexisting relative mitochondrial polarization) should typically utilize dyes in non-­quenching mode [25, 26, 28], since at very low concentrations, the fluorescent signal shows a linear relationship with the concentration of the dye. Thus, the concentration of the dye reflects the Nernstian distribution of the dye between compartments in response to local changes in potential [27]. The use of more permeant dyes, like TMRM, is therefore advantageous for this experiment. Importantly, empirical determination of the right concentration of TMRM for non-quenching experiments should be the first step in such a study. 3. The mitochondrial uptake of NAO has been shown to be independent of ∆ψm, unlike many other mitochondrial dyes. At a

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high concentration shows toxicity and can bind to cardiolipin in all mitochondria, regardless of their energetic state. 4. EDTA must not be used, as it can interfere with the Ni beads. 5. The assay undergoes two incubation steps. The first at 37 °C and the second, which uses the developer, at room temperature. It is best to keep the developer and nicotinamide for the second incubation frozen until just prior to use. Do not refreeze the assay buffer between the incubations. 6. The concentration of NAD+ can be changed to increase or decrease the amount of signal produced by SIRT1 activity. This allows for better optimization for signal vs. reagent use. 7. Using a white plate can give ~5× higher signal than that of an transparent plate, which when read in a top-reading fluorometer can greatly increase the sensitivity of the assay. 8. Having a negative control of no SIRT1 added is recommended to ensure there is no increased background and/or interactions with the compounds of interest. 9. Taking a measurement of the plate prior to adding developer can be useful as another background measurement. 10. Due to the fluorescence, this assay has a good tolerance for extracts that may discolor the wells. However, proper background readings should still be taken to ensure accuracy. 11. For enzymes that have lower activity than SIRT1, the protein amount added should be increased (e.g., SIRT6), and longer incubations performed. 12. NAD+ could exhibit fluorescence at high concentrations (>200  μM). If high NAD+ concentrations are required, the background reaction formulation can be altered. Instead of not adding NAD+, background reactions can include NAD+ but exclude the enzyme or use the corresponding non-­ acetylated peptide. 13. DMSO may inhibit the enzyme activity and lower the signal. Therefore, adding equal amount of DMSO in the same concentration to control wells is important. Typically,  13 (200 mM NaOH, 1 mM EDTA) for running the gel. 13. Blocking solution (10% goat serum, 1 M glycine in 1× PBS, 0.2% TritonX-100 and 0.005% sodium azide). 2.4  Oligonucleotides

A 43-mer oligonucleotide containing an AP-site analog tetrahydrofuran (THF) at nucleotide position 31. Label this oligonucleotide either with [γ-32P]ATP using T4 polynucleotide kinase as described previously [15] or with Cy5 (fluorescent 678/694 nm) at 5′ end. Mix equimolar amounts of 5′-end-labeled oligonucleotide containing THF with an unlabeled complementary strand in annealing buffer (10 mM Tris–HCl (pH 8), 50 mM NaCl, 1 mM EDTA) at 95 °C for 5 min then slowly cooling it to room temperature to form duplex oligomer (see Note 2). The following oligonucleotide sequence was used, 5′-GATCTGATTCCCCATCTCCTCAGTTTCACT(THF)CTG CACCGCATG-­3′. ­5′-CTAGACTAAGGGGTAGAGGAGTCAAAGTGAGGACGT GGCGTAC-­3′.

3  Methods 3.1  In Vitro Biochemical Assays to Assess the Effect of Acetylation on APE1 Enzyme

Acetylation of a BER protein can affect its activity or its affinity for the substrate DNA or the product. Thus, acetylation can positively or negatively affect the enzymatic turnover in vitro. Acetylation could act via multiple mechanisms, including effects on DNA binding and protein-protein interactions to regulate DNA damage repair. In this section, we will discuss three major in vitro assays to assess the impact of acetylation of APE1 on its functions: in vitro AP-endonulcease assay, AP-site-binding assay, and complete repair assay.

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3.1.1  In Vitro Acetylation of Recombinant APE1 Protein

1. Incubate purified recombinant APE1 protein 5–8 nM with immunopurified p300 HAT domain (100 nM) together with 2 mM acetyl-CoA in 30 μL of HAT buffer at 30 °C for 1 h. As a control reaction, incubate recombinant APE1 protein without acetyl-CoA. These proteins can be used directly for assays or can be subjected to further purification by dialysis or column to remove HAT and Acetyl-CoA (see Note 3).

3.1.2  Effect of Acetylation of APE1 on AP-Site Substrate Binding by EMSA

1. Before casting the native polyacrylamide gel, clean 1.0 mm glass plates. 2. After the plates are thoroughly cleaned, cast a 7.5% native polyacrylamide gel. A 15 mL solution is enough to cast two 1 mm gels, which can be made by mixing: 10.33 mL deionized water, 3.75 mL 30% Acrylamide/Bis solution, 0.75 mL 10× TBE, 150 μL 10% APS, and 10 μL TEMED. 3. Mix the solution thoroughly to create a homogenous mixture and cast the gel. Insert a 10-well gel comb immediately without introducing air bubbles and allow it to polymerize for 30 min–1 h. 4. Prepare 0.5× TBE buffer for electrophoresis and keep it at 4 °C. 5. Pre-run the gel with cold 0.5× TBE at 80–120 V (approximately 30 min), in the meantime prepare the samples. 6. Prepare each binding reaction mix (20 μL) in an Eppendorf tube as follows: labeled DNA (25 nM), 1–2 μg Poly dI-dC, appropriate concentrations (35 and 350 pmol) of in vitro acetylated or unmodified APE1, and make up the volume by adding 1X EMSA binding buffer. If there are multiple samples, please add the protein in the last step of the binding reaction. After adding all components, centrifuge briefly and incubate at room temperature for 10–20 min. 7. After incubation, place the tubes on ice and add 4 μL of 6× loading dye. 8. Load the samples and run the gel for approximately 2 h at 80–120 V until the dye front reaches the bottom of the gel. 9. Scan the gel using the LI-COR imaging system at an excitation wavelength around 649 nm (Fig. 2) (see Note 4).

3.1.3  Effect of Acetylation of APE1 Enzyme on its Catalytic Activity

Effect of acetylation on the enzymatic activity of a protein can be assessed by using in vitro acetylated recombinant protein and unmodified protein and lesion-containing labeled DNA substrate in vitro [15]. As an example, we will describe AP-endonuclease activity assay of APE1. 1. Prepare a denaturing 20% polyacrylamide gel containing 8 M urea. A 10 mL solution can be made by mixing: 4.8 g of 8 M urea, 5 mL of 40% Acrylamide/Bis solution, 1 mL 10× TBE,

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+

APE1

-

AcAPE1

-

+

-

+

+

+

-

-

-

DNA-protein complex Free probe

Fig. 2 5′Cy5 labeled 43-mer duplex oligonucleotide containing AP site analog THF (25 nM) was incubated with unmodified APE1 (35 or 350 pmol) or in vitro-­ acetylated APE1 (AcAPE1). DNA-protein complex was resolved in 7.5% native gel by electrophoretic mobility shift assay (EMSA)

33  μL 30% APS, 4 μL TEMED. Make up the remaining volume with deionized water (see Note 5). 2. Cast the gel in 1.0 mm glass plate (pre-cleaned with 75% ethanol). Allow the gel to solidify for 30 min–1 h. 3. Place the pre-casted gel in a tank containing 1× TBE buffer and pull out the comb carefully. Clean the wells by flushing with buffer. 4. Pre-run the gel at 80 V for 30 min (see Note 6). 5. The assay can be done either in a time- or in a dose-dependent manner to examine the reaction kinetics. 6. Prepare each reaction mixture (10 μL) in an Eppendorf tube as follows: 2.5 pmol of the 5′-32P end-labeled duplex oligonucleotide containing THF and 200−1000 pg of in vitro acetylated or unmodified APE1 in reaction buffer. After adding all components, incubate it at 37 °C for 5 min or for various time points (2, 5, and 10 min) during which the reaction rate is linear. 7. As soon as the chosen time periods are over, stop the reaction by adding an equal volume of the loading buffer (95% formamide, 10 mM EDTA, 0.1% bromophenol blue and 0.1% xylene cyanol) to each of the reaction mixtures and incubate at 95 °C for 5 min. 8. Clean the wells of the gel again by flushing with gel running buffer to ensure there is no urea in the wells before loading the sample (see Note 7). 9. Load the samples and run the gel slowly at 65 V until the dye front reaches two-third of the length of the gel. 10. Scan the gel using the standard phosphorimager system or wrap it with Saran wrap and expose to film for autoradiography.

Assays to Examine the Effect of Acetylation on BER Enzymes 3.1.4  Effect of Acetylation of APE1 on a Reconstituted Complete Repair Assay

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1. Prepare denaturing 12.5% polyacrylamide gel containing 8 M urea. For a 10 mL solution, mix 4.8 g of 8 M urea, 3.12 mL of 40% Acrylamide/Bis solution, 1 mL 10× TBE, 2 mL of deionized water, 33 μL 30% APS, 4 μL TEMED. 2. Cast the gel in 1.0 mm glass plate (pre-cleaned with 75% ethanol). Allow the gel to solidify for 30 min–1 h. 3. Place the pre-casted gel in a tank containing 1× TBE buffer and pull out the comb. Clean the wells by flushing with buffer. 4. Pre-run the gel at 80 V for 30 min (see Note 6). 5. Prepare reaction mixtures as follows (50 μL): 20 μM each of dATP, dGTP, and dTTP, 2 μM dCTP including 2 μCi [α-32P] dCTP (3000 Ci mmol−1), 1 ng of APE1 or AcAPE1, 1 ng DNA polymerase β, and 5 ng DNA ligase III in reaction buffer. The components were mixed at 0 °C and reactions can be initiated by addition of 2 pmol duplex AP site (THF) containing oligonucleotide. Now, transfer the reaction mixture to 37 °C for 30 min. The protein concentrations should be optimized for maximum repair synthesis [21] (see Note 8). 6. Stop the reaction by adding an equal volume of the loading buffer (95% formamide, 10 mM EDTA, 0.1% bromophenol blue, and 0.1% xylene cyanol) to each of the reaction mixtures and incubate at 95 °C for 5 min. 7. Clean the wells of the gel again by flushing with gel running buffer to ensure there is no urea in the wells before loading the samples. 8. Load the samples and run the gel slowly at 65 V until the dye front reaches two-third of the length of the gel. Scan the gel using the standard phosphorimager or wrap it with Saran wrap and expose to film for autoradiography.

3.2  Cellular Assays to Assess the Effects of Acetylation on BER Enzymes

Spatiotemporal acetylation of BER enzymes may modulate the overall efficiency of the DNA damage repair processes in the context of chromatin in cells. Moreover, in the event of induction of damage by exogenous sources, an immediate increase in BER activity is imperative which could be achieved by acetylating BER enzymes, without increasing the amount of the protein. However, if acetylation is slow or nonexistent, overall BER pathway efficacy can be greatly reduced, leading to accumulation of BER intermediates, single strand breaks in the DNA, impacting genomic stability and cellular survival. Cells expressing the non-acetylable (Lysine to Arginine) mutant proteins are often used to monitor DNA damage and repair capacity by comet assay and cell survival by colony formation assay. Moreover, generation of antibody against an acetylated peptide of a protein of interest is useful in understanding the subcellular localization of the acetylated proteins under different cellular context. In this section,

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we will discuss comet assay, immunofluorescence, and cell survival assay, which are frequently used to assess the effect of acetylation on BER proteins in mammalian cells. 3.2.1  Examine the Effect of Acetylation of BER Enzymes on DNA Damage Repair In Vivo by Comet Assay

Comet assay is a powerful, single-cell electrophoresis technique that can measure the overall presence of DNA damage in the nuclear genome in vivo [22]. Neutral comet assay measures DNA double-strand breaks (DSBs) while the alkaline comet assay measures both DSBs, single-strand breaks and alkali-labile sites such as AP lesions. Trevigen comet assay protocol is beneficial to standardize abovementioned assay system in the lab. 1. Plate cells that are expressing wild-type or acetylation-­defective mutant enzyme in a six-well culture plate. Treat these cells with genotoxic agents of interest (such as gamma radiation, alkylating agent MMS, or oxidizing agent H2O2) for specific time periods and release cells in fresh media for 2–24 h to allow repair. 2. Once the cells are ready to be harvested, place the low melting agarose (LMA) in boiling water for approximately 5 min or until the solution is completely liquefied. Open the lid of the LMA container slightly to relieve pressure during this process. Once liquid, aliquot 100 μL of LMA to the desired number of microcentrifuge tubes and place in a 37 °C incubator to cool. 3. To harvest the cells, add 0.25% trypsin to each well. Remove trypsin from the cells and place in a 37 °C incubator for approximately 2 min or until cells detach. 4. Add 1 mL media containing 10% FBS. Pipette media over the plate until all cells are resuspended. Collect cell solution into microcentrifuge tubes. 5. Pellet cells by centrifugation at 100 × g for 5 min and wash twice with 1× PBS. Count the cells and resuspend in 1× PBS to generate a suspension of approximately 1 × 105 cells/mL. 6. Add 10 μL of the collected cell suspensions to the respective microcentrifuge tube containing 37 °C LMA. Using a p200 with the tip cut back, create a homogenous mixture of LMA and cells by pipetting up and down. Quickly add 50 μL of the solution to the comet slide, carefully avoiding solidification, and spread over the marked circular area on comet slide (see Note 9). 7. Place the comet slides in the dark at 4 °C until the agarose is completely solidified (approximately 10–20 min). After a while, submerge the samples in Trevigen lysis solution in the dark at 4 °C for overnight. 8. Prepare alkaline unwinding solution by dissolving 0.6 g NaOH pellets in 250 μL of 200 mM EDTA and 49.75 mL deionized water.

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9. Drain out excess lysis buffer, and submerge the slide in alkaline unwinding solution for 20 min at room temperature. Meanwhile, prepare 1-L alkaline electrophoresis solution by dissolving 8 g of NaOH pellets in 2 mL of 500 mM EDTA (pH 8) and deionized water (see Note 10). 10. Fill electrophoresis unit with alkaline electrophoresis solution just enough for the slides to be submerged. Once the alkaline unwinding step is complete, place the slides equidistant from the electrodes with the label adjacent to the cathode. Perform electrophoresis at 21 V for 30 min (see Note 11). 11. After electrophoresis, drain off excess solution from the slides and submerge in deionized water followed by 70% ethanol for 5 min. Place slides at 37 °C for 10–15 min before staining with SYBR Green. 12. Dilute SYBR Green 1:1000 and add 100 μL to each sample for 30 min. After staining, submerge the comet slides in deionized water to rinse off excess SYBR Green. Place slides in the dark until completely dry. 13. Visualize samples under the microscope at 10× magnification. Comet tails can be scored quantitatively by open comet software (Fig. 3). 3.2.2  Effect of Acetylation of BER Enzymes on Cell Survival; Clonogenic Survival Assay

Ability of tumor cells to form colonies upon genotoxic insult can be measured by clonogenic survival assay [15]. 1. Plate equal numbers (approximately 500) of cells on 60 mm plates in triplicate, containing appropriate media (see Note 12). 2. After 16–20 h (log phase culture), treat cells without or with increasing doses of genotoxic agents. 3. Wash cells with 1× PBS for two times and add fresh growth media. 4. Allow cells to grow for 2 weeks until visible colonies appear (see Note 13). 5. Pour the medium from the Petri dish into a container for disposal and carefully add about 5 mL of PBS to wash off the remaining medium. 6. Pour off the PBS and add about 5 mL of 100% methanol for fixation and leave for 5 min. Repeat the step one more time. 7. Add 5 mL of crystal violet to each Petri dish and leave for 5 min. Pour off the stain and rinse the dish under running tap water to remove excess dye. Invert dishes and leave to dry. 8. Count colonies for each treatment. 9. Calculate the mean colony count for each of the treatments. Divide the number of colonies in the drug-treated dishes by the number of colonies in the control dishes and express as a

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Fig. 3 Human medulloblastoma cells ONS-76 cells were exposed to gamma-radiation (2 Gy) and subjected to alkaline comet assay. (a) Representative images from the comet assay (10×). (b) The length of the comet tails was quantitated by open comet software. ****p 13) of the alkaline unwinding solution and electrophoresis solution is important for proper separation in the gel. 2. During the duplex oligonucleotide annealing process, gradually cool it from 95 °C to room temperature. Do not use PCR machine as temperature drop is rapid. 3. After preparation of acetylated protein, it can be separated from unmodified protein by FPLC as described previously [16, 24] or can be used directly for in vitro assays as described. 4. When working with Cy5 labeled oligonucleotides, the gel should be scanned immediately after completion of running on the same day, to obtain crisp and distinct bands. Leaving the gel without scanning for long time periods may lead to increased diffusion and decreased fluorescence of the bands. 5. While preparing the 12.5% polyacrylamide gel containing 8 M urea, care should be taken to dissolve the 8 M urea properly by heating mildly before addition of APS and TEMED. It should be kept in mind that the APS and TEMED are added once the gel mixture cools down. 6. Pre-running the gel is important to warm up and equilibrate the gel to maintain a uniform denaturing condition. 7. Before as well as after pre-running the gel clean the wells by flushing with the running buffer to make sure that there is no urea left in the wells, otherwise, urea can interfere with the sample loading. 8. In complete repair assay, the reaction mixture will have cold oligos and specific 2 μCi of [α-32P]dNTP chosen according to lesion site sequence. 9. Place comet slides in 37 °C incubator before adding the cells to prevent non-uniform solidification of agarose. 10. Ensure Lysis solution was chilled before use.

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11. Electrophoresis should be performed at 4 °C. 12. In cell survival assay, do not label the lids of the culture plate because these are removed when the colonies are fixed. Make sure that the marker pen used is resistant to methanol. 13. The incubation time for colony formation will vary depending on the doubling time of the cell line used but is usually between 8–12 days. This allows for about 10 doubling times. It is advisable to check the dishes after about 8 days, and colonies should be clearly visible to the naked eye. 14. The second fixation step with 100% methanol for 10 min in immunostaining is purely optional. It depends on the antibody we use in the subsequent steps of the protocol. 15. In immunostaining, the incubation period for the primary antibody and secondary antibody is very crucial. Over-­ incubation of secondary antibody can give rise to high background staining which might interfere with the results of the experiment.

Acknowledgment This research was supported by NIH/NCI R01CA148941 and Nebraska state DHS LB506 grants. References 1. Choudhary C, Kumar C, Gnad F, Nielsen ML, Rehman M, Walther TC, Olsen JV, Mann M (2009) Lysine acetylation targets protein complexes and co-regulates major cellular functions. Science 325:834–840. https://doi. org/10.1126/science.1175371 2. Polevoda B, Sherman F (2002) The diversity of acetylated proteins. Genome Biol 3:reviews0006 3. Piekna-Przybylska D, Bambara RA, Balakrishnan L (2016) Acetylation regulates DNA repair mechanisms in human cells. Cell Cycle 15:1506–1517. https://doi.org/10.10 80/15384101.2016.1176815 4. Spange S, Wagner T, Heinzel T, Kramer OH (2009) Acetylation of non-histone proteins modulates cellular signalling at multiple levels. Int J Biochem Cell Biol 41:185–198. https:// doi.org/10.1016/j.biocel.2008.08.027 5. Busso CS, Lake MW, Izumi T (2010) Posttranslational modification of mammalian AP endonuclease (APE1). Cell Mol Life Sci 67:3609–3620. https://doi.org/10.1007/ s00018-010-0487-3

6. Carter RJ, Parsons JL (2016) Base excision repair, a pathway regulated by posttranslational modifications. Mol Cell Biol 36:1426–1437. https://doi.org/10.1128/MCB.00030-16 7. David SS, O’Shea VL, Kundu S (2007) Base-­ excision repair of oxidative DNA damage. Nature 447:941–950. pii: nature05978 8. Dianov GL, Hubscher U (2013) M ammalian base excision repair: the forgotten archangel. Nucleic Acids Res 41:3483–3490. https://doi.org/10.1093/nar/gkt076 9. Parikh SS, Mol CD, Hosfield DJ, Tainer JA (1999) Envisioning the molecular choreography of DNA base excision repair. Curr Opin Struct Biol 9:37–47. pii: S0959440X(99)80006-2 10. Mitra S, Izumi T, Boldogh I, Bhakat KK, Hill JW, Hazra TK (2002) Choreography of oxidative damage repair in mammalian genomes. Free Radic Biol Med 33:15–28. pii: S0891584902008195 11. Bhakat KK, Hazra TK, Mitra S (2004) Acetylation of the human DNA glycosylase NEIL2 and inhibition of its activity. Nucleic

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Acids Res 32:3033–3039. https://doi. org/10.1093/nar/gkh632 12. Sengupta S, Yang C, Hegde ML, Hegde PM, Mitra J, Pandey A, Dutta A, Datarwala AT, Bhakat KK, Mitra S (2018) Acetylation of oxidized base repair-initiating NEIL1 DNA glycosylase required for chromatin-bound repair complex formation in the human genome increases cellular resistance to oxidative stress. DNA Repair (Amst) 66–67:1–10. pii: S15687864(18)30072-7 13. Tini M, Benecke A, Um SJ, Torchia J, Evans RM, Chambon P (2002) Association of CBP/p300 acetylase and thymine DNA glycosylase links DNA repair and transcription. Mol Cell 9:265–277. pii: S1097-2765(02) 00453-7 14. Bhakat KK, Mokkapati SK, Boldogh I, Hazra TK, Mitra S (2006) Acetylation of human 8-oxoguanine-DNA glycosylase by p300 and its role in 8-oxoguanine repair in vivo. Mol Cell Biol 26:1654–1665. pii: 26/5/1654 15. Roychoudhury S, Nath S, Song H, Hegde ML, Bellot LJ, Mantha AK, Sengupta S, Ray S, Natarajan A, Bhakat KK (2017) Human apurinic/apyrimidinic endonuclease (APE1) is acetylated at DNA damage sites in chromatin, and acetylation modulates its DNA repair activity. Mol Cell Biol 37:e00401-16. Print 15 Mar 2017. pii: e00401-16 16. Bhakat KK, Izumi T, Yang SH, Hazra TK, Mitra S (2003) Role of acetylated human AP-endonuclease (APE1/Ref-1) in regulation of the parathyroid hormone gene. EMBO J 22:6299–6309. https://doi.org/10.1093/ emboj/cdg595 17. Lirussi L, Antoniali G, Vascotto C, D’Ambrosio C, Poletto M, Romanello M, Marasco D, Leone M, Quadrifoglio F, Bhakat KK, Scaloni A, Tell G (2012) Nucleolar accumulation of APE1 depends on charged lysine residues that undergo acetylation upon genotoxic stress and modulate its BER activity in

cells. Mol Biol Cell 23:4079–4096. https:// doi.org/10.1091/mbc.E12-04-0299 18. Hasan S, El-Andaloussi N, Hardeland U, Hassa PO, Burki C, Imhof R, Schar P, Hottiger MO (2002) Acetylation regulates the DNA end-trimming activity of DNA polymerase beta. Mol Cell 10:1213–1222. pii: S10972765(02)00745-1 19. Sengupta S, Mantha AK, Song H, Roychoudhury S, Nath S, Ray S, Bhakat KK (2016) Elevated level of acetylation of APE1 in tumor cells modulates DNA damage repair. Oncotarget 7:75197–75209. https://doi. org/10.18632/oncotarget.12113 20. Yamamori T, DeRicco J, Naqvi A, Hoffman TA, Mattagajasingh I, Kasuno K, Jung SB, Kim CS, Irani K (2010) SIRT1 deacetylates APE1 and regulates cellular base excision repair. Nucleic Acids Res 38:832–845. https://doi. org/10.1093/nar/gkp1039 21. Das A, Wiederhold L, Leppard JB, Kedar P, Prasad R, Wang H, Boldogh I, Karimi-Busheri F, Weinfeld M, Tomkinson AE, Wilson SH, Mitra S, Hazra TK (2006) NEIL2-initiated, APE-independent repair of oxidized bases in DNA: evidence for a repair complex in human cells. DNA Repair (Amst) 5:1439–1448. pii: S1568-7864(06)00216-3 22. Olive PL, Banath JP (2006) The comet assay: a method to measure DNA damage in individual cells. Nat Protoc 1:23–29. pii: nprot.2006.5 23. Chattopadhyay R, Das S, Maiti AK, Boldogh I, Xie J, Hazra TK, Kohno K, Mitra S, Bhakat KK (2008) Regulatory role of human AP-endonuclease (APE1/Ref-1) in YB-1-­ mediated activation of the multidrug resistance gene MDR1. Mol Cell Biol 28:7066–7080. https://doi.org/10.1128/MCB.00244-08 24. Bhakat KK, Yang SH, Mitra S (2003) Acetylation of human AP-endonuclease 1, a critical enzyme in DNA repair and transcription regulation. Methods Enzymol 371:292–300. https://doi. org/10.1016/S0076-6879(03)71022-2

Chapter 12 Analysis of DNA Processing Enzyme FEN1 and Its Regulation by Protein Lysine Acetylation Onyekachi E. Ononye, Catherine W. Njeri, and Lata Balakrishnan Abstract Cellular proteins are modified by lysine acetylation wherein an acetyltransferase transfers an acetyl group from acetyl co enzyme A onto the e-amino group of lysine residues. This modification is extremely dynamic and can be reversed by a deacetylase that removes the acetyl group. Addition of acetyl group to the lysine residue neutralizes its positive charge, thereby functioning as a molecular switch in regulating the enzymatic functions of the protein, its stability, and it cellular localization. Since this modification is extremely dynamic within the cell, biochemical studies characterizing changes in protein function are imperative to understand how this modification alters protein function in a specific cellular pathway. This unit describes in detail expression and purification of a recombinant nuclease and acetyltransferase, in vitro acetylation of the recombinant protein and biochemical assays to study the changes in enzymatic activity of the in vitro acetylated nuclease. Key words Lysine acetylation, In vitro acetylation, Lysine acetyltransferase, p300, Flap endonuclease 1 (FEN1), Nuclease enzyme assays

1  Introduction Addition of chemical groups to specific side chain residues on amino acids are called posttranslational modifications (PTMs) [1]. Transfer of an acetyl group from acetyl coenzyme A to a specific amino acid is known as acetylation. Proteins can be N-terminally acetylated (Nt-ac) by modifying the α-amino group of the first amino acid residue of the protein. This is carried out by a group of enzymes known as N-terminal acetyltransferases (NATs) [2]. Proteins can also be specifically acetylated by the addition of an acetyl group to the ε-amino group of lysine (K) residues. Lysine acetylation is a dynamic, readily reversible modification which is mediated by lysine acetyl transferases (KATs) and lysine deacetylases (KDACs). Acetylation of the ε-amino group of a lysine residue neutralizes its positive charge, which in turn may have a myriad of effects on the protein, such as regulation of enzymatic activity Robert M. Brosh, Jr. (ed.), Protein Acetylation: Methods and Protocols, Methods in Molecular Biology, vol. 1983, https://doi.org/10.1007/978-1-4939-9434-2_12, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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[3–5], modulation of protein-nucleic acids and/or protein-protein interactions [6], alteration of sub-cellular localization [7–10], and/or changes in protein stability and degradation [11]. While lysine acetylation of histone proteins has been the focus of study for many decades, recent advances in proteomics have identified thousands of cellular, nonhistone proteins that are acetylated in response to cellular perturbations [12–14]. In fact, it is now widely accepted that lysine acetylation is as abundant as protein phosphorylation in regulating protein characteristics. Assessing changes in protein characteristics on account of lysine acetylation has proved to be technically challenging, since this modification is extremely dynamic and other forms of modification, such as ubiquitination and sumoylation, can also compete with acetylation for the same lysine residues [15]. Biochemical characterization of protein enzymatic function relies on the use of purified recombinant proteins for various assays. Commonly used biochemical techniques used to study protein acetylation are to create acetylation mimic (lysine to glutamine) or acetylation inhibitory (lysine to arginine) mutants. In order to create these mutant proteins, one needs to first identify sites of cellular lysine acetylation. A significant drawback of this technique is that if the lysine acetylation occurs in the active site of the protein or at a site that is known to interact with other proteins or DNA for functional activity, then modifying the lysine residue to either mimic or inhibit acetylation may in itself change the activity of the protein. Other techniques involve modifying the lysine residues that are known to be acetylated to cysteine residues and then using cysteine alkylation technology to incorporate acetyl analogs on cysteine residues that mimic an acetylated lysine residue [16]. Altering the lysine to a cysteine group can cause inherent changes in enzymatic function and it is also difficult to control naturally occurring cysteines in the protein from being modified. An extremely innovative technology has also been developed to genetically modify site specific lysine residues using nonsense suppression methodology [17, 18]. However, the limitation of this technique is that multiple lysine sites cannot be modified at the same time and the yield of purification is significantly low. Since each in vitro acetylation techniques have their pros and cons, it is best to use multiple forms of in vitro acetylated protein to characterize lysine acetylation-based enzymatic changes. The method that is outlined in this protocol uses purified recombinant acetyltransferases and acetyl coenzyme A to generate acetylated proteins. While this method is fairly simple to perform in the laboratory, there are a few caveats that need to be considered while interpreting the data. The cell contains numerous KATs and KDACs that play both specific and redundant roles within the cell. Each KAT and KDAC identifies specific signatures in the protein amino acid sequence that allow it to modify the protein.

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While performing in vitro acetylation reactions, one needs to first identify the KAT that is responsible for modifying the protein within the cell, in order to mimic cellular conditions. In vitro acetylation reactions are not fully efficient and KATs can frequently be promiscuous in transferring acetyl groups to lysine residues which may not function as its substrate in vivo. These reactions also yield a mixed population of modified recombinant proteins that contain different stoichiometries of acetylated residues. However, since the biochemical assays provide an output for the combined group of this mixed population of modified protein, we will still be able to obtain important information about changes in enzymatic activity after lysine acetylation. In the current chapter we are outlining the expression and purification of a DNA replication/repair associated nuclease, namely, flap endonuclease 1 (FEN1) and histone acetyltransferase p300. We further describe a protocol for in vitro acetylation of recombinant FEN1 and enzymatic assays to study the effect of this modification on enzyme activity. The technique described here can be adapted to in vitro modify any recombinant protein using a suitable acetyltransferase and study biochemical changes to their enzyme activity.

2  Materials 2.1  Expression and Purification of Recombinant Human Flap Endonuclease 1 (hFEN1) 2.1.1  Buffers and Reagents Transformation and Protein Expression

Protein Purification

1. Competent BL21(DE3) cells. 2. pET-FCH FEN1 plasmid construct (see Note 1). 3. Luria Broth (LB) media. 4. Kanamycin sulfate: 50 mg/mL. 5. Isopropyl β-d-1-thiogalactopyranoside (IPTG): 0.4 M IPTG dissolved in distilled water. 6. LB/Kan plate: LB Agar supplemented with 50 μg/mL kanamycin sulfate. 7. LB/Kan media: LB supplemented with 50 μg/mL kanamycin sulfate. 1. hFEN1 Lysis Buffer: 50 mM Tris–HCl [pH 8.0], 500 mM NaCl, 1 mM beta mercaptoethanol (BME), 1 mM phenylmethylsulfonyl fluoride (PMSF), 1 mM EDTA, 1 tablet complete protease inhibitor, 10 mM imidazole, 10% glycerol. 2. hFEN1 Storage Buffer: 50 mM KPi [pH 7.0], 100 mM NaCl, 10% glycerol. 3. hFEN1 His Wash Buffer: 50 mM Tris–HCl [pH 8.0], 500 mM NaCl, 1 mM BME, 1 mM PMSF, 10 mM imidazole, 10% glycerol.

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4. hFEN1 His Elution Buffer: 50 mM Tris–HCl [pH 8.0], 500 mM NaCl, 1 mM BME, 1 mM PMSF, 1 M imidazole, 10% glycerol. 5. Imidazole: 1 M dissolved in distilled water. 6. Distilled water. 7. Ethanol: Diluted to 20% in distilled water. 8. Ni-NTA agarose resin: Pack approximately 5 mL of resin. 9. Size exclusion column (SEC) 500–70 kDa. 2.1.2  Equipment

1. 200 or 250 mL conical flasks. 2. 2800 mL flasks. 3. High speed centrifuge with compatible rotors that can hold up to 4 °C. 4. NGC™ medium pressure gas chromatography system.

2.2  Expression and Purification of Histone Acetyl Transferase p300 2.2.1  Buffers and Reagents

1. p300-HAT-pUC57-Kan construct (see Note 2). 2. pET28b plasmid. 3. BL21 (DE3) cells. 4. LB/Kan plates. 5. LB/Kan media.

Transformation

6. Ni-NTA Agarose resin.

Protein Purification

1. Autoinduction media: 90 mM KPi [pH 7.0], 1 g/L glucose, 2 g/L lactose, 0.5% glycerol, 12 g peptone, 24 g yeast extract, and distilled water to a final volume of 1 L supplemented with 50 μg/mL kanamycin (see Note 3). 2. P300 Lysis Buffer: 25 mM HEPES [pH 7.5], 300 mM NaCl, 10% glycerol, and 10 mM imidazole. 3. P300 Elution Buffer: 25 mM HEPES [pH 7.5], 300 mM NaCl, 10% glycerol, and 300 mM imidazole. 4. P300 Storage Buffer: 25 mM HEPES [pH 7.5], 300 mM NaCl, and 10% glycerol.

2.2.2  Equipment

1. Dialysis tubing. 2. NGC™ medium pressure gas chromatography system or equivalent.

2.3  In Vitro Acetylation 2.3.1  Buffers and Reagents

1. Acetyl-CoA: 14C labeled and unlabeled sodium salt. 2. 1× Histone Acetyltransferase (HAT) Buffer: 50 mM Tris–HCl [pH 8.0], 1 mM dithiothreitol (DTT), 10 mM sodium butyrate (NaBt), 10 mM NaCl, 1 mM PMSF, and 10% (v/v) glycerol.

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3. Purified recombinant histone acetyltransferase: p300 (or appropriate KAT). 4. Purified recombinant human FEN1 (or any protein to be modified). 2.3.2  Equipment

1. 37 °C Heat block.

2.4  Detection of In Vitro Acetylation

1. 4–15% gradient SDS PAGE gel.

2.4.1  Buffers and Reagents

2. Pre-stained molecular weight standard. 3. MagicMark™ XP western protein standard. 4. 1× Laemmli sample buffer. 5. Tris/Glycine/SDS electrophoresis running buffer: 25 mM Tris, 192 mM glycine, 0.1% SDS, [pH 8.3]. 6. Tris/Glycine transfer buffer: 25 mM Tris, 192 mM glycine, [pH 8.3], and 20% methanol. 7. TBS-T: 1× Tris buffered saline (20 mM Tris, 500 mM NaCl, [pH 7.5]), 0.1% Tween 20. 8. Blocking buffer: 5% Milk dissolved in TBS-T. 9. Polyvinylidene difluoride (PVDF) membrane activated in methanol. 10. Primary antibody: Anti-acetyl lysine. 11. Horse radish peroxidase (HRP) conjugated secondary antibody. 12. ECL reagent. 13. 3 mm Chromatography paper (cut into squares) (see Note 4).

2.4.2  Equipment

1. ImageQuant LAS 4000 mini biomolecular imager or equivalent. 2. Vacuum pump. 3. Gel drier. 4. Autoradiography cassette to expose film. 5. Autoradiography film. 6. OPTIMAX™ X-ray processor.

2.5  Enzyme Assays

1. Nicked flap substrate (see Note 5).

2.5.1  Buffers and Reagents

2. React 2 Buffer: 1 M Tris–HCl [pH 8.0], 1 M MgCl2, 1 M NaCl.

Labeling of Oligonucleotides

3. Nuclease-free water. 4. α-32P dCTP. 5. DNA Polymerase I, large Klenow fragment. 6. 10× TBE: 1 M Tris-Borate, 0.02 M EDTA, dissolved in 1 L of distilled water.

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7. 1× TBE (running buffer): Dilute 100 mL of 10× TBE in 900 mL of distilled water. 8. 10% ammonium persulfate (APS). 9. Tetramethylethylenediamine (TEMED). 10. 12% sequencing gel: 45 mL acrylamide:bisacrylamide (40% w/v), 63 g urea, 15 mL 10× TBE, 300 μL 10% APS, 30 μL TEMED, and 37.5 mL distilled water. Spacers used to make sequencing gel are 0.8 mm thick. 11. Ethanol: 95% and 70%. 12. Duplex annealing buffer: 100 mM potassium acetate, 30 mM HEPES [pH 7.5]. 13. Micro bio-spin P-30 column. 14. Whatman filter paper. 15. Scintillation vials. 16. Ecoscint scintillation fluid. 17. Spin-X filter column 0.22 μm. Nuclease and Binding Assays

1. 10× and 1× TBE (running buffer): 0.1 M Tris-Borate, 0.002 M EDTA, dissolved in 1 L of distilled water. 2. 15% Denaturing gel: 22.5 mL acrylamide:bisacrylamide (40% w/v), 33.8 g Urea, 8 mL 10× TBE, 400 μL 10% APS, 40 μL TEMED, and 29.2 mL distilled water. Spacers used to make denaturing gel are 0.35 mm thick. 3. Nuclease reaction buffer: 50 mM Tris–HCl [pH 8.0], 2 mM DTT, 0.25 mg/mL bovine serum albumin (BSA), 30 mM NaCl, 4 mM MgCl2, 2 mM ATP, and 5% glycerol. 4. 6% Native gel: 15 mL acrylamide:bisacrylamide (40% w/v), 10 mL 10× TBE, 550 μL 10% APS, 55 μL TEMED, and 74.5 mL distilled water. Spacers used to make native gel are 1.5 mm thick. 5. EMSA reaction buffer: 50 mM Tris–HCl [pH 8.0], 2 mM DTT, 0.25 mg/mL BSA, 30 mM NaCl, 20 μM EDTA, and 5% glycerol. 6. Termination Dye: 90% formamide (v/v), 10 mM EDTA, 0.1% xylene cyanol, and 0.1% bromophenol blue.

2.5.2  Equipment

1. OPTIMAX™ X-ray processor. 2. Scintillation counter. 3. Sequencing S2 gel electrophoresis apparatus. 4. Polyacrylamide gel electrophoresis system for 17 cm × 15 cm vertical glass plates. 5. Power supply. 6. Typhoon™ FLA 9500 laser scanner.

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3  Method 3.1  Expression and Purification of Recombinant Human Flap Endonuclease 1 (hFEN1) 3.1.1  Large-Scale Expression of hFEN1

1. Thaw 50 μL of competent BL21 (DE3) on ice. 2. Add 2 μL of pET-FCH FEN1 plasmid and incubate on ice for 30 min. 3. Heat shock the transformation reaction at 42 °C for 45 s. Following this, incubate the reaction on ice for 2 min. 4. Add 500 μL of LB medium to the transformation reaction and incubate at 37 °C for 1 h. 5. Plate 100 μL of the transformation reaction on an LB/Kan plate and incubate overnight at 37 °C. 6. Pick a single colony from the overnight plate and inoculate into 100 mL LB/Kan media. 7. Incubate media at 37 °C overnight with shaking. 8. On the same day, autoclave four 1 L LB media in 2800 mL flasks to be used the following day for larger expression of the protein. 9. The next day, add 1 mL of 50 mg/mL Kan into each of the autoclaved 1 L LB media. 10. Inoculate 20 mL of the overnight culture into each flask and incubate the cultures at 37 °C with shaking until OD600 ~ 0.6. 11. Induce the culture with 0.4 mM IPTG and express the protein for an additional 3 h at 37 °C. 12. Harvest cells by centrifuging at 4400 × g for 20 min at 4 °C. 13. Resuspend the cell pellet (from all 4 L) in 100 mL lysis buffer. Transfer the resuspended cells into a clean 250 mL plastic beaker. 14. Sonicate the resuspended cells using the following parameters: 30 s on; 30 s off for 15 min at amplitude of 35% (see Note 6). 15. Spin down the cell lysate at 2900 × g for 30 min at 4 °C and collect the supernatant.

3.1.2  Purification Procedure

1. Pack Ni-NTA agarose resin into column and equilibrate with hFen1 His Wash Buffer. 2. Apply the supernatant to the column and collect the flow through (see Note 7). 3. Wash the column with 50 mL hFEN1 His wash buffer and collect the wash as a single fraction. 4. Elute the protein with a 70 mL gradient of 10 mM to 1 M imidazole and collect 1.5 mL fractions. 5. Electrophorese protein peak fractions on an SDS-PAGE gel to assess purity.

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p300 (HAT)

250 -150 -100 -75 --

hFEN1

250 -150 --

50 --

100 -75 --

37 --

50 -37 --

25 --

25 -20 -15 -10 --

20 -15 -10 --

1

2

4

5

6

Fig. 1 Purified recombinant hFEN1 and p300

6. Pool the fractions containing the protein and dialyze overnight with 2 L of hFEN1 storage buffer. Change dialysis buffer in the morning and dialyze for an additional 3 h (see Note 8). 7. Following dialysis, measure protein concentration with preferred method. 8. Make 500 μL aliquots of dialyzed protein since you can only inject 500 μL at a time into the SEC (see Notes 9 and 10). 9. Wash the SEC column with 2 column volumes (CV) filtered milli-Q water at a flow rate of 0.5 mL. 10. Equilibrate the column with 2 CV hFEN1 storage buffer. 11. Inject 500  μL of hFEN1 and elute with hFEN1 storage buffer. 12. Run protein peak fractions on an SDS-PAGE gel to assess purity (see Fig. 1). 13. Pool the fractions containing the protein. 14. Measure protein concentration, aliquot, and store at −80 °C. 15. Clean up columns as described in Note 11. 3.2  Expression and Purification of Histone Acetyl Transferase p300 3.2.1  Large-Scale Expression of p300

1. Digest and ligate the p300-HAT-pUC57-Kan plasmid into the EcoR1-Xho1 site of pET28b to yield p300-HAT-pET28b plasmid for protein expression. 2. Transform p300-HAT-pET28b plasmid into competent BL21 (DE3) as described in Subheading 3.1.1. 3. Plate 100 μL of transformation reaction onto LB/Kan plates and incubate overnight at 37 °C.

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4. Inoculate 100 mL LB-Kanamycin starter culture with a single colony from the successfully transformed plate and incubate overnight at 37 °C. 5. Prepare two 1 L autoinduction media (see Note 3). 6. The next day, supplement the two flasks containing 1 L autoinduction media with Kanamycin to a final concentration of 50 μg/mL. Add 20 mL of overnight culture to each flask and grow at room temperature for 48 h. 7. Harvest cells by centrifugation using the same specifications as outlined in Subheading 3.1.1. 8. Resuspend the cell pellet in 100 mL p300 lysis buffer and lyse by sonication as previously described. 9. Centrifuge the cell lysate to remove cellular debris and collect the cleared lysate in the supernatant. 3.2.2  p300 Purification Procedure

1. Prepare Ni-NTA agarose column as earlier described in Subheading 3.1.2 but equilibrate column with p300 Wash buffer. 2. Apply the cleared lysate to the column and collect flow through (see Note 7). 3. Wash the column with 10 CVs of p300 wash buffer and collect the flow through as a single fraction (see Note 12). 4. Elute the p300 HAT domain in 1 mL fractions with 100 mL of a linear gradient of p300 elution buffer. 5. Electrophorese protein peak fractions on an SDS-PAGE gel to assess purity (see Fig. 1). 6. Pool the fractions containing the p300 HAT domain together and dialyze overnight in the p300 storage buffer. 7. Measure the protein concentration of the dialyzed protein using a preferred method. Aliquot and store protein at −80 °C. 8. Measured the acetyl transferase activity (see Note 13).

3.3  In Vitro Acetylation

1. For the acetylation reaction to be successful, the order in which the components are introduced into the reaction is vital (see Note 14). 2. Add the appropriate volume of 1× HAT required to make up a final reaction volume of 20 μL to a 1.5 mL microfuge tube. Recall that the volume to be used is dependent on the amount of protein to be acetylated and the standard acetylation ratio (see Note 15). 3. Combine all the components in the right order and incubate at 37 °C for 30 min. 4. We utilize either of the two types of Acetyl-CoA (14C labeled and unlabeled) in our laboratory and compare the levels of acetylation using methods that will be described in the next section.

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3.4  Detection of In Vitro Acetylation

1. To a microfuge tube, add 5 μM of the acetylated protein and 10 μL of 1× Laemmli buffer. 2. Heat the solution at 95 °C for 5 min and spin for 30 s to recover samples lost due to condensation. 3. Load on an SDS-PAGE gel alongside the molecular weight marker and the magic marker. Electrophorese in Tris/glycine/ SDS buffer for 30 min at 180 V (see Note 16). 4. Transfer gel to PVDF membrane using the sandwich method and electrophorese at 55 V for 1 h (see Note 17). 5. Briefly wash the blot with TBS-T and block for an hour with mild shaking. 6. Discard blocker, add acetyl lysine primary antibody in 5% milk (1:1000), and incubate with shaking overnight in 4 °C. 7. The next day, wash the blot three times with TBS-T for 10 min each. 8. Add the appropriate HRP conjugated secondary antibody diluted in 5% milk and incubate for an hour with mild shaking at room temperature. 9. Wash the blot three times with TBS-T for 10 min each. 10. Add ECL reagent to the blot and develop. 11. Once the SDS-PAGE gel has been electrophoresed, remove gel from casing and place on a piece of chromatography paper. 12. Cover gel with plastic wrap and dry in a gel dryer for 1 h. 13. Once the gel is dry, tape it to the inside of the cassette and cover the gel with an autoradiography film (see Note 18). 14. Place the cassette into the −80 °C freezer for a few days and scan the film in a dark room using the OPTIMAX™ X-ray processor (see Fig. 2).

3.5  Enzyme Assays 3.5.1  Labeling of Oligonucleotides

1. Anneal 2 μL of 10 μM DP (downstream primer) to 5 μL of 10  μM T (template) in the presence of 10 μL of water and 3 μL of 10× React2 buffer. 2. Incubate reaction at 95 °C for 5 min and transfer to 70 °C heat block. Turn off heat block to slowly cool down the reaction to room temperature for 1.5 h. 3. Upon the completion of the incubation reaction, add 4 μL of label, 1 μL of Klenow fragment, and 5 μL of nuclease-free water to make a final volume of 30 μL. Incubate reaction for at 1.5 h at 37 °C. 4. While this incubation is going on, prepare a sequencing gel (see Note 19). Additionally, prepare spin columns by following the manufacturer’s instructions.

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CBB

+ + -

H3/H4 FEN1 p300 Ac-CoA

+ + +

217

X-Ray

+ + -

+ + +

+ + -

+ + +

+ + -

+ + + -- p300

-- FEN1

-- H3/H4

1

2

3

4

1

2

3

4

Fig. 2 Assessing FEN1 in vitro lysine acetylation by autoradiography. In vitro acetylated H3/H4 peptide and FEN1 was subjected to SDS-PAGE analysis and stained using Coomassie brilliant blue (CBB). The same gel was subsequently analyzed by autoradiography (X-ray). Acetylated H3/H4, FEN1, and autoacetylated p300 are indicated

5. Add 10 μL of nuclease-free water to incubated samples to bring the final volume up to 40 μL. 6. Pipet the labeled sample into the spin column and centrifuge for 5 min at 1000 × g. 7. While the sample is centrifuging, cut up a piece of Whatman filter paper into one-inch squares. Also, pre-run the sequencing gel in 1× TBE at 80 W. 8. In triplicates, spot 0.5 μL of each sample on a filter paper and place in the scintillation vials filled with 4 mL of Ecoscint. Measure the amount of radionucleotide incorporated using the scintillation counter. Save counts as “after” reading. 9. To the remaining sample, add 20 μL of 2× termination dye and heat at 95 °C for 5 min. 10. Load the samples on the gel and electrophorese for 1 h 45 min (see Note 20). 11. Carefully remove one of the plates used to expose the gel. Cover it with plastic wrap and using autoradiography tape, mark different positions on the gel to ensure proper orientation required for gel extraction in later steps. 12. In a dark room, expose the gel to an autoradiography film for about a minute and develop in the X-ray processor.

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13. Position the developed film on the gel and cut out the bands containing the labeled oligonucleotide. Put the gel piece in 500 μL of TE and rotate overnight. 14. Transfer the gel pieces and DNA in TE to a Spin-X filter column and centrifuge for 2 min at 1000 × g to separate the gel pieces from the DNA. Discard the column, but retain the flow through. 15. To the tube containing the flow through, add 1500 μL 90% Ethanol, 12.5 μL sodium acetate, and 3 μL paint pellet. Incubate for at least 1 h in −80 °C (see Note 21). 16. Centrifuge at 15,000 × g for 20 min. Discard supernatant and ensure pellet is not disturbed in the process. 17. Add 500  μL of 70% ethanol and centrifuge at 15,000 × g for 10 min to wash the pellet. Discard supernatant. 18. Allow the pellet to dry in the 37 °C heat block for 10 min. Keep the lid open, but the work area covered with appropriate PPE as mentioned in Note 20. 19. While the pellet is drying, aliquot 4 mL Ecoscint into scintillation vials. 20. Resuspend pellet in 30 μL of TE. 21. In triplicates, spot 0.5 μL of sample on precut filter paper and place in scintillation vials. Measure the amount of radionucleotide incorporated using the scintillation counter. Save counts as “recovery” reading. 22. Calculate the volume of labeled primer that accounts for 1 μM or 1 pmol/μL (see Note 22). Anneal to the template and upstream unlabeled oligos in the ratio 1:2:4, respectively. Annealing reaction should be done in duplex annealing buffer. Heat at 95 °C for 5 min and gradually cool down to room temperature by turning off incubator overnight (see Notes 23 and 24). 3.5.2  Nuclease Assay

1. Make a 15% denaturing gel and allow to polymerize (see Note 19). 2. Set up a standard reaction with varying concentrations of unmodified and acetylated forms of FEN1 in nuclease reaction buffer to a final volume of 20 μL (see Note 25). 3. Add 5 nM of the annealed substrate to each reaction and incubate the reaction for 10 min at 37 °C. 4. Pre-run gel at 80 W on S2 gel electrophoresis apparatus in 1× TBE while samples are incubating (see Note 26). 5. After incubation, add 20 μL of termination dye to samples and heat at 95 °C for 5 min. Spin tubes to recover sample in condensate. 6. Load 20 μL of sample onto the denaturing gel for 90 min (see Note 27).

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* 5’

FEN1

-

3’

+ p300

Unmodified

Acetylated

-- Substrate

-- FEN1 Cleavage Product

1

2

3

4

5

6

7

Fig. 3 FEN1 Cleavage Assay. A 15-nt flap substrate labeled at the 5′ end of the downstream primer was used to measure the 5′–3′ endonuclease activity of hFEN1. Reactions containing 5 fmol of substrate and increasing concentrations (10, 20, and 50 fmol) of unmodified FEN1 and acetylated FEN1 (Ac-FEN1) were incubated for 10 min at 37 °C. The labeled substrate is depicted above the gel with the asterisk indicating the site of the 32P label. The substrate alone, FEN1 cleavage product, and cleavages beyond the base of the flap are indicated beside the gel

7. Using vacuum, dry gel in the gel dryer for about 1 h. 8. Expose dried gel to phosphor screen in a cassette overnight. 9. Scan and image results using Typhoon™ FLA 9500 laser scanner as shown in Fig. 3. 10. Compare the amount of nuclease activity by quantifying the intensity of each band produced using densitometry (see Note 28). 3.5.3  Electromobility Gel Shift Assay to Analyze Protein-DNA Binding

1. Make a 6% native gel and allow to polymerize. Once polymerized, set up native gel electrophoresis apparatus and pre-run gel in 1× TBE buffer. 2. Set up a standard reaction with varying concentrations of unmodified and acetylated forms of FEN1 in EMSA reaction buffer to final volume of 20 μL. 3. Add 5 nM of the annealed substrate to each reaction and incubate the reaction for 10 min at 37 °C. 4. Add 20 μL of 2× Termination dye to stop the reaction and load onto the pre-run gel for 1 h at 180 V. 5. Similarly, dry gel for an hour and expose to a phosphor screen in a cassette overnight. 6. Scan and image results using Typhoon™ FLA 9500 laser scanner as shown in Fig. 4.

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* 5’

FEN1

-

Unmodified

3’

+ p300

Acetylated

-- Substrate Bound FEN1 -- Substrate

1

2

3

4

5

6

7

Fig. 4 FEN1 binding assay. Substrate binding efficiency of FEN1 and Ac-FEN1 was studied using electromobility gel shift assay. Five fmol of 15-nt substrate was incubated with increasing concentrations (10, 20, and 50 fmol) of either unmodified FEN1 and acetylated FEN1, and the reactions were incubated for 10 min at room temperature and separated on a 6% polyacrylamide gel. The labeled substrate is depicted above the gel with the asterisk indicating the site of the 32P label. The substrate alone and complexes containing FEN1-bound substrate are indicated beside the gel at the right

4  Notes 1. pET-FCH FEN1 plasmid is designed and developed as previously outlined [19]. 2. The minimal HAT domain (residues 1284–1669) of the human p300 sequence was synthesized by Genscript Biotech Science Research Company. The p300-HAT-pUC57-Kan plasmid was created by inserting the sequence into the ECoR1-­ Xho1 site of pUC57-Kan. Plasmid can be obtained by contacting corresponding author. 3. When making autoinduction media, it is important to autoclave the ingredients separately. Peptone and yeast extract are dissolved in 700 mL of distilled water and autoclaved in a 2800 mL flask. We find that autoclaving stocks of 100 mL aliquots of glucose and lactose and 90 mL of KPi allows for a smooth experimental set up. Similarly, autoclave glycerol in a 250 mL bottle. Simply add the individual aliquots to the 700 mL media to make 1 L of autoinduction medium. 4. One sheet of chromatography paper can be cut into 4 squares. 5. The nicked flap substrate consists of a template (T), a downstream flap (DS), and an upstream (US) annealed together in duplex buffer. The sequences used can be found in Table 1.

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Table 1 Designed primer sequence for creating annealed nicked substrate Primer

Sequence

Template (T)

5′/biotin/GTACCGAGCTCGAATTCGCCCGTTTCACGCCTGTTAG TTAATTCACTGGCCGTCGTTTTACAACGACGTGACTGGG-3′

Downstream primer (DP) 5′-GCCCAGTCACGTCGTTGTAAAACGGGTCGTGACTGGGAAAAC CCTGGCG-3′ Upstream primer (UP)

5′-CGCCAGGGTTTTCCCAGTCACGACA-3′

6. After sonication, the cell lysate should appear light and not too viscous. If the sample is still viscous, sonicate for an additional 5 min. Also ensure that the pellet is completely resuspended and there are no pellet chunks floating around. 7. Set about 20 μL of supernatant aside to load in the crude-free extract lane on the gel. 8. The protein tends to precipitate during dialysis. Fortunately for this prep, most of the protein still remained in solution. To circumvent this problem, dilute the protein from the His column tenfold with hFEN1 storage buffer before dialysis. 9. Inject 500 μL into the SEC and freeze the other samples at −80 °C. 10. All buffers must be filtered prior to using them with the SEC. 11. Column care:

(a) Ni-NTA column cleanup and storage: wash with 10 CV of 1 M imidazole, then wash with 10 CV of water, then wash with 5 CV of 20% ethanol. Store the column in 20% ethanol.



(b) The size exclusion column should be cleaned with 5 CV Milli-Q water and stored in 20% ethanol.

12. Analyze this sample on an SDS-page gel to ascertain if some of the protein was lost during this step. 13. One can use any commercially available HAT activity kit or use the HPLC methodology outlined in a different chapter in this book. 14. We have found that the acetylation reaction is the most efficient when reagents are added in this particular order: 1× HAT buffer, protein to be acetylated, HAT (p300) and Acetyl Co-A. 15. In vitro acetylation of proteins should be performed in a 1:0.1:10 ratio (protein to be acetylated:HAT:Acetyl-CoA). However, this may differ for every protein and needs to be empirically determined.

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16. Note that steps 1–3 are common for both detection methods, but steps 4–10 are for western blot detection and steps 11–14 are for detection using autoradiography. 17. The sandwich is made in this order: sponge on the bottom, a filter paper, the gel to be transferred, the PVDF membrane, a filter paper, and finally another sponge. PVDF membrane should be activated in methanol for a minute and soaked in transfer buffer alongside the other components before the sandwich is made. 18. This step should be done in a dark room as exposure to light destroys the film. 19. When making a sequencing or denaturing gel, add distilled water, TBE and urea to a container with a stir bar. Microwave for 20 s to aid in dissolving urea. Place container on the stir plate and add the other components until all the urea is completely dissolved and the solution is clear. 20. It is important to use proper protective equipment when working with radioactivity. Gels should be electrophoresed behind a shield. 21. You can add 1 μL of glycogen to the sample to ensure you get a well-formed pellet if you obtain a diffused pellet. 22. To calculate the volume of labeled oligo required for 1 pmol/μL, use the calculations in the excel sheet as shown in Fig. 5.

Fig. 5 Calculations for determining the concentration of 5′ end labeled substrates. This figure provides an example of the template used to calculate the amount of radiolabeled primer to be used for annealing substrates

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23. Final concentration for annealing reaction should be 10 nM (fmol/uL) of labeled primer. An example of an annealing reaction with a final volume of 100uL and concentration of 10 nM: Labeled primer = 3.6 μL calculated volume required for 1 μM Template (1 μM) = 2 μL Upstream (1 μM) = 4 μL Duplex buffer = 90.4 μL Total = 100 μL of 10 nM annealed substrate 24. Check to ensure complete annealing occurs by electrophoresing annealed sample and labeled primer on a 6% native gel. Annealed primer should have a higher band than the labeled primer. 25. Our working range is between 10–50 nM of protein for these reactions. 26. It is important to wash the wells to get rid of gel pieces that might interfere with the samples and prevent them from properly migrating through the gel. 27. We generally save the remaining 20 μL sample and store in −20 °C in case that batch of experiments need to be repeated. 28. Densitometry calculation: {(b)/(b + a)} × 100. For % cleavage, b is the cleaved product and a is the uncleaved substrate remaining while for % binding, b is the bound product and a is the unbound substrate remaining.

Acknowledgments This work was supported by NIH Grant GM0938328 and New Faculty Start-Up Funds from Indiana University Purdue University Indianapolis (IUPUI). References 1. Dawson MA, Kouzarides T (2012) Cancer epigenetics: from mechanism to therapy. Cell 150(1):12–27. https://doi.org/10.1016/j. cell.2012.06.013 2. Drazic A, Myklebust LM, Ree R, Arnesen T (2016) The world of protein acetylation. Biochim Biophys Acta 1864(10):1372–1401. https:// doi.org/10.1016/j.bbapap.2016.06.007 3. Hasan S, Stucki M, Hassa PO, Imhof R, Gehrig P, Hunziker P, Hubscher U, Hottiger MO (2001) Regulation of human flap endonuclease-1 activity by acetylation through the tran-

scriptional coactivator p300. Mol Cell 7(6):1221–1231. pii: S1097-2765(01)00272-6 4. Hasan S, El-Andaloussi N, Hardeland U, Hassa PO, Burki C, Imhof R, Schar P, Hottiger MO (2002) Acetylation regulates the DNA end-trimming activity of DNA polymerase beta. Mol Cell 10(5):1213–1222. pii: S1097276502007451 5. Balakrishnan L, Stewart J, Polaczek P, Campbell JL, Bambara RA (2010) Acetylation of Dna2 endonuclease/helicase and flap endonuclease 1 by p300 promotes DNA stability by

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creating long flap intermediates. J Biol Chem 285(7):4398–4404. pii: M109.086397. https://doi.org/10.1074/jbc.M109.086397 6. Kang K, Choi JM, Fox JM, Snyder PW, Moustakas DT, Whitesides GM (2016) Acetylation of surface lysine groups of a protein alters the organization and composition of its crystal contacts. J Phys Chem B 120(27): 6461–6468. https://doi.org/10.1021/ acs.jpcb.6b01105 7. Naryzhny SN, Lee H (2004) The post-­ translational modifications of proliferating cell nuclear antigen: acetylation, not phosphorylation, plays an important role in the regulation of its function. J Biol Chem 279(19):20194– 20199. https://doi.org/10.1074/jbc. M312850200. pii: M312850200 8. Thevenet L, Mejean C, Moniot B, Bonneaud N, Galeotti N, Aldrian-Herrada G, Poulat F, Berta P, Benkirane M, Boizet-Bonhoure B (2004) Regulation of human SRY subcellular distribution by its acetylation/deacetylation. EMBO J 23(16):3336–3345. https://doi. org/10.1038/sj.emboj.7600352 9. Valacco MP, Varone C, Malicet C, Canepa E, Iovanna JL, Moreno S (2006) Cell growth-­ dependent subcellular localization of p8. J Cell Biochem 97(5):1066–1079. https://doi. org/10.1002/jcb.20682 10. di Bari MG, Ciuffini L, Mingardi M, Testi R, Soddu S, Barila D (2006) c-Abl acetylation by histone acetyltransferases regulates its nuclearcytoplasmic localization. EMBO Rep 7(7):727–733. https://doi.org/10.1038/ sj.embor.7400700 11. Caron C, Boyault C, Khochbin S (2005) Regulatory cross-talk between lysine acetylation and ubiquitination: role in the control of protein stability. Bioessays 27(4):408–415. https://doi.org/10.1002/bies.20210 12. Choudhary C, Kumar C, Gnad F, Nielsen ML, Rehman M, Walther TC, Olsen JV, Mann M (2009) Lysine acetylation targets protein com-

plexes and co-regulates major cellular functions. Science 325(5942):834–840. pii: 1175371. https://doi.org/10.1126/science.1175371 13. Choudhary C, Weinert BT, Nishida Y, Verdin E, Mann M (2014) The growing landscape of lysine acetylation links metabolism and cell signalling. Nat Rev. Mol Cell Biol 15(8):536– 550. https://doi.org/10.1038/nrm3841 14. Scholz C, Weinert BT, Wagner SA, Beli P, Miyake Y, Qi J, Jensen LJ, Streicher W, McCarthy AR, Westwood NJ, Lain S, Cox J, Matthias P, Mann M, Bradner JE, Choudhary C (2015) Acetylation site specificities of lysine deacetylase inhibitors in human cells. Nat Biotechnol 33(4):415–423. https://doi. org/10.1038/nbt.3130 15. Gill G (2004) SUMO and ubiquitin in the nucleus: different functions, similar mechanisms? Genes Dev 18(17):2046–2059. https://doi.org/10.1101/gad.1214604 16. Huang R, Holbert MA, Tarrant MK, Curtet S, Colquhoun DR, Dancy BM, Dancy BC, Hwang Y, Tang Y, Meeth K, Marmorstein R, Cole RN, Khochbin S, Cole PA (2010) Site-­ specific introduction of an acetyl-lysine mimic into peptides and proteins by cysteine alkylation. J Am Chem Soc 132(29):9986–9987. https://doi.org/10.1021/ja103954u 17. Guo J, Wang J, Lee JS, Schultz PG (2008) Site-specific incorporation of methyl- and acetyl-­ lysine analogues into recombinant proteins. Angew Chem Int Ed Engl 47(34): 6399–6401. https://doi.org/10.1002/ anie.200802336 18. Neumann H, Peak-Chew SY, Chin JW (2008) Genetically encoding N(epsilon)-acetyllysine in recombinant proteins. Nat Chem Biol 4(4):232–234. https://doi.org/10.1038/ nchembio.73 19. Bornarth CJ, Ranalli TA, Henricksen LA, Wahl AF, Bambara RA (1999) Effect of flap modifications on human FEN1 cleavage. Biochemistry 38(40):13347–13354

Chapter 13 Examining the Role of HDACs in DNA Double-Strand Break Repair in Neurons Ping-Chieh Pao, Jay Penney, and Li-Huei Tsai Abstract Histone deacetylases (HDACs) modulate chromatin structure by removing acetyl groups from histones. Upon DNA double-strand breaks (DSBs), deacetylation of H3K56 and H4K16 by HDACs occurs immediately at break sites, and is crucial for DSB repair. Here we describe two assays that examine defective DSB repair caused by HDAC inhibition in primary cortical neurons: single-cell gel electrophoresis to assay DNA integrity (the comet assay) and western blot analysis for γH2AX, a phosphorylated histone variant associated with DSBs. Key words Genome stability, DNA damage, Histone deacetylation, Histone deacetylases (HDACs), DNA repair, DNA double-strand breaks (DSBs), Single-cell gel electrophoresis assay, Comet assay, γH2AX

1  Introduction Histone deacetylases (HDACs) are a group of enzymes that remove acetyl groups from lysine residues on histones to regulate multiple cellular functions including gene expression and DNA double-­strand break (DSB) repair [1]. Deacetylation of H3K56 and H4K16 by HDACs occurs rapidly after DSB induction, and is essential for DNA repair [2]. Upon DNA DSB formation in post-­mitotic neurons, HDAC1, one of the HDAC family members, is recruited to break sites, where it promotes DNA repair through the non-homologous end-joining pathway. Neurons lacking HDAC1 exhibit higher susceptibility to DSB-inducing agents and repair DSBs less efficiently [3, 4]. Blocking HDAC activity using HDAC inhibitors increases DSBs in mouse brain [5] and compromises DSB repair in U2OS and MCF-7 cells [2, 6]. Here, we describe the comet assay and γH2AX immunoblotting, two methods commonly used in our laboratory to assess the effects of HDAC inhibitors on DSB repair in neurons. To induce DSBs, we treated mouse primary neurons with the DSB-inducing Robert M. Brosh, Jr. (ed.), Protein Acetylation: Methods and Protocols, Methods in Molecular Biology, vol. 1983, https://doi.org/10.1007/978-1-4939-9434-2_13, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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agent ­etoposide (ETP) for 1 h. After washing out ETP, neurons were allowed to recover (and to repair DNA damage) with or without trichostatin A (TSA), a HDAC inhibitor. Following recovery for 10 h, cells were harvested for the comet assay and western blotting. The comet assay is a sensitive method to detect physical breaks in DNA on a single-cell level. In this assay, cells are first embedded in agarose, lysed, and then subjected to electrophoresis. Fragmented or damaged DNA migrates faster than intact DNA and resembles a comet, with the length and intensity of tail indicating the extent of damage [7] (Fig. 1). Unwinding DNA and performing electrophoresis in alkaline buffer (pH > 13) is able to detect single-strand breaks (SSBs), DSBs, and alkali-labile lesions [8]. A neutral pH comet assay can be used (see Note 1) alternatively if you are not using a DSB-inducing agent, like ETP. As a complimentary assay to examine the extent of DSBs in a population of cells, we utilized western blotting to determine the level of histone H2A variant X with phosphorylation at Ser139 (γH2AX), a well-­ established DSB marker [9]. Cells harvested before and immediately after ETP treatment (baseline DSBs and before DSB repair has occurred, respectively) and cells allowed to recover in the absence of TSA for 10 h were included as controls.

2  Materials Prepare all solutions using ddH2O, unless indicated otherwise. 2.1  Compound Treatment of Primary Cortical Neurons

1. Primary mouse cortical neurons, cell density: 5 × 106 cells in a 10-cm culture dish. 2. Neuronal media: Neurobasal media, B27 supplement, 5 mM l-glutamine and 1% penicillin and streptomycin. 3. Dimethyl sulfoxide (DMSO). 4. 10 mM Trichostatin A (TSA): Dissolve 1 mg of TSA in 0.33 mL DMSO. Aliquot and store at −20 °C. 5. 25 mM Etoposide (ETP): Dissolve 25 mg of ETP in 1.7 mL DMSO. Aliquot and store at −20 °C.

2.2  Comet Assay

1. PBS (Phosphate-buffered saline): 1.06 mM KH2PO4, 155.17 mM NaCl, 2.97 mM Na2HPO4, pH 7.4. Store at 4 °C. 2. 1% Low melting point (LMP) agarose: Mix 1 g UltraPure™ LMP agarose and 100 mL PBS in a glass flask. Microwave for 2–3 min until the agarose is completely dissolved. 3. Heat block for microcentrifuge tubes. 4. CometSlide™ 2 Well.

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Electrophoresis -

+

Head

Tail

Intact DNA

Fragmented or damaged DNA

Fig. 1 A typical damaged cell in comet assay

5. CometAssay® Lysis solution. 6. Alkaline buffer: 300 mM NaOH, 1 mM EDTA. 7. CometAssay® Electrophoresis System II and power supply. 8. Neutralization buffer: 0.4 M Tris–HCl (pH 7.4). Store at 4 °C. 9. 70% Ethanol. Store at room temperature. 10. TE buffer: 10 mM Tris–HCl (pH 8.0), 1 mM EDTA (pH 8.0). Store at room temperature. 11. SYBR Gold nucleic acid staining solution: Add 10 μL SYBR Gold stock (10,000× concentrate in DMSO) into 100 mL TE buffer for 10 CometSlides. Aliquot SYBR Gold stock and store at −20 °C. Discard excess SYBR Gold once thawed. 12. Fluoromount-G slide mounting medium. 13. Rectangular coverglass: No. 1½, 24 × 60 mm. 14. Clear nail polish. 15. Epifluorescence or confocal microscope. 16. OpenComet software [10]: a free plugin for ImageJ, which can be downloaded at http://www.cometbio.org/index.html. 2.3  Western Blot Analysis for γH2AX

1. PBS (Phosphate-buffered saline): 1.06 mM KH2PO4, 155.17 mM NaCl, 2.97 mM Na2HPO4, pH 7.4. Store at 4 °C. 2. Cell scrapers. 3. Cytoplasmic extraction buffer (CEB) and nuclear extraction buffer (NEB) from subcellular protein fractionation kit for tissues (Thermo Fisher 87790). 4. Protease inhibitor cocktail.

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5. Phosphatase inhibitor cocktail. 6. Histone extraction buffer: 1 mL NEB, 50 μL 100 mM CaCl2 and 300 units of micrococcal nuclease. 7. BCA protein assay kit. 8. 5× SDS protein loading buffer: 250 mM Tris–HCl (pH 6.8), 10% SDS, 50% glycerol, 0.5% bromophenol blue and 5% β-mercaptoethanol. 9. 16% SDS polyacrylamide gel. 10. Running buffer: 0.2 M glycine, 0.025 M Tris base and 0.1% SDS. 11. Wet transfer buffer: Dissolve 3.032 g Tris, 14.416 g glycine and 1 g SDS in 800 mL ddH2O, and mix with 200 mL methanol. 12. 3MM filter paper. 13. 0.45 μm PVDF membrane. 14. 100% Methanol. 15. Tris buffered saline with Tween (TBST): 10 mM Tris–HCl (pH 8.0), 150 mM NaCl and 0.05% Tween-20. 16. 5% Nonfat milk blocking buffer: Dissolve 5 g nonfat dry milk in 100 mL TBST. 17. γH2AX antibody (Abcam, ab81299). 18. Histone H3 antibody (Abcam, ab24834). 19. HRP conjugated secondary antibody. 20. ECL Western blotting substrate. 21. Film. 22. Western blot stripping buffer (Thermo Fisher 46430).

3  Methods Please refer to the following chapter for preparation of primary neuronal cultures [11]. 3.1  Compound Treatment of Primary Cortical Neurons

1. Plate 5 × 106 cells in a 10-cm culture dish with 10 mL neuronal media. 2. The next day (1 day in vitro, 1 DIV), add an additional 5 mL fresh neuronal media. 3. Withdraw 3 mL of old media from the dish, and replenish with 5 mL fresh neuronal media on 3, 7 and 10 DIV. 4. On 13 DIV, adjust media volume to 10 mL in each 10-cm dish using 25 mL serological pipet.

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5. Feed neurons with 5 mL fresh neuronal media. 6. On 14 DIV, transfer 8 mL media (“conditioned media”) from each dish to a 15 mL tube. 7. Keep conditioned media in a 37 °C water bath. 8. Add 1.4 μL of 25 mM ETP, or DMSO as control, to each dish for induction of DSBs. 9. Incubate treated cells at 37 °C in a tissue culture incubator for 1 h. 10. Prepare TSA-containing conditioned media. Add 2 μL of 10 mM TSA (or DMSO as control) to 8 mL warm conditioned media collected previously. 11. Replace all old media with TSA (or DMSO)-containing conditioned media for 10 h to allow neurons to recover (see Note 2). 3.2  Comet Assay 3.2.1  Harvesting Cells for Comet Assay

1. Aspirate culture media, and wash cells with cold PBS once. 2. Add 5 mL cold PBS, and collect cells using a cell scraper. 3. Transfer cells to 15 mL tubes, and pellet down at 500 × g for 5 min at 4 °C. 4. Aspirate the supernatant and gently resuspend cells in 1 mL cold PBS. 5. Transfer to 1.5 mL Eppendorf tubes. 6. Centrifuge at 500 × g for 5 min at 4 ° C, and wash once with 1 mL of cold PBS then pellet the cells again. 7. Remove the supernatant, and store cell pellets at −80 °C until use.

3.2.2  Embedding Cells in LMP Agarose and Lysis Overnight

1. Prepare 1% LMP agarose, and keep 450 μL LMP agarose aliquots at 42 °C for least 30 min before embedding. 2. Aliquot 100 mL Trevigen lysis solution (enough for ~10 CometSlides) and cool to 4 °C for at least 30 min prior to use. 3. Label CometSlides using a pencil. Avoid using a pen as the label may come off in ethanol. 4. Thaw cell pellets on ice for 5 min. 5. Resuspend cells in 1 mL cold PBS gently. 6. Adjust cell density to ~4 × 105 cells/mL with cold PBS. 7. Add 50 μL diluted cells to 450 μL LMP agarose at 42 °C. 8. Vortex gently, and then apply 50 μL mixture onto each well of CometSlides. Spread agarose using the side of the pipette tip. 9. Keep slides at 4 °C in the dark for 30 min. 10. Immerse slides in cold lysis solution overnight (14–16 h) in the dark at 4 °C.

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3.2.3  Alkaline Unwinding, Electrophoresis, and Staining

1. Freshly prepare 1 L of alkaline buffer, and cool to 4 °C before use. 2. Incubate slides with cold alkaline buffer at 4 °C for 1 h in the dark (see Note 3). 3. Perform electrophoresis in 850 mL alkaline buffer at 4 °C for 30 min at 1 V/cm (distance between electrodes) (see Note 4). 4. Immerse slides in cold neutralization buffer at 4 °C 2× 15 min with shaking in the dark. 5. Incubate slides with 70% ethanol for 30 min at room temperature with shaking. 6. To bring cells into the sample plane, dry samples at 37 °C for 15–20 min. Samples can be stored at room temperature with desiccant prior to imaging. 7. Stain cells with SYBR Gold (1:10,000) for 30 min in the dark at room temperature with shaking. 8. Wash briefly with water and dry slides at 37 °C for 15–20 min. 9. Mount slides and cover with coverslip. Seal slides with clear nail polish. 10. Slides can be stored at 4 °C in the dark at least a month.

3.2.4  Imaging and Analysis

1. Visualize slides with epifluorescence or confocal microscope, and take tile scan image using 5× objective to obtain more comets if possible. The excitation/emission wavelength of SYBR Gold is 496/522 nm. 2. Analyze comet images using default settings in OpenComet software: Use background correction and auto head finding. Detailed instructions and video tutorial are available at: http:// www.cometbio.org/index.html. 3. Open the output images and exclude overlapping comets identified by OpenComet manually. 4. Analyze images (at least 100 comets in each group) for several features including tail length, percent DNA in tail, and tail moment (tail length × % DNA in tail) (Fig. 2b). 5. Take representative images using 20× objective after the analysis (Fig. 2a).

3.3  Western Blot Analysis for γH2AX

1. Aspirate culture media, and wash cells with cold PBS once. 2. Add 5 mL cold PBS, and collect cells using a cell scraper. 3. Transfer to 15 mL tubes, and pellet down at 500 × g for 5 min at 4 °C. 4. Aspirate the supernatant and gently resuspend cells in 1 mL cold PBS.

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(A)

ETP_1 h ETP_1 h Rec. 10 h with DMSO Rec. 10 h with TSA

ETP_1 h

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(B)

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Fig. 2 Pharmacological inhibition of HDAC by TSA impairs DSB repair in neurons. Comet assays were performed on the untreated cells, cells treated with ETP for 1 h, and ETP treatment followed by 10 h recovery (Rec.) with DMSO or TSA. Representative images (a), and the quantification of tail length, % tail DNA and tail moment (b) from comet assays. Data are presented as mean ± s.e.m. (**P