Homologous Recombination: Methods and Protocols [1st ed.] 9781071606438, 9781071606445

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Methods in Molecular Biology 2153

Andrés Aguilera Aura Carreira Editors

Homologous Recombination Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

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

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

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-bystep fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.

Homologous Recombination Methods and Protocols

Edited by

Andrés Aguilera Centro Andaluz de Biología Molecular y Medicina Regenerativa, CABIMER, Universidad de Sevilla, Sevilla, Spain

Aura Carreira Institut Curie, PSL Research University, CNRS, UMR3348, F-91405, Orsay, France; University Paris Sud, Paris-Saclay University, CNRS, UMR3348, F-91405, Orsay, France

Editors Andre´s Aguilera Centro Andaluz de Biologı´a Molecular y Medicina Regenerativa, CABIMER Universidad de Sevilla Sevilla, Spain

Aura Carreira Institut Curie, PSL Research University, CNRS UMR3348, F-91405 Orsay, France University Paris Sud, Paris-Saclay University, CNRS UMR3348, F-91405 Orsay, France

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-0643-8 ISBN 978-1-0716-0644-5 (eBook) https://doi.org/10.1007/978-1-0716-0644-5 © Springer Science+Business Media, LLC, part of Springer Nature 2021 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, expressed 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: This image was prepared by Aura Carreira and Teresa Moreno Rodriguez. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface DNA double-strand breaks (DSBs) are the most harmful lesions to DNA in the cell. To cope with these insults, all organisms have devised two main types of evolutionary conserved mechanisms for their repair, homologous recombination (HR), and non-homologous end joining (NHEJ). The first one operates predominantly during the S/G2 phase of the cell cycle, when the sister chromatid is available for repair. Because HR requires homology for repair, this pathway is considered essentially error-free. NHEJ is the pathway of choice in the other phases of the cell cycle, including G1. This pathway is generally faithful but can be prone to errors. Most spontaneous DNA breaks arising in somatic cells occur randomly as a consequence of DNA replication failure caused by either DNA lesions or generated by obstacles that impede the progression of the replication fork (e.g., protein-bound to DNA, DNA secondary structures, replication–transcription conflicts, etc.). For this reason, HR is a major DNA repair pathway during S/G2 phases of the cell cycle. Thus, HR is intimately ligated to the prevention of genome instability in replicating somatic cells. In meiotic cells however, DSBs are developmentally controlled by the action of specific endonucleases where HR is essential; gametogenesis is not possible in the absence of HR. Genome instability and in particular defective HR is a common feature of a number of genetic diseases including cancer. Defects in HR in meiotic cells can lead to birth defects such as Down syndrome. Considering the relevance of HR as one of the major DSB repair pathways in mitotically cycling cells, as well as its essential role in meiosis, understanding the molecular mechanisms and factors that participate in HR is of key importance in Molecular Biology and Biomedicine. In this book, we compile a series of laboratory protocols covering the analysis of different steps of the homologous recombination process from the genetic, molecular biology, and cell biology perspectives. As these steps are very well conserved through evolution, taking advantage of different model organisms have led to accelerated discoveries in this field. Thus, when appropriate, some of the protocols we present here are explained in the context of more than one model system. We hope this book will facilitate the use of both classical and more recent approaches to answer specific questions on HR mechanisms as well as to decipher the function of novel factors involved in HR. We expect that this compilation of protocols elaborated by leading experts in the field will be useful not only to the scientific community working in genome integrity but also to scientists working in other areas such as cancer biology or cell cycle with renovated interests in HR and DSB repair. Sevilla, Spain Orsay, France

Andre´s Aguilera Aura Carreira

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

1 Detection of DNA Double-Strand Breaks by γ-H2AX Immunodetection. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sonia I. Barroso and Andre´s Aguilera 2 END-seq: An Unbiased, High-Resolution, and Genome-Wide Approach to Map DNA Double-Strand Breaks and Resection in Human Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nancy Wong, Sam John, Andre´ Nussenzweig, and Andres Canela 3 Resection of a DNA Double-Strand Break by Alkaline Gel Electrophoresis and Southern Blotting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Erika Casari, Elisa Gobbini, Michela Clerici, and Maria Pia Longhese 4 Analysis of DNA Double-Strand Break End Resection and Single-Strand Annealing in S. pombe . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Zhenxin Yan, Sandeep Kumar, and Grzegorz Ira 5 Quantifying DNA End Resection in Human Cells . . . . . . . . . . . . . . . . . . . . . . . . . . Yi Zhou and Tanya T. Paull 6 Genetic and Molecular Approaches to Study Chromosomal Breakage at Secondary Structure–Forming Repeats . . . . . . . . . . . . . . . . . . . . . . . . . Anissia Ait Saada, Alex B. Costa, and Kirill S. Lobachev 7 Biochemical Analysis of D-Loop Extension and DNA Strand Displacement Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Youngho Kwon and Patrick Sung 8 DNA Strand Exchange to Monitor Human RAD51-Mediated Strand Invasion and Pairing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sudipta Lahiri and Ryan B. Jensen 9 Monitoring Homologous Recombination Activity in Human Cells . . . . . . . . . . . ˚ sa Ehle´n, and Aura Carreira Domagoj Vugic, A 10 Interhomolog Homologous Recombination in Mouse Embryonic Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fabio Vanoli, Rohit Prakash, Travis White, and Maria Jasin 11 Branch Migration Activity of Rad54 Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Olga M. Mazina and Alexander V. Mazin 12 Holliday Junction Resolution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Raquel Carreira, F. Javier Aguado, Tomas Lama-Diaz, and Miguel G. Blanco 13 Identification and Analysis of Different Types of UFBs . . . . . . . . . . . . . . . . . . . . . . Simon Gemble and Mounira Amor-Gue´ret 14 Intrachromosomal Recombination in Yeast . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anastasiya Epshtein, Lorraine S. Symington, and Hannah L. Klein

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Genome-Wide Analysis of Mitotic Recombination in Budding Yeast . . . . . . . . . . Lydia R. Heasley, Nadia M. V. Sampaio, and Juan Lucas Argueso Monitoring Gene Conversion in Budding Yeast by Southern Blot Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Miyuki Yamaguchi and James E. Haber DNA Double-Strand Break-Induced Gene Amplification in Yeast . . . . . . . . . . . . . Tomas Strucko, Michael Lisby, and Uffe Hasbro Mortensen Measuring Chromosome Pairing During Homologous Recombination in Yeast. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fraulin Joseph, So Jung Lee, Eric Edward Bryant, and Rodney Rothstein Cytological Monitoring of Meiotic Crossovers in Spermatocytes and Oocytes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yan Yun, Masaru Ito, Sumit Sandhu, and Neil Hunter Detection of DSBs in C. elegans Meiosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tatiana Garcı´a-Muse Methods to Map Meiotic Recombination Proteins in Saccharomyces cerevisiae. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Aurore Sanchez and Vale´rie Borde Investigation of Break-Induced Replication in Yeast . . . . . . . . . . . . . . . . . . . . . . . . . Beth Osia, Rajula Elango, Juraj Kramara, Steven A. Roberts, and Anna Malkova Measurement of Homologous Recombination at Stalled Mammalian Replication Forks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nicholas A. Willis and Ralph Scully Super-Resolution Imaging of Homologous Recombination Repair at Collapsed Replication Forks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Donna R. Whelan and Eli Rothenberg The Analysis of Recombination-Dependent Processing of Blocked Replication Forks by Bidimensional Gel Electrophoresis . . . . . . . . . . . . . Karol Kramarz, Anissia Ait Saada, and Sarah A. E. Lambert The Sister-Chromatid Exchange Assay in Human Cells . . . . . . . . . . . . . . . . . . . . . . Emanuela Tumini and Andre´s Aguilera Analysis of Recombination at Yeast Telomeres . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Marie-Noelle Simon, Dmitri Churikov, and Vincent Ge´li Gel Electrophoresis Analysis of rDNA Instability in Saccharomyces cerevisiae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mariko Sasaki and Takehiko Kobayashi Analyzing Homologous Recombination at a Genome-Wide Level . . . . . . . . . . . . Coline Arnould, Vincent Rocher, and Gae¨lle Legube CRISPR/Cas9-Induced Breaks in Heterochromatin, Visualized by Immunofluorescence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ioanna Mitrentsi and Evi Soutoglou

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In Vivo Binding of Recombination Proteins to Non-DSB DNA Lesions and to Replication Forks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Roma´n Gonza´lez-Prieto, Marı´a J. Cabello-Lobato, and Fe´lix Prado Live Cell Imaging of Nuclear Actin Filaments and Heterochromatic Repair foci in Drosophila and Mouse Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Colby See, Deepak Arya, Emily Lin, and Irene Chiolo In Vitro Characterization of Sumoylation of HR Proteins. . . . . . . . . . . . . . . . . . . . Veronika Altmannova and Lumir Krejci High-Throughput Analysis of Heteroduplex DNA in Mitotic Recombination Products . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Dionna Gamble, Yee Fang Hum, and Sue Jinks-Robertson Fluorescence Microscopy for Analysis of Relocalization of Structure-Specific Endonucleases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Carl P. Lehmann, Irene Saugar, and Jose´ Antonio Tercero Physical and Genetic Assays for the Study of DNA Joint Molecules Metabolism and Multi-invasion-Induced Rearrangements in S. cerevisiae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Aure`le Piazza, Pallavi Rajput, and Wolf-Dietrich Heyer

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors F. JAVIER AGUADO • Departamento de Bioquı´mica e Bioloxı´a Molecular, CIMUS, Universidade de Santiago de Compostela-Instituto de Investigacion Sanitaria (IDIS), Santiago de Compostela, Spain ANDRE´S AGUILERA • Centro Andaluz de Biologı´a Molecular y Medicina Regenerativa, CABIMER, University of Seville-CSIC-UPO, Seville, Spain ANISSIA AIT SAADA • School of Biological Sciences, Institute for Bioengineering and Bioscience, Georgia Institute of Technology, Atlanta, GA, USA VERONIKA ALTMANNOVA • Department of Biology, Masaryk University, Brno, Czech Republic; International Clinical Research Center, St. Anne’s University Hospital, Brno, Czech Republic MOUNIRA AMOR-GUE´RET • Institut Curie, PSL Research University, CNRS UMR 3348, Orsay, France; CNRS UMR 3348, Paris Saclay University, Institut Curie, Research Center, Orsay, France JUAN LUCAS ARGUESO • Department of Environmental and Radiological Health Sciences, Colorado State University, Fort Collins, CO, USA; Cell and Molecular Biology Graduate Program, Colorado State University, Fort Collins, CO, USA COLINE ARNOULD • LBCMCP, Centre de Biologie Integrative (CBI), CNRS, Universite´ de Toulouse, UT3, Toulouse, France DEEPAK ARYA • Molecular and Computational Biology Department, University of Southern California, Los Angeles, CA, USA SONIA I. BARROSO • Centro Andaluz de Biologı´a Molecular y Medicina Regenerativa, CABIMER, University of Seville-CSIC-UPO, Seville, Spain MIGUEL G. BLANCO • Departamento de Bioquı´mica e Bioloxı´a Molecular, CIMUS, Universidade de Santiago de Compostela-Instituto de Investigacion Sanitaria (IDIS), Santiago de Compostela, Spain VALE´RIE BORDE • Institut Curie—Research Center, UMR3244 CNRS, Pavillon Trouillet Rossignol, PSL Research University, Paris Cedex 05, France; Paris Sorbonne Universite´, Paris, France ERIC EDWARD BRYANT • Department of Biological Sciences, Columbia University, New York, NY, USA MARI´A J. CABELLO-LOBATO • Division of Cancer Sciences, Manchester Cancer Research Center, University of Manchester, Manchester, UK ANDRES CANELA • Laboratory of Genome Integrity, National Cancer Institute, NIH, Bethesda, MD, USA; The Hakubi Center for Advanced Research and Radiation Biology Center, Graduate School of Biostudies, Kyoto University, Kyoto, Japan AURA CARREIRA • Institut Curie, PSL Research University, CNRS, UMR3348, F-91405, Orsay, France; University Paris Sud, Paris-Saclay University, CNRS, UMR3348, F91405, Orsay, France RAQUEL CARREIRA • Departamento de Bioquı´mica e Bioloxı´a Molecular, CIMUS, Universidade de Santiago de Compostela-Instituto de Investigacion Sanitaria (IDIS), Santiago de Compostela, Spain ` degli Studi di ERIKA CASARI • Dipartimento di Biotecnologie e Bioscienze, Universita Milano-Bicocca, Milano, Italia

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Contributors

IRENE CHIOLO • Molecular and Computational Biology Department, University of Southern California, Los Angeles, CA, USA DMITRI CHURIKOV • Marseille Cancer Research Center (CRCM), U1068 Inserm, UMR7258 CNRS, Institut Paoli-Calmettes, Aix Marseille University, Marseille, France ` degli Studi di MICHELA CLERICI • Dipartimento di Biotecnologie e Bioscienze, Universita Milano-Bicocca, Milano, Italia ALEX B. COSTA • School of Biological Sciences, Institute for Bioengineering and Bioscience, Georgia Institute of Technology, Atlanta, GA, USA A˚SA EHLE´N • Institut Curie, PSL Research University, CNRS, UMR3348, F-91405, Orsay, France; University Paris Sud, Paris-Saclay University, CNRS, UMR3348, F-91405, Orsay, France RAJULA ELANGO • Department of Medicine, Division of Hematology-Oncology, Cancer Research Institute, Harvard Medical School, Boston, MA, USA; Beth Israel Deaconess Medical Center, Harvard Medical School, Boston, MA, USA ANASTASIYA EPSHTEIN • Department of Biochemistry and Molecular Pharmacology, New York University School of Medicine, New York, NY, USA DIONNA GAMBLE • Department of Molecular Genetics and Microbiology, Duke University, Durham, NC, USA; University Program in Genetics and Genomics, Duke University, Durham, NC, USA TATIANA GARCI´A-MUSE • Centro Andaluz de Biologı´a Molecular y Medicina RegenerativaCABIMER, Universidad de Sevilla-CSIC-UPO, Seville, Spain VINCENT GE´LI • Marseille Cancer Research Center (CRCM), U1068 Inserm, UMR7258 CNRS, Institut Paoli-Calmettes, Aix Marseille University, Marseille, France SIMON GEMBLE • Institut Curie, PSL Research University, CNRS UMR144, Paris, France ` degli Studi di ELISA GOBBINI • Dipartimento di Biotecnologie e Bioscienze, Universita Milano-Bicocca, Milano, Italia ROMA´N GONZA´LEZ-PRIETO • Department of Cell and Chemical Biology, Leiden University Medical Center, Leiden, The Netherlands JAMES E. HABER • Department of Biology, Rosenstiel Basic Medical Sciences Research Center, Brandeis University, Waltham, MA, USA LYDIA R. HEASLEY • Department of Environmental and Radiological Health Sciences, Colorado State University, Fort Collins, CO, USA WOLF-DIETRICH HEYER • Department of Microbiology and Molecular Genetics, University of California, Davis, CA, USA; Department of Molecular and Cellular Biology, University of California, Davis, CA, USA YEE FANG HUM • Department of Cancer Biology, Abramson Family Cancer Research Institute Basser Center for BRCA, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA, USA NEIL HUNTER • Howard Hughes Medical Institute, Department of Microbiology and Molecular Genetics, Davis, University of California, Davis, CA, USA GRZEGORZ IRA • Department of Molecular and Human Genetics, Baylor College of Medicine, Houston, TX, USA MASARU ITO • Howard Hughes Medical Institute, Department of Microbiology and Molecular Genetics, Davis, University of California, Davis, CA, USA MARIA JASIN • Department of Pathology, Memorial Sloan Kettering Cancer Center, New York, NY, USA RYAN B. JENSEN • Department of Therapeutic Radiology, Yale University School of Medicine, New Haven, CT, USA

Contributors

xiii

SUE JINKS-ROBERTSON • Department of Cancer Biology, Abramson Family Cancer Research Institute Basser Center for BRCA, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA, USA SAM JOHN • Laboratory of Genome Integrity, National Cancer Institute, NIH, Bethesda, MD, USA FRAULIN JOSEPH • Department of Genetics and Development, Columbia University Irving Medical Center, New York, NY, USA HANNAH L. KLEIN • Department of Biochemistry and Molecular Pharmacology, New York University School of Medicine, New York, NY, USA TAKEHIKO KOBAYASHI • Laboratory of Genome Regeneration, Institute for Quantitative Biosciences, University of Tokyo, Tokyo, Japan JURAJ KRAMARA • Department of Biology, University of Iowa, Iowa City, IA, USA KAROL KRAMARZ • Institut Curie, CNRS, UMR3348, PSL Research University, Orsay, France; Institut Curie, Paris-Saclay University, Unite Mixte de Recherche, Centre National de la Recherche Scientifique, Orsay, France; Genome Integrity, RNA and Cancer Unit (UMR3348), Equipe Labelise´e Ligue, Institut Curie—Research Center, Orsay, France LUMIR KREJCI • Department of Biology, Masaryk University, Brno, Czech Republic; International Clinical Research Center, St. Anne’s University Hospital, Brno, Czech Republic; National Center for Biomolecular Research, Masaryk University, Brno, Czech Republic SANDEEP KUMAR • Department of Molecular and Human Genetics, Baylor College of Medicine, Houston, TX, USA YOUNGHO KWON • Department of Biochemistry and Structural Biology, University of Texas Health Science Center at San Antonio, San Antonio, TX, USA SUDIPTA LAHIRI • Department of Therapeutic Radiology, Yale University School of Medicine, New Haven, CT, USA TOMAS LAMA-DIAZ • Departamento de Bioquı´mica e Bioloxı´a Molecular, CIMUS, Universidade de Santiago de Compostela-Instituto de Investigacion Sanitaria (IDIS), Santiago de Compostela, Spain SARAH A. E. LAMBERT • Institut Curie, CNRS, UMR3348, PSL Research University, Orsay, France; Institut Curie, Paris-Saclay University, Unite Mixte de Recherche, Centre National de la Recherche Scientifique, Orsay, France; Genome Integrity, RNA and Cancer Unit (UMR3348), Equipe Labelise´e Ligue, Institut Curie—Research Center, Orsay, France SO JUNG LEE • Department of Genetics and Development, Columbia University Irving Medical Center, New York, NY, USA GAE¨LLE LEGUBE • LBCMCP, Centre de Biologie Integrative (CBI), CNRS, Universite´ de Toulouse, UT3, Toulouse, France CARL P. LEHMANN • Centro de Biologı´a Molecular Severo Ochoa (CSIC/UAM), Madrid, Spain EMILY LIN • Molecular and Computational Biology Department, University of Southern California, Los Angeles, CA, USA MICHAEL LISBY • Department of Biology, University of Copenhagen, Copenhagen, Denmark KIRILL S. LOBACHEV • School of Biological Sciences, Institute for Bioengineering and Bioscience, Georgia Institute of Technology, Atlanta, GA, USA ` degli Studi di MARIA PIA LONGHESE • Dipartimento di Biotecnologie e Bioscienze, Universita Milano-Bicocca, Milano, Italia

xiv

Contributors

ANNA MALKOVA • Department of Biology, University of Iowa, Iowa City, IA, USA ALEXANDER V. MAZIN • Department of Biochemistry and Molecular Biology, Drexel University College of Medicine, Philadelphia, PA, USA OLGA M. MAZINA • Department of Biochemistry and Molecular Biology, Drexel University College of Medicine, Philadelphia, PA, USA IOANNA MITRENTSI • Institut de Ge´ne´tique et de Biologie Mole´culaire et Celullaire, INSERM U964, CNRS, UMR7104, Illkirch, France; Universite´ de Strasbourg, Strasbourg, France UFFE HASBRO MORTENSEN • Department of Biotechnology and Biomedicine, Technical University of Denmark, Kongens Lyngby, Denmark ANDRE´ NUSSENZWEIG • Laboratory of Genome Integrity, National Cancer Institute, NIH, Bethesda, MD, USA BETH OSIA • Department of Biology, University of Iowa, Iowa City, IA, USA TANYA T. PAULL • The Department of Molecular Biosciences, The University of Texas at Austin, Austin, TX, USA AURE`LE PIAZZA • Spatial Regulation of Genomes, Institut Pasteur, UMR3525 CNRS, Paris, France; Department of Microbiology and Molecular Genetics, University of California, Davis, CA, USA; Univ Lyon, ENS, UCBL, CNRS, INSERM, Laboratory of Biology and Modelling of the Cell, UMR5239, Lyon, France FE´LIX PRADO • Department of Genome Biology, Andalusian Molecular Biology and Regenerative Medicine Center (CABIMER), CSIC-University of Seville-UPO, Seville, Spain ROHIT PRAKASH • Developmental Biology Program, Memorial Sloan Kettering Cancer Center, New York, NY, USA PALLAVI RAJPUT • Department of Microbiology and Molecular Genetics, University of California, Davis, CA, USA STEVEN A. ROBERTS • School of Molecular Biosciences, Center for Reproductive Biology, Washington State University, Pullman, WA, USA VINCENT ROCHER • LBCMCP, Centre de Biologie Integrative (CBI), CNRS, Universite´ de Toulouse, UT3, Toulouse, France ELI ROTHENBERG • Department of Biochemistry and Molecular Pharmacology, Perlmutter Cancer Center, New York University School of Medicine, New York, NY, USA RODNEY ROTHSTEIN • Department of Genetics and Development, Columbia University Irving Medical Center, New York, NY, USA NADIA M. V. SAMPAIO • Department of Environmental and Radiological Health Sciences, Colorado State University, Fort Collins, CO, USA; Cell and Molecular Biology Graduate Program, Colorado State University, Fort Collins, CO, USA AURORE SANCHEZ • Institut Curie—Research Center, UMR3244 CNRS, Pavillon Trouillet Rossignol, PSL Research University, Paris Cedex 05, France; Paris Sorbonne Universite´, Paris, France SUMIT SANDHU • Howard Hughes Medical Institute, Department of Microbiology and Molecular Genetics, Davis, University of California, Davis, CA, USA MARIKO SASAKI • Laboratory of Genome Regeneration, Institute for Quantitative Biosciences, University of Tokyo, Tokyo, Japan IRENE SAUGAR • Centro de Biologı´a Molecular Severo Ochoa (CSIC/UAM), Madrid, Spain RALPH SCULLY • Department of Medicine, Division of Hematology-Oncology, Cancer Research Institute, Harvard Medical School, Boston, MA, USA; Beth Israel Deaconess Medical Center, Harvard Medical School, Boston, MA, USA

Contributors

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COLBY SEE • Molecular and Computational Biology Department, University of Southern California, Los Angeles, CA, USA MARIE-NOELLE SIMON • Marseille Cancer Research Center (CRCM), U1068 Inserm, UMR7258 CNRS, Institut Paoli-Calmettes, Aix Marseille University, Marseille, France EVI SOUTOGLOU • Institut de Ge´ne´tique et de Biologie Mole´culaire et Celullaire, INSERM U964, CNRS, UMR7104, Illkirch, France; Universite´ de Strasbourg, Strasbourg, France; Genome Damage and Stability Center, School of Life Sciences, University of Sussex, Brighton, UK TOMAS STRUCKO • Department of Biotechnology and Biomedicine, Technical University of Denmark, Kongens Lyngby, Denmark PATRICK SUNG • Department of Biochemistry and Structural Biology, University of Texas Health Science Center at San Antonio, San Antonio, TX, USA LORRAINE S. SYMINGTON • Department of Microbiology and Immunology, Columbia University Irving Medical Center, New York, NY, USA JOSE´ ANTONIO TERCERO • Centro de Biologı´a Molecular Severo Ochoa (CSIC/UAM), Madrid, Spain EMANUELA TUMINI • Centro Andaluz de Biologı´a Molecular y Medicina Regenerativa, CABIMER, Universidad de Sevilla-CSIC-UPO, Seville, Spain FABIO VANOLI • Developmental Biology Program, Memorial Sloan Kettering Cancer Center, New York, NY, USA DOMAGOJ VUGIC • Institut Curie, PSL Research University, CNRS, UMR3348, F-91405, Orsay, France; University Paris Sud, Paris-Saclay University, CNRS, UMR3348, F-91405, Orsay, France DONNA R. WHELAN • Department of Pharmacy and Biomedical Sciences, La Trobe Institute for Molecular Science, La Trobe University, Bundoora, VIC, Australia; Department of Biochemistry and Molecular Pharmacology, Perlmutter Cancer Center, New York University School of Medicine, New York, NY, USA TRAVIS WHITE • Department of Pathology, Memorial Sloan Kettering Cancer Center, New York, NY, USA NICHOLAS A. WILLIS • Department of Medicine, Division of Hematology-Oncology, Cancer Research Institute, Harvard Medical School, Boston, MA, USA; Beth Israel Deaconess Medical Center, Harvard Medical School, Boston, MA, USA NANCY WONG • Laboratory of Genome Integrity, National Cancer Institute, NIH, Bethesda, MD, USA MIYUKI YAMAGUCHI • Department of Biology, Rosenstiel Basic Medical Sciences Research Center, Brandeis University, Waltham, MA, USA ZHENXIN YAN • Department of Molecular and Human Genetics, Baylor College of Medicine, Houston, TX, USA YAN YUN • Howard Hughes Medical Institute, Department of Microbiology and Molecular Genetics, Davis, University of California, Davis, CA, USA YI ZHOU • The Department of Molecular Biosciences, The University of Texas at Austin, Austin, TX, USA

Chapter 1 Detection of DNA Double-Strand Breaks by γ-H2AX Immunodetection Sonia I. Barroso and Andre´s Aguilera Abstract DNA double-strand breaks (DSBs) are the most deleterious type of DNA damage and a cause of genetic instability as they can lead to mutations, genome rearrangements, or loss of genetic material when not properly repaired. Eukaryotes from budding yeast to mammalian cells respond to the formation of DSBs with the immediate phosphorylation of a histone H2A isoform. The modified histone, phosphorylated in serine 139 in mammals (S129 in yeast), is named γ-H2AX. Detection of DSBs is of high relevance in research on DNA repair, aging, tumorigenesis, and cancer drug development, given the tight association of DSBs with different diseases and its potential to kill cells. DSB levels can be obtained by measuring levels of γ-H2AX in extracts of cell populations or by counting foci in individual nuclei. In this chapter some techniques to detect γ-H2AX are described. Key words DNA damage, Double-strand breaks, γ-H2AX, Immunofluorescence, Immunoblotting

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Introduction DNA can be damaged by endogenous and exogenous sources causing, among other lesions, double-strand breaks (DSBs). These can cause from cell death, if unrepaired, to loss of genetic material, if mis-repaired. Mammalian cells respond to agents that introduce DSBs with the phosphorylation of histone H2AX [1]. After the exposure to a damage-inducing agent, such as ionizing radiation, thousands of γ-H2AX molecules accumulate per DSB [2]. Given the high amplification of the signal, every DSB can be detected as a focus of γ-H2AX by immunological techniques using specific antibodies. These foci are crucial to the proper recruitment of repair factors to the site of damage [3]. The H2AX C-terminal tail that has the phosphorylated motif in mammals is also present in histone H2A from yeast, and it is also phosphorylated after DNA damage to facilitate DNA repair [4]. There are several detection techniques to evaluate DNA lesions such as DSBs: single-cell electrophoresis (Comet assay), terminal

Andre´s Aguilera and Aura Carreira (eds.), Homologous Recombination: Methods and Protocols, Methods in Molecular Biology, vol. 2153, https://doi.org/10.1007/978-1-0716-0644-5_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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deoxynucleotidyl transferase dUTP nick-end labeling (TUNEL), linear amplification-mediated high-throughput genome-wide translocation sequencing (LAM-HTGTS), DNA breakage detection-fluorescence in situ hybridization (DBD-FISH), and detection of DSB-associated molecular markers such as the Ku protein or γ-H2AX. These techniques have been thoroughly reviewed [5–11] and in this chapter we focus on the detection of phosphorylated H2AX as a measurement of DSBs because it is one of the earliest events after DSB formation and it is easily immunodetected. Given that DSBs can lead to genetic instability and eventually to cancer, and that paradoxically DSB induction is also used as cancer treatment, it is very useful to have a tool to monitor disease progression and/or treatment effectiveness based on the accumulation of γ-H2AX [12]. It is also useful for the understanding of the DNA damage response, repair, and all processes that control genomic integrity when factors of such pathways are missing or do not work properly [13, 14].

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Materials

2.1 γ-H2AX Immunofluorescence in Mammalian Cells

1. DMEM-Dulbecco’s Modified Eagle Medium. 2. 24-well plates. 3. Tweezers. 4. Round coverslips. 5. Bovine serum albumin (BSA). 6. PBS tablets. 7. Blocking solution: 3% BSA in phosphate-buffered saline (PBS). 8. anti-γ-H2AX antibody (Merck-Millipore). 9. Formaldehyde (methanol free) 10% ultra pure. 10. Alexa Fluor 594 goat anti-mouse. 11. Vacuum line. 12. DAPI solution (1 μg/ml in PBS). 13. Immu-Mount Mounting Medium. 14. Microscope slides. 15. Fluorescence microscope.

2.2 Detection of γ-H2AX in Mammalian Cells by Western Blotting

1. Lysis buffer (RIPA): 0.3 M NaCl, 1% (v/v) NP-40, 5% (w/v) sodium deoxycholate, 0.5% (w/v) SDS, 50 mM Tris–HCl pH 8.0. 2. Laemmli buffer 2: 4% SDS, 20% glycerol (v/v), 2% 2-mercaptoethanol, 0.004% (w/v) bromophenol blue, 125 mM Tris–HCl pH 6.8.

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3. Difco Skim Milk. 4. PBS tablets. 5. Protein ladder. 6. Running buffer: 25 mM Tris, 192 mM glycine. 7. Nitrocellulose blotting membrane with a pore size of 0.2 μm. 8. Transfer buffer: 25 mM Tris, 192 mM glycine, 20% (v/v) methanol. 9. TBS-T: 20 mM Tris–HCl pH 7.5, 150 mM NaCl, 0.1% Tween 20. 10. Blocking solution: 5% (w/v) Difco Skim Milk in TBS-T. 11. anti-γ-H2AX antibody (Merck-Millipore). 12. Goat anti-mouse HRP. 13. Supersignal West Pico PLUS Chemiluminescent Substrate. 14. High-sensitivity chemiluminescent films. 15. Phosphatase inhibitors. 16. Mini-PROTEAN Tetra Cell. 2.3 Detection of γ-H2AX in Yeast by Western Blotting

1. Trichloroacetic acid (TCA). 2. Difco Skim Milk. 3. Phosphatase inhibitors. 4. Laemmli buffer 2: 4% SDS, 20% glycerol (v/v), 2% 2-mercaptoethanol, 0.004% (w/v) bromophenol blue, 125 mM Tris–HCl pH 6.8. 5. Glass beads acid-washed. 6. Multi-vortex mixer. 7. Running buffer: 25 mM Tris, 192 mM glycine. 8. Transfer buffer: 25 mM Tris, 192 mM glycine, 20% (v/v) methanol. 9. Nitrocellulose blotting membrane with a pore size of 0.2 μm. 10. Blocking solution: 5% (w/v) Difco Skim Milk in TBS-T. 11. TBS-T: 20 mM Tris–HCl pH 7.5, 150 mM NaCl, 0.1% Tween 20. 12. Anti-histone H2A (phospho S129) antibody. 13. Horseradish peroxidase conjugated goat anti-rabbit antibody. 14. Supersignal West Pico PLUS Chemiluminescent Substrate. 15. High-sensitivity chemiluminescent films. 16. Protein ladder. 17. Tris 1 M. 18. Mini-PROTEAN Tetra cell.

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Methods

3.1 γ-H2AX Immunofluorescence in Mammalian Cells

1. HeLa cells are cultured on coverslips (see Note 1) at a concentration of 2  105 cells/well in a 6-well plate. 2. After 24 h, coverslips are transferred to a 24-well plate with cold PBS (see Note 2). 3. Aspirate the PBS with the vacuum system and add 1 ml of 2% formaldehyde in PBS (see Note 3). Incubate for 20 min at RT. 4. Remove formaldehyde solution (see Note 3). 5. Incubate the cells for 5 min at

20  C in 70% ethanol.

6. Replace this solution with fresh 70% ethanol. Incubate 5 min at 4  C (see Note 4). 7. Incubate the cells with blocking solution (1 ml/well) and incubate 1 h at room temperature. 8. Remove blocking solution and add anti-γ-H2AX (1:1000 in blocking solution) antibody in a total volume of 250 μl/well. Incubate 1 h at room temperature. 9. Wash twice in PBS (5 min each) and incubate with Alexa Fluor 594 goat anti-mouse secondary antibody (1:1000 in blocking solution) 1 h at room temperature. 10. Wash twice for 5 min each in PBS. Incubate with DAPI solution for 5 min at room temperature and wash twice more for 5 min each in PBS. Wash once in distilled H2O. 11. Coverslips are removed using tweezers and placed on a microscope slide with Immu-Mount mounting medium (see Note 5). 12. The slides are placed flat at room temperature for 24 h and then stored at 4  C (see Fig. 1). 3.2 Detection of γ-H2AX in Mammalian Cells by Western Blotting

1. Collect the cells and wash with ice-cold PBS. 2. Centrifuge at 13,000  g 5 min. Discard the supernatant. 3. Add 100 μl of RIPA buffer with phosphatase inhibitors to 106 cells and incubate 30 min on ice. 4. Centrifuge 13,000  g 10 min at 4  C. Transfer the supernatant to a new tube. 5. Add an equal volume of Laemmli buffer 2 to a volume of cell extract. Boil samples at 95  C for 5 min. Centrifuge at 13,000  g 5 min (see Note 6). 6. Load 15–20 μl of each sample in a 12% acrylamide gel. Include a protein ladder. Run the electrophoresis at 200 V 45 min. 7. Transfer the acrylamide gel to a nitrocellulose membrane at 400 mA for 2 h (see Note 7).

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Fig. 1 Representative image of HeLa cells stained with DAPI (DNA) and γ-H2AX antibody. γ-H2AX foci can be observed in the nuclei

Fig. 2 Representative image of γ-H2AX immunoblotting. An increase in γ-H2AX is observed after DSB induction (+). The lower band corresponds to the loading control

8. Incubate the membrane in blocking solution. 9. Wash the membrane with TBS-T and add anti-γ-H2AX antibody (1:1000) in blocking solution. Incubate overnight at 4  C with gentle shaking. 10. Wash twice with TBS-T. Add secondary antibody (1:10000) in blocking solution. Incubate 1 h at room temperature with gentle shaking. 11. Wash three times with TBS-T, 5 min each. 12. Prepare the detection reagent by mixing equal volumes of each solution. 13. Place the membrane face-up on a parafilm-coated plate and add the detection reagent. Incubate 5 min at room temperature in the dark. 14. Acquire the image using development techniques for chemiluminescence (see Fig. 2). 3.3 Detection of γ-H2AX in Yeast by Western Blotting

1. Grow 20 ml yeast cultures in the appropriate medium and temperature to an absorbance at 600 nm of about 0.8. 2. Harvest cells by centrifugation at 4000  g 3 min. Discard supernatant. 3. Wash the cells with 1 ml of cold 20% TCA (w/v in H2O) (see Note 8).

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4. Spin down in a microcentrifuge tube for 75 s at 12,000  g. Discard supernatant (see Note 9). 5. Resuspend the cell pellet in 200 μl of 10% TCA and add 200 μl of glass beads (see Note 8). 6. Vortex at high speed 10 min at 4  C in a multi-vortex mixer. Spin down for 75 s at 12,000  g. 7. Transfer supernatant to a new tube and wash the beads twice with 200 μl 10% TCA (see Note 8). Transfer the supernatant to the same tube (600 μl final volume). 8. Centrifuge 10 min at 1000  g at room temperature. Discard supernatant. 9. Add 100 μl of Laemmli buffer 2 plus 50 μl of H2O to the cell pellet (see Note 6). Resuspend by vortex and add 50 μl of Tris 1 M to neutralize (see Note 10). Mix by vortex. 10. Boil samples at 95  C for 10 min. Mix by vortex. 11. Centrifuge at 1000  g 10 min at room temperature. 12. Load 15–20 μl of each sample in a 12% acrylamide gel. Include a protein ladder. Run the electrophoresis at 200 V 45 min. 13. Transfer the acrylamide gel to a nitrocellulose membrane at 30 V overnight at 4  C (see Note 7). 14. Incubate the membrane in blocking solution. 15. Wash the membrane with TBS-T and add anti-histone H2A (phospho S129) antibody (1:2000) in blocking solution. Incubate overnight at 4  C with gentle shaking. 16. Wash twice with TBS-T. Add secondary antibody (1:2000) in blocking solution. Incubate 1 h at room temperature with gentle shaking. 17. Wash three times with TBS-T, 5 min each. 18. Prepare the detection reagent by mixing equal volumes of each solution. 19. Place the membrane face-up on a parafilm-coated plate and add the detection reagent. Incubate 5 min at room temperature in the dark. 20. Acquire the image using development techniques for chemiluminescence.

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Notes 1. Use tweezers to place two to three round coverslips on a well before seeding the cells. 2. Fill the 24-well plate with 1 ml of PBS per well and place one coverslip per well with the cells on the upper side.

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3. Formaldehyde is a sensitizing agent and a cancer hazard. Wear gloves and a lab coat and always work in a chemical fume hood. 4. In this step, coverslips can be kept in 70% ethanol at 4  C for several weeks. 5. Place a drop of about 30 μl of mounting medium on a slide and place the coverslip with the cells facing the mounting medium. 6. 2-mercaptoethanol is toxic if inhaled and may cause damage to organs. Wear gloves and a lab coat and always work in a chemical fume hood. 7. Methanol is a hazardous chemical with significant toxic, flammable, and reactive properties that can produce deleterious impacts on human health and the environment when not properly handled. Wear a lab coat and gloves when working with methanol and always manipulate it in a chemical fume hood. 8. Trichloroacetic acid can cause skin burns and eye damage. Wear gloves and a lab coat when working with this reagent. 9. Pellets can be kept frozen at

20  C.

10. Add Tris 1 M till the solution becomes blue.

Acknowledgments AA’s lab is funded by the Spanish Ministry of Economy and Competitiveness, Junta de Andalucı´a, European Research Council, and the European Union (FEDER). References 1. Rogakou EP, Pilch DR, Orr AH, Ivanova VS, Bonner WM (1998) DNA double-strand breaks induce histone H2AX phosphorylation on serine 139. J Biol Chem 273 (10):5858–5868 2. Rogakou EP, Boon C, Redon C, Bonner WM (1999) Megabase chromatin domains involved in DNA double-strand breaks in vivo. J Cell Biol 146(5):905–916 3. Paull TT, Rogakou EP, Yamazaki V, Kirchgessner CU, Gellert M, Bonner WM (2000) A critical role for histone H2AX in recruitment of repair factors to nuclear foci after DNA damage. Curr Biol 10(15):886–895 4. Downs JA, Lowndes NF, Jackson SP (2000) A role for Saccharomyces cerevisiae histone H2A in DNA repair. Nature 408(6815):1001–1004 5. Figueroa-Gonza´lez G, Pe´rez-Plasencia C (2017) Strategies for the evaluation of DNA damage and repair mechanisms in cancer.

Oncol Lett 13(6):3982–3988. https://doi. org/10.3892/ol.2017.6002 6. Ostling O, Johanson KJ (1984) Microelectrophoretic study of radiation-induced DNA damages in individual mammalian cells. Biochem Biophys Res Commun 123(1):291–298 7. Anderson D, Dhawan A, Laubenthal J (2013) The comet assay in human biomonitoring. Methods Mol Biol 1044:347–362. https:// doi.org/10.1007/978-1-62703-529-3_18 8. Kumari S, Rastogi R, Singh K, Singh S, Sinha R (2008) DNA damage: detection strategies. EXCLI J 7:44–62 9. Hu J, Meyers RM, Dong J, Panchakshari RA, Alt FW, Frock RL (2016) Detecting DNA double-stranded breaks in mammalian genomes by linear amplification-mediated highthroughput genome-wide translocation sequencing. Nat Protoc 11(5):853–871. https://doi.org/10.1038/nprot.2016.043

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10. Levsky JM, Singer RH (2003) Fluorescence in situ hybridization: past, present and future. J Cell Sci 116(Pt 14):2833–2838 11. Jones JM, Gellert M, Yang W (2001) A Ku bridge over broken DNA. Structure 9 (10):881–884 12. Bonner WM, Redon CE, Dickey JS, Nakamura AJ, Sedelnikova OA, Sollier S, Pommier Y (2008) Gamma H2AX and cancer. Nat Rev Cancer 8:957–967 13. Domı´nguez-Sa´nchez M, Barroso S, Go´mezGonza´lez B, Luna R, Aguilera A (2011)

Genome instability and transcription elongation impairment in human cells depleted of THO/TREX. PLoS Genet 7(12):e1002386. https://doi.org/10.1371/journal.pgen. 1002386 14. Bhatia V, Barroso SI, Garcı´a-Rubio ML, Tumini E, Herrera-Moyano E, Aguilera A (2014) BRCA2 prevents R-loop accumulation and associates with TREX-2 mRNA export factor PCID2. Nature 511(7509):362–365. https://doi.org/10.1038/nature13374

Chapter 2 END-seq: An Unbiased, High-Resolution, and Genome-Wide Approach to Map DNA Double-Strand Breaks and Resection in Human Cells Nancy Wong, Sam John, Andre´ Nussenzweig, and Andre´s Canela Abstract DNA double-strand breaks (DSBs) represent the most toxic form of DNA damage and can arise in either physiological or pathological conditions. If left unrepaired, these DSBs can lead to genome instability which serves as a major driver to tumorigenesis and other pathologies. Consequently, localizing DSBs and understanding the dynamics of break formation and the repair process are of great interest for dissecting underlying mechanisms and in the development of targeted therapies. Here, we describe END-seq, a highly sensitive next-generation sequencing technique for quantitatively mapping DNA double-strand breaks (DSB) at nucleotide resolution across the genome in an unbiased manner. END-seq is based on the direct ligation of a sequencing adapter to the ends of DSBs and provides information about DNA processing (end resection) at DSBs, a critical determinant in the selection of repair pathways. The absence of cell fixation and the use of agarose for embedding cells and exonucleases for blunting the ends of DSBs are key advances that contribute to the technique’s increased sensitivity and robustness over previously established methods. Overall, END-seq has provided a major technical advance for mapping DSBs and has also helped inform the biology of complex biological processes including genome organization, replication fork collapse and chromosome fragility, off-target identification of RAG recombinase and gene-editing nucleases, and DNA end resection at sites of DSBs. Key words DNA double-strand breaks (DSBs), DNA damage, DNA repair, DSB mapping and quantification, Next-generation sequencing, Nucleotide resolution, End resection, Adapter ligation, Exonucleases, Agarose plugs

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Introduction Central to a cells ability to maintain genome stability are systems that monitor and repair DSBs. DSBs have been shown to occur in response to exogenous insults such as exposure to irradiation, chemotherapeutic and gene-editing agents, and from physiological processes such as replication, transcription, meiotic recombination in germ cells, and antigen receptor rearrangements in lymphocytes. If not rapidly and faithfully repaired, DSBs can serve as substrates

Andre´s Aguilera and Aura Carreira (eds.), Homologous Recombination: Methods and Protocols, Methods in Molecular Biology, vol. 2153, https://doi.org/10.1007/978-1-0716-0644-5_2, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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for aberrant chromosomal rearrangements, which, in turn, can promote cancer and other pathologies. 1.1 Current and Historical Approaches to Map DSBs

A major limitation of current cytological and genomic approaches that assess DNA damage is their inability to map the chromosomal locations of DSBs and the resulting structures at breaks with precision and detail. Identifying the exact location of DSBs in the genome will greatly expand our understanding of the mechanism of their formation and will additionally allow for the monitoring of the kinetics of DNA processing and repair at DSBs. Precise DSB mapping will also allow for the integration of genome context (i.e., DNA composition, chromatin modifications, and processes like transcription and replication) in assessing their influence on the formation and repair of DSBs. In recent years, the need for mapping and quantifying on- and off-target specificities of DSB-generating gene-editing nucleases in combination with the advances in next-generation sequencing has helped accelerate the development of genome-wide techniques to map DSBs. Many of these approaches are based on indirect readouts of DSB formation such as chromatin immunoprecipitation of DNA repair proteins (e.g., H2AX, SMC5, RPA, and other proteins [1–4]) or by detecting the outcome of improper DSB repair such as chromosomal translocations [5] or via the incorporation of an exogenously provided DNA repair sequence [6, 7]. Collectively, these outputs serve as surrogate marks for the site of a DSB. Each of these indirect approaches is not only unable to map the exact position of DSBs directly with sensitivity and robustness but also fails to provide features regarding the structure and processing of damaged DNA ends. These methodological limitations have prompted us to develop END-seq—a genome-wide, high-resolution, direct and quantitative readout for measuring DSBs in an unbiased manner that provides technological resolution to each of these major inadequacies. BLESS, and its recent modification BLISS, also map broken DNA ends in an unbiased genome-wide manner [8, 9]. These techniques are, however, associated with additional noise, primarily caused by the use of formaldehyde as a fixative. Importantly, neither BLESS/BLISS provides information about end structures at DSBs. END-seq, therefore, provides a major advance in the detection of double-strand breaks.

1.2 Overview of the END-seq Procedure

Briefly, in END-seq, native and unfixed human or mouse cells are embedded in agarose (Fig. 1). This critical step avoids the artificial generation of DSBs from mechanical processing and fixation of samples. After protein and RNA digestion, the DNA ends of DSBs are blunted with single-strand DNA-specific exonucleases, A-tailed (i.e., one “A” is added in a nontemplated manner to each blunted DNA end), and ligated within agarose plugs to a T-tailed DNA adapter which is labeled with biotin and compatible with

High-Resolution Mapping of DSBs and Resection Cells embedded in agarose plugs

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Proteinase K and RNAseA digestion Blunting of DSB ends with exonuclease VII and/or exonuclease T

DSB

5’

3’ 5’

3’

Streptavidin capture, end-repair & A-tailing

Ligate END-seq adapter 2 B

A

B Hairpin digestion

A-tailing of blunted ends

Ligate END-seq adapter 1

A

B B

Melt agarose plug & shear DNA B

B

B

B

Illumina PCR library amplification Illumina sequencing single-end

Barcode 1

Barcode 2

Fig. 1 Detailed schema of the END-seq methodology. Live cells are embedded in low-melting agarose following which agarose plugs are treated with proteinase K and RNase A to digest proteins and RNA, respectively. After blunting the single-strand DNA overhangs at DSBs with exonucleases (exonuclease VII and exonuclease T), A-tailing of the 30 ends allows the ligation of a biotinylated hairpin adapter containing a 30 T overhang and the Illumina’s p5 adapter sequence (“END-seq adapter 1”) within the plug. The agarose plug is then melted and DNA is extracted and sheared by sonication. Sheared fragments containing END-seq adapter 1 (and the DSB end) are captured with streptavidin-coated beads. The new ends created by sonication are end repaired and A-tailed, allowing ligation of a second hairpin adaptor containing Illumina’s p7 sequence (named “END-seq adapter 2”). Adapter hairpins are digested away with the USER enzyme (a combination of an uracil DNA-glycosylase and endonuclease VIII) which digests DNA to create single-nucleotide gaps at uracil residues. PCR amplification using Illumina TruSeq primers with barcodes (denoted in yellow and green) result in a ready-to-use library in which the first base sequenced (read number 1) corresponds to the first base of the blunted DSB. (Figure has been adapted from [11])

Illumina sequencing. Next, high-molecular-weight DNA is extracted from the agarose plugs and fragmented by sonication. Subsequent purification of the biotinylated ends and ligation of a second adapter followed by PCR amplification result in a ready-tosequence library in which the first base sequenced corresponds precisely to the first base of the blunted DSB. Furthermore, the number of reads mapping to the DSB is proportional to the frequency of DSBs in the cell population, which allows for the quantification of each DSB. One of the initial steps of DSB repair by homologous recombination (HR) is the generation of 30 single-stranded DNA (ssDNA) via the 50 to 30 nucleolytic degradation of broken ends—a process termed DNA end resection [10]. DNA end resection inhibits an error-prone repair pathway called nonhomologous end joining and initiates repair using the high-fidelity homologous recombination (HR) pathway by recruiting the RAD51 recombinase that promotes strand invasion to complete HR. Since the blunting step in the END-seq protocol removes single-stranded overhangs and sequencing initiates at the first nucleotide in blunted doublestranded DNA, the resected interval can be calculated as the distance from the sequencing start site to the original position of the

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WT

Lig4-/-

Lig4-/-53bp1-/-

Fig. 2 Screen shot: a visual example of resection and DNA processing at a single DSB. A genome browser screen shot of one DSB in the murine genome located at chr3: chr3:9,004,213-9,004,220. The site of the break is highlighted with a vertical blue bar above the top track. The y-axis represents the total number of accumulated reads at the AsiSI site depicted in WT, Lig4/ and Lig4/53bp1/ pre-B-cell lines. Lig4/ are repair-deficient cells that allow for DSB persistence and resection while the combined ablation of LIG4 and the pro-NHEJ factor 53BP1 results in hyperresection. The accumulation of reads away from the DSB indicates end resection. Read coverage is colored in black or red for plus and minus strand alignments, respectively. Horizontal blue bars at the bottom of each track represent resected regions in the vicinity of the DSB

DSB. Given that each cell undergoes resection to differing extents, END-seq reveals a resection distribution profile of ends from the original site of the DSB (i.e., non resected) to several kilobases on each side of the break (i.e., resected) (Fig. 2). 1.3 Normalization Using Spike-Ins

Normalization of END-seq data is crucial to accurately compare the frequency of DSBs between samples. Although the same number of cells is used in all samples, normalization of DSBs by the total number of sequenced reads can lead to misleading interpretations. Simple read counts are unable to correct global changes in DSBs caused by specific experimental conditions (e.g., drug treatments, genotypes, time points) or even noise from apoptotic cells. Accurate comparison between samples, therefore, requires the use of an exogenous spike-in control. The spike-in control, used in the

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END-seq procedure, are cells where a single, unique DSB can be induced by a zinc-finger nuclease in a repair-deficient cell line. This site has been shown to be broken in 100% of induced spike-in cells. Consequently, any differences in the total amount of reads at the spike-in DSB between samples can be used as a normalization control. In addition, the spike-in normalization allows for the estimation of the frequency of cells harboring any given DSB across the entire genome. For a 5% spike-in, the read counts at the unique zinc finger nuclease–induced DSB signifies a frequency of 5% of cells carrying a DSB in the total cellular population. Spike-in normalization, therefore, allows for the calculation of break frequency and enables quantitative comparisons between different samples in the experiment. 1.4 Applications and Uses of END-seq

END-seq has been successfully applied to detect and quantify DSBs produced by restriction endonucleases in vivo, genome editing nucleases, recombination-activating genes (RAGs) during VDJ recombination [11], activation-induced cytidine deaminase (AID) during class switch recombination, topoisomerase-2 induced breaks [12, 13], and at collapsed replication forks upon replicative stress [14]. DSB hotspots and resection during meiotic recombination may also be detected. END-seq has been effectively implemented in both murine and human cells across numerous genotypes and treatment conditions and represents a tool with great versatility and flexibility.

1.5 Sensitivity, Limitations, and Caveats

The ability to detect a DSB (i.e., sensitivity) using END-seq depends primarily on two factors: (1) the frequency of DSBs and (2) the spatial distribution at any given DSB in the pool of cells in a sample. Simply put, a higher signal (read count) will be detected by END-seq if more cells in the population have a DSB (i.e., high frequency) and the breaks at a specified location are proximal to each other between individual cells. We estimate that detection of a single DSB by END-seq necessitates that a break occurs at least in 1 in every 10,000 cells [11]. If DSBs are not coincident in their position between cells, they are not detected by END-seq as they will be indistinguishable from background. In other words, a major END-seq requirement for DSB detection is that the break sites be recurrent. Random DSBs, such as those generated after irradiation, cannot be detected by END-seq and their presence in the sample is reflected in increased noise which decreases the overall sensitivity of the method. It is worth stating that DNA repair–deficient genotypes increase END-seq sensitivity because it prolongs the time DSBs stay unrepaired.

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Materials

2.1 Common Reagents and Instruments

1. Two water baths. 2. Eppendorf Thermomixer C fitted with 50 mL and 1.5 mL adapters or equivalent. 3. Wide-bore pipette filtered tips, 200 μL, VWR, Catalog # 46620-642 or equivalent. 4. Orbital platform shaker that can reach at least 180 rpm, FINEPCR shaker model SH30 or equivalent.

2.2 Embedding Cells in Agarose Plugs

1. Metal block from a dry bath or a thermoblock. 2. Puregene Proteinase K enzyme (Qiagen Catalog # 158920, 5 mL). 3. Plug mold, cell suspension buffer, and 2% agarose from Bio-Rad CHEF Mammalian Genomic DNA Plug Kit (Catalog # 170-3591). Prepare aliquots of the agarose included in the kit by melting the entire bottle in boiling water for 15 min and distributing the melted agarose into 500 μL aliquots in 1.5 mL tubes. Aliquots can be stored at 4  C. Use one or more 500 μL aliquots, as needed, for each new procedure. 4. Lysis buffer (10 mM Tris–HCl pH 8, 50 mM EDTA, 150 mM NaCl, 1% SDS in nuclease free water. The lysis buffer should be stored at room temperature to avoid SDS precipitation.)

2.3 RNAse Treatment of Plugs

1. Plug wash buffer (10 mM Tris, pH 8.0, 50 mM EDTA, in nuclease free water). 2. Bio-Rad screened cap, Catalog # 1703711. 3. Puregene RNAse A solution, 5 mL, Qiagen, Catalog # 158924. 4. TE buffer (10 mM Tris, pH 8, 1 mM EDTA, in nuclease-free water).

2.4 Blunting of DSBs Ends, A-Tailing, and Ligation of END-Seq Adapter 1

1. EB buffer (10 mM Tris, pH 8.0, in nuclease-free water). 2. NEB Exonuclease VII and 5 Exonuclease VII buffer, Catalog # M0379L, 10,000 units/mL. 3. Homemade 5 Exonuclease VII buffer (250 mM Tris–HCl (pH 8.0), 250 mM sodium phosphate (pH 8.0), 50 mM 2-mercaptoethanol, 40 mM EDTA, pH 8 at 25  C). Store in 1.8 mL aliquots at 20  C. 4. NEB Exonuclease T, Catalog # M0265L, 5000 units/mL. 5. Rotatory mixer for 1.5 mL tubes, Labnet mini-labroller rotator H5500.

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Table 1 Sequence of adapters and primers used in the END-seq protocol Adapter or primer name

Sequence 50 –30

END-seq adapter 1

Phosphate GATCGGAAGAGCGTCGTGTAGGGAAAGAGTGUU biotindTU biotin-dTUUACAC TCTTTCCCTACACGACGCTCTTCCGATC∗T

END-seq adapter 2

Phosphate GATCGGAAGAGCACACGTCUU UUUUUUAGACGTGTGCTCTTCCGATC∗T

TruSeq barcoded primer p5

AATGATACGGCGACCACCGAGATCTACACN NNNNNNNACACTCTTTCCCTACACGACGCT CTTCCGATC∗T

TruSeq barcoded primer p7

CAAGCAGAAGACGGCATACGAGANNNNN NNNGTGACTGGAGTTCAGACGTGTGCTCTTC CGATC∗T

See Note 14 for adapter annealing NNNNNNNN represents 8 nucleotide barcodes, following Illumina’s recommendations (https://support.illumina. com/downloads/index-adapters-pooling-guide-1000000041074.html). “∗” denotes a phosphothiorate bond

6. NEBuffer 4, NEB Catalog # B7004S (also used in Subheading 2.6). 7. NEB Klenow Fragment (30 ! 50 exo-), Catalog # M0212L, 5000 units/mL (also used in step 2.6). 8. NEBNext dA-Tailing Reaction Buffer, alternatively NEBuffer 2 supplemented with 0.2 mM of dATP (1) (also used in Subheading 2.6). 9. NEB Quick Ligation Kit includes Quick Ligase (2000 U/μL) and 2 Quick Ligase Buffer, Catalog # M2200L (also used in Subheading 2.6). 10. END-seq adapter 1 (see Table 1 and note indicated for adapter annealing). 11. NEBuffer 2, Catalog # B7002S (also used in Subheading 2.6). 2.5 DNA Sonication and Shearing

1. Covaris S2 or S220 focused ultrasonicator or alternatively any sonicator (e.g., Bioruptor Plus, Diagenode, Catalog # B01020001) that gives a narrow size distribution of DNA fragments around 175 bp. 2. Covaris Holder microTUBE, Catalog # 500114. 3. NEB β-Agarase I, Catalog # M0392L, 1000 units/mL. 4. Dialysis membrane filter, MF, 0.1 μm, Millipore, Catalog # VCWP04700. 5. SDS 10% Solution, Ambion, Catalog # AM9823. 6. Covaris microTUBE AFA Fiber Pre-Slit Snap-Cap 6  16 mm, Catalog # 520045. 7. Sodium acetate (3 M), pH 5.5 Ambion, Catalog # AM9740.

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8. Glycogen from mussels, 20 mg/mL, Roche, Catalog # 10901393001 or equivalent. 9. Nanodrop spectrophotometer or equivalent. 2.6 End-Repair, A-Tailing, and Ligation of END-Seq Adapter 2

1. ThermoFisher DynaMag-2 Magnet Catalog # 12321D or equivalent. 2. PCR machine, Bio-Rad T100 or equivalent. 3. END-seq adapter 2 (see Table 1 and Note indicated for adapter annealing). 4. Dynabeads MyOne Streptavidin C1 Catalog # 65002. 5. Dynabeads binding buffer 2 (10 mM Tris–HCl, 1 mM EDTA, 2 M NaCl, pH 7.5 in nuclease-free water). 6. Dynabeads wash buffer 1 (5 mM Tris–HCl, 0.5 mM EDTA, 1 M NaCl, pH 7.5 in nuclease-free water). 7. dNTP 100 mM Set Bioline Catalog # 39025 (make 10 mM mix combining dATP, dTTP, dCTP, dGTP). 8. NEB T4 ligase buffer 10, Catalog # B0202S. 9. NEB T4 Polynucleotide Kinase, Catalog # M0201L, 10,000 units/mL. 10. NEB T4 DNA Polymerase, Catalog # M0203L, 3000 units/ mL. 11. NEB DNA Polymerase I, Large (Klenow) Fragment, Catalog # M0210L, 5000 units/mL. 12. NEB USER Enzyme, Catalog # M5505L, 50 units, 1000 units/mL. 13. KAPA HiFi HotStart ReadyMix, Catalog # KK2601. 14. Illumina Truseq 8 nucleotide barcoded primer p5 (Table 1). 15. Illumina Truseq 8 nucleotide barcoded primer p7 (Table 1).

2.7 PCR Product Cleanup, Gel Purification, qPCR, and Sequencing

1. ThermoFisher DynaMag-PCR Magnet Catalog # 492025 or equivalent. 2. DNA gel electrophoresis system. 3. Standard blue LED transilluminator. 4. Agencourt AMPure XP beads, Catalog # A63880 or equivalent. 5. QIAquick Gel Extraction Kit, Catalog # 28706. 6. KAPA Library Quantification Kit for Illumina platforms. 7. Bio-Rad CFX96 Touch Real-Time PCR detection system or equivalent. 8. NextSeq 550 Series or Illumina sequencer equivalent (HiSeq) and sequencing kits, Catalog # 20024906.

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3

17

Methods Each sample of END-seq corresponds to one agarose plug. The cell size limits the number of cells that can be used to make each plug (from two million of senescent MEFs (large cells) to 80 million of mouse thymocytes (small cells)). The volume of the cell pellet cannot exceed 12 μL. It is possible to make and process more than one plug per sample to increase the total number of cells and enhance the sensitivity of the method.

3.1 Day 1: Plug Making and Proteinase K Digestion

Prepare in advance two water baths or heat blocks, at 70  C and 37  C. Melt 500 μL aliquot(s) of 2% agarose (Bio-Rad kit) in a 70  C water bath for 10 min until liquified and equilibrate at 37  C water bath for at least 10 min before use. 1. Harvest cells, count and aliquot the desired number of cells. Depending on cell type, we recommend the following number of cells per plug: for stimulated B-cells, 20 million cells; T-cells,15 million; HCT116, seven million; HEK293T, seven million; HeLa, five million and MCF7, four million. As noted above, several plugs for the same sample can be made to increase sensitivity (usually to a total of 15 million) and processed in parallel until day 3 at which point the individual plugs can be combined (see Note 1 for removing dead cells). 2. Spin cells and resuspend them in 1 mL of PBS and transfer to a 1.5 mL tube. 3. Add a fixed number of spike-in cells (2–5%) to each sample. 4. Spin down cells at 400  g for 7 min at 4  C. 5. Remove PBS from the 1.5 mL tube and leave approximately 50 μL covering the cell pellet. Slowly add 1 mL of PBS with a P1000 micropipette tip, do not disturb the pellet, and spin at 400  g for 5 min at 4  C (see Note 2). 6. Repeat step 5. 7. After the second PBS wash, remove PBS from the 1.5 mL tube and leave approximately 50 μL covering the cell pellet. Spin again for 10 s to remove any residual PBS droplets from the walls of the tube. 8. Use a 200 μL micropipette tip to remove the rest of the PBS and resuspend the pellet in 57 μL of Bio-Rad Cell Suspension Buffer. The cell pellet should occupy a volume of 5–13 μL and the mixture of cells and suspension buffer should be ~62 μL (60–70 μL range) (see Note 3). 9. Place one gel plug cast (10 plugs per cast, included in Bio-Rad CHEF Mammalian Genomic DNA Plug Kit) on an inverted metal block on ice. Set the smooth metal surface in contact

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with the adhesive of the base of the gel plug cast and avoid contact with condensed water. This will allow the agarose to cool faster once the well is filled (see Note 4). 10. Equilibrate cells in a cell suspension buffer at room temperature (RT) for 5 min. 11. Prepare two pipettes, set one to 37.5 μL and the other one to 110 μL. Use a wide-bore p200 tip and take 37.5 μL of melted 2% agarose at 37  C (pipet up and down twice in agarose) and add it to the 62.5 μL of cell suspension without mixing. Use the other P200 with a normal tip set at 110 μL and slowly mix ten times avoiding air bubble formation. Make sure the cell suspension and agarose are homogeneous. At this point, transfer the cell-agarose suspension to one well of the plug cast, creating a “dome” if needed. Repeat this step for each sample (Fig. 3a). 12. Place the plug cast on the inverted metal block in 4  C refrigerator for 30 min to allow the agarose to polymerize. Avoid contact with condensed water. 13. For each sample prepare one 50 mL tube with proteinase K solution by adding 170 μL of Qiagen Proteinase K enzyme and 2.5 mL of lysis buffer (each tube can have up to four plugs from the same sample). 14. Transfer plugs into the 50 mL tube containing proteinase K solution by removing the tape from the bottom of the plug cast and ejecting plugs into a 50 mL tube using the plug mold plunger (included in the Bio-Rad kit) (Fig. 3a). Make sure all plugs are fully submerged in lysis buffer. 15. Cap the 50 mL tube and incubate in a Thermomixer C or water bath for 1 h at 50  C with intermittent mixing (cycle 1: 15 s at 450 rpm followed by 15 min at 0 rpm) followed by 7 h at 37  C (cycle 2: 15 s at 450 rpm followed by 15 min at 0 rpm) and finally hold at 23  C (cycle 3: infinite 0 rpm). If using a water bath instead of a thermomixer, mix by hand every 10 min using a swirling motion for 1 h at 50  C. Then, transfer the 50 mL tube to a water bath at 37  C for 7 h to overnight. If plugs are immediately processed after 8 h of proteinase K treatment, then proceed until step 3 in Subheading 3.2, and then keep plugs in plug wash buffer at room temperature overnight. Start with step 4 in Subheading 3.2 the following day (see Notes 5 and 6). 3.2 Day 2: RNAseA Digestion

1. Prepare plug wash buffer (50 mM EDTA pH 8.0 and 10 mM TrisHCl pH 8.0) and TE buffer (10 mM Tris, pH 8.0, 1 mM EDTA) in molecular biology grade water. Mix thoroughly and leave at room temperature until use.

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B. Plug transfer to 1.5ml from screened cap A. Plug Preparation

30mins at 4ºC

C. Plug transfer between 1.5ml tubes before enzymatic reaction

D. Dialysis assembly

Fig. 3 Detailed depictions of individual steps in the END-seq protocol. (a) Plug preparation: cell suspension embedded in agarose is pipetted into a plug mold and allowed to solidify at 4  C. Polymerized plugs are then transferred to a 50 mL conical tube containing lysis buffer with proteinase K. (b) Plug transfer from cap: Plugs are transferred after washes from a screened cap to a 1.5 mL tube using a disposable spatula. (c) Plug transfer between tubes: Transfer plug to a new tube for enzymatic reactions by superimposing the opening of two 1.5 mL tubes and tapping on a benchtop until the agarose plug slides down into the new tube. (d) Dialysis assembly: a dialysis membrane is floated on the surface of 15 mL TE in a 6 cm petri dish. Melted agarose sample is pipetted onto the center of the membrane. The dish is covered and samples are dialyzed for 1 h

2. Drain proteinase K solution through the screen cap fitted on the 50 mL tube and tap each tube on a benchtop to push plugs down to the bottom of the tube. 3. Rinse plugs three times with 15 mL of plug wash buffer by adding and draining plug wash buffer through the screened cap. Swirl tube and make sure plugs are fully immersed in wash buffer during each rinse (see Note 7). Plugs can be kept in plug wash buffer for several hours to overnight at room temperature. 4. Drain plug wash buffer through screened cap and tap the tube to ensure that the plugs moved down to the bottom of each tube.

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5. Rinse plugs twice with 15 mL of TE buffer (as in step 3). 6. Wash plugs twice in 15 mL of TE with constant shaking for 15 min at RT (~180 rpm using an orbital platform shaker, or the minimum speed to see plugs moving). Make sure plugs are not settled at the bottom of the tube during washes. 7. Discard the last wash of TE through the screened cap. Remove the screened cap and add 2.5 mL of TE and 50 μL of Puregene RNAseA solution. Recap tubes, swirl to mix, make sure plugs are submerged, and incubate in a Thermomixer C for 1 h at 37  C with intermittent mixing (cycle: 10 s at 450 rpm followed by 10 min at 0 rpm). 8. Drain RNase solution through the screened cap and tap plugs to the bottom of the 50 mL tube. 9. Rinse plugs three times with 15 mL of plug wash buffer. Swirl tube and discard buffer through the screened cap and tap plugs to the bottom of the tube (as in Step 4) before adding the next wash. 10. Wash plugs four times with 15 mL of plug wash buffer for 15 min at RT with constant agitation (~180 rpm using an orbital platform shaker). Do not discard final 15 mL of plug wash buffer. 11. Leave the plugs in plug wash buffer at 4  C. This is a convenient stopping point. Plugs can be stored in plug wash buffer for up to 2 weeks at 4  C. 3.3 Day 3: DSB End Blunting, A-Tailing, and Ligation of END-Seq Adapter 1

Please be aware that if there are more than two plugs per sample, perform the reactions and washes in 50 mL tubes instead of 1.5 mL tubes. Multiply the volume of reaction according to the number of plugs in each tube. 1. Discard plug wash buffer, rinse twice with EB buffer (10 mM Tris pH 8.0 in nuclease free water), and perform four washes with agitation (~180 rpm using an orbital platform shaker) for 15 min each at RT with EB buffer. 2. After discarding the last EB wash, invert the 50 mL tube and remove the screened cap. Use a disposable spatula to transfer the plugs from the interior of the screened cap to 1.5 mL tubes containing 1 mL of EB (Fig. 3b; see Note 8). Steps 3 and 4 are for treatment with exonuclease VII for resection and end structure studies. If this is not the aim of the experiment, proceed to step 5. 3. Remove EB buffer with a P1000 micropipette tip (see Note 9). Then, add 1 mL of 1 homemade exonuclease VII buffer and rotate for 15 min at RT in a rotatory mixer at 8 rpm. Repeat with 500 μL of 1 NEB Exonuclease VII buffer and rotate for 15 min at RT in a rotatory mixer at 8 rpm.

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4. Remove the exonuclease VII buffer with a P1000 micropipette tip. Transfer plugs to a new 1.5 mL tube by inverting the opening of the tube over a new 1.5 mL tube (Fig. 3c). Tap on the bench until the plug moves down into the new tube. Remove any residual liquid from the wall and bottom of the Eppendorf tube with a P20 tip. Then, add 125 μL of exonuclease VII reaction mix (per plug): 25.00 μL 5 NEB Exonuclease VII buffer. 6.25 μL Exonuclease VII (10 U/μL). 93.75 μL H2O. Mix by tapping the tube with your fingers. Incubate at 37  C for 1 h with mixing at 400 rpm (in the thermomixer, mix by tapping the tube with your fingers every 20 min). 5. Remove the exonuclease VII reaction mix with a P200 micropipette tip and wash twice with 1 mL of 1 NEBuffer 4 for 15 min at RT in a rotatory mixer at 8 rpm. 6. Remove NEBuffer 4 with a P1000 micropipette tip and transfer plug to a new 1.5 mL tube (Fig. 3c). Remove any residual liquid from the tube with a P20 tip. Then, add 125 μL per plug of ExoT reaction mix on the plug: 12.5 μL 10 NEBuffer 4. 6.25 μL exonuclease T (5 U/μL). 106.25 μL H2O. Mix by tapping the tube with your fingers. Incubate at 25  C for 1 h with mixing at 400 rpm (in the thermomixer, mix by tapping the tube with your fingers every 20 min). 7. Remove the exonuclease T reaction mix and rinse with 1 mL of EB buffer. Decant and transfer plugs to a 50 mL tube containing 15 mL of EB and shake (~180 rpm using an orbital platform shaker) for 15 min at room temperature. Discard EB wash through the screened cap and tap plugs to the bottom of the tube before adding the next wash. Make sure plugs are submerged in EB buffer. Repeat the EB wash two additional times for a total of three washes. 8. After discarding the last EB wash, invert the 50 mL tube and remove the screened cap. Using a disposable spatula, scoop every plug from the interior of the screened cap to one 1.5 mL tube filled with 1 mL of 1 NEBNext dA-Tailing buffer and mix for 15 min at RT in a rotatory mixer at 8 rpm. Repeat the NEBNext dA-Tailing buffer wash one more time.

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9. Remove the 1 NEB dA-Tailing buffer with a P1000 micropipette tip and transfer each plug to a new 1.5 mL tube (Fig. 3c). Remove any residual buffer droplets and add 125 μL of dA-Tailing reaction mix (per plug): 12.5 μL NEB dA-Tailing buffer (10). 7 μL Klenow fragment (30 ! 50 exo-) (5 U/μL). 105.5 μL of H2O. Mix by tapping the tube with your fingers. Incubate at 37  C for 1 h with mixing at 400 rpm (in the thermomixer, mix by tapping the tube with your fingers every 20 min). 10. Remove the dA-Tailing reaction mix with a P200 micropipette tip, then add 1 mL of 1 NEBuffer 2 and mix for 15 min at RT in a rotatory mixer at 8 rpm. 11. Remove the 1 NEBuffer 2 with a P1000 micropipette tip and transfer plugs to a new 1.5 mL tube (Fig. 3c). Remove any residual buffer droplets and add 125 μL per plug of ice-cold Quick Ligation mix (prepare on ice) to the plug: 112 μL 2 Quick Ligase Buffer (NEB). 4 μL Annealed END-seq adapter 1 (10 μM). 4 μL Quick Ligase (2000 U/μL NEB). 5 μL H2O. Incubate at 25  C for 1 h with shaking at 400 rpm (in the thermomixer, mix by tapping the tube with the fingers every 20 min). 12. Remove the Quick Ligase Buffer mix with a P200 micropipette tip and rinse with 1 mL of plug wash buffer by pipetting up and down (be careful not to draw the plug into the pipette tip). Remove the plug wash buffer and add 1 mL of plug wash buffer and mix for 15 min at RT in a rotatory mixer at 8 rpm. Repeat this for a total of four washes. 13. Transfer plugs to a 50 mL tube with 45 mL of plug wash buffer by decanting and mix minimally for 8 h or overnight in a thermomixer C at 23  C with intermittent mixing (cycle: 10 s at 450 rpm followed by 15 min at 0 rpm). This wash step helps eliminate any unligated and free END-seq adapter 1. The presence of the free adapter will increase the amount of primerdimer formation in the PCR reaction. 3.4 Day 4: Plug Melting and DNA Shearing

Prepare in advance two water baths or heat blocks at 70  C and 43  C. Turn on the Covaris S220 sonicator and start the degassing and cooling procedure while performing drop dialysis (see step 8 below).

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1. Discard plug wash buffer from tubes in step 13, Subheading 3.3 above and rinse twice with TE buffer and perform four TE washes with agitation (~180 rpm on an orbital platform shaker) for 15 min at RT. 2. Discard TE Buffer through the screened cap and tap the tube to push plugs to the bottom of the tube. 3. Transfer one plug at a time with a disposable spatula to an empty 1.5 mL tube. Avoid any carryover of TE by touching the plug to the border of the screened cap to drain any excess liquid. If several plugs from the same sample are being processed simultaneously, pool them together in this step. 4. Pulse spin the microfuge tube for ~5 s at 2400  g to spin the agarose plug to the bottom of the tube. This will facilitate efficient heat transfer and rapid melting of the agarose in the next step. 5. Melt the agarose plugs by incubating tubes in a water bath or heat block at 70  C for 2 min. Incubate for 3 min if the volume is greater than 200 μL. 6. Immediately transfer tubes to a 43  C water bath and incubate for 5 min. 7. In a separate 1.5 mL tube, place the total volume of betaagarase enzyme needed for the melting step (1.5 μL  number of tubes) and leave at RT for at least 1 min before use. Add 1.5 μL of beta-agarase enzyme in the center of the melted plug in each tube. Slowly pipet up and down 3–5 times the entire volume of the melted plug with a P200 wide-bore tip and incubate at 43  C for 45 min (a temperature difference of 3  C can inactivate the enzyme). 8. Setup drop dialysis as follows: pipet 15 mL of TE buffer into a 6 cm Petri dish for each sample and float a 0.1 μm dialysis membrane above the TE buffer (Fig. 3d). Place the lid back on the Petri dish and let the membrane hydrate for 10 min. 9. Pulse spin microfuge tube containing beta-agarase digested samples for 1 s at 2400  g to spin down condensation. 10. Slowly pipet up and down the entire DNA solution with a P200 wide-bore tip and transfer it to the center of the dialysis membrane in a single drop (Fig. 3d). Do not touch the membrane with the tip as this could submerge the membrane resulting in sample loss. Place the lid on the petri dish and let samples dialyze for 1 h at room temperature (see Note 10). 11. Transfer DNA to a newly labelled 1.5 mL tube with a wide bore tip without touching the membrane. Add 1 μL of 10% SDS, 4 μL of proteinase K (20 mg/mL), vortex, and incubate for 15 min at 50  C.

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12. Spin down for 5 min at maximum speed in a microcentrifuge at RT to remove air bubbles. 13. Transfer DNA to a Covaris microTUBE AFA Fiber Pre-Slit Snap-Cap 6  16 mm using a P200 wide-bore tip. Pipet the entire DNA solution and slowly release it in the bottom of the Covaris tube to avoid making air bubbles. Fill with TE to the rim of the covaris tube (total tube volume is ~140 μL). If sample volume is more than 140 μL, add TE to have a total volume of 280 μL and distribute it between two Covaris tubes. Shear the DNA to a median size of 175 bp using the below cycle settings on a Covaris S220 series ultrasonicator. Cycle 1—duty 10%, intensity 175.0, cycles/burst 200, 240 s , 4–7  C. 14. Transfer the sheared DNA to a new tube and add 80 μL of TE to a total volume of 200ul. To this add 1 μL of glycogen, 20 μL of 3 M sodium acetate pH 5.2, vortex well, and add 500 μL of 100% ethanol at RT. Vortex (do not pipette) and incubate tubes in dry ice for 15 min. In the case of multiple tubes of the same sample, combine the sheared DNA and adjust volumes accordingly. 15. Spin chilled samples at maximum speed for 20 min at 4  C in a microcentrifuge. A white pellet should be visible. 16. Decant the supernatant, wash the pellet twice with 1 mL of ethanol 70% at RT, and spin at max speed for 5 min at 4  C in a microcentrifuge. 17. Decant the supernatant, carefully remove all the ethanol droplets with a vacuum aspirator and a micropipette, and air dry the DNA pellet at RT. 18. After the pellets are dry, add 70 μL of TE to the pellet, vortex, and incubate at 50  C for 5 min. Resuspend pellets with a micropipette P200 tip. 19. Spin at maximum speed for 5 min at RT in a microcentrifuge to pellet any insoluble material. Transfer solubilized DNA (supernatant) to a new tube and avoid taking any insoluble material or debris. Repeat this step as many times as is necessary until you have a clear DNA solution (see Note 11). 20. Measure DNA concentration with a nanodrop spectrometer (see Note 12 on the expected recovered DNA amounts based on starting cell type and numbers). 21. Store DNA at 20  C until ready to proceed to the next steps in Subheading 3.5. This is a convenient stopping point. Samples can be stored at 20  C for several weeks.

High-Resolution Mapping of DSBs and Resection

3.5 Day 5: Streptavidin Capture of Labeled DSB Ends, Blunting, A-Tailing Ligation of END-Seq Adapter 2, and PCR Library Amplification

25

Prepare in advance 9 mL of 1 bead wash and binding buffer (1 WB), 7 mL of EB per sample, and 10 mL of 2 bead binding buffer (2 BB) (see Note 13). 1. Add 35 μL of Dynabeads MyOne C1 in a 1.5 mL tube; wash beads with 1 mL of 1 WB. Place the tube on a DynaMag2 Magnet for 1 min to ensure that all the beads are collected on the tube wall and remove the 1 WB with a P1000 tip. Repeat for a total of two washes. 2. Resuspend beads in 70 μL (twice the original volume of the beads) of 2 WB buffer; then add 70 μL of DNA sample (from step 21, Subheading 3.4), vortex, and mix (700 rpm) for 30 min at RT in a thermomixer. Vortex every 15 min to avoid beads settling to the bottom of the tube. 3. Place tubes in the magnet, remove supernatant, and wash beads three times with 1 mL of 1 WB, twice with 1 mL of EB, and once with 1 mL of 1 NEB T4 Ligase Buffer. Change tubes after the first EB wash. Beads can be kept on the magnet in 1 NEB T4 Ligase Buffer while preparing for the end repair reaction (steps below). 4. Remove the 1 NEB T4 Ligase buffer using a P1000 tip, resuspend beads in 50 μL of end repair reaction buffer, and incubate for 30 min at 24  C with mixing (700 rpm) in the thermomixer. Vortex samples every 15 min. Prepare end-repair reaction buffer: 5 μL 10 NEB T4 DNA Ligase buffer. 2 μL 10 mM dNTPs. 2 μL end repair enzyme mix (1:5:5 large Klenow fragment: T4PolI:T4PNK). 41 μL H2O to make final volume 50 μL. Repair enzyme mix: 5 μL 3 U/μL T4 DNA Polymerase (NEB). 1 μL 5 U/μL large Klenow fragment (NEB). 5 μL 10 U/μL T4 DNA Polynucleotide Kinase (NEB). 5. Place tubes in the magnet, remove the END-repair reaction mix, and wash beads once with 1 mL 1 WB, twice with 1 mL of EB, and once with 1 mL of 1 NEBNext dA-Tailing Reaction Buffer. Change tubes after the first EB wash. Hold beads on the magnet in 1 NEBNext dA-Tailing Reaction Buffer while preparing the dA-Tailing reaction mix. 6. Remove 1 NEBNext dA-Tailing Reaction Buffer using a P1000 tip, resuspend beads in 50 μL of dA-tailing reaction mix, and incubate for 30 min at 37  C with mixing (700 rpm) in the thermomixer. Vortex every 15 min.

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Prepare dA-Tailing reaction mix: 5 μL 10 NEBNext dA-Tailing Reaction Buffer. 3 μL 5 U/μL NEB Klenow Fragment (30 ! 50 exo-). 42 μL of H2O. 7. Place tubes in the magnet, remove dA-Tailing reaction mix and wash beads once with 1 mL 1 NEBuffer 2. Hold beads on the magnet in 1 NEBuffer 2 while preparing the ligation reaction. 8. Remove 1 NEBuffer 2 using a P1000 tip and resuspend in 115 μL of ligation mix and incubate for 20 min at 25  C with mixing (700 rpm) in the thermomixer. Vortex every 15 min. Prepare ligation mix: (prepare on ice). 52.5 μL H2O. 57.5 μL 2 Quick Ligase Buffer (NEB). 2 μL END-seq adapter 2 (0.50 μM). 3 μL Quick Ligase (2000 U/μL NEB). 9. Inactivate the ligation reaction by adding 12 μL of EDTA 0.5 M, vortex, and place tubes in the magnet. Remove the ligation reaction using a P200 tip and wash beads three times with 1 mL 1 WB. Wash three times with 1 mL of EB. Change tubes after the first EB wash. 10. Remove the last EB wash and resuspend beads in 8 μL of EB. Add 10 μL of USER reaction, mix well, and incubate for 30 min at 37  C with mixing (700 rpm) in the thermomixer. Vortex every 15 min. USER reaction: 9 μL 2 Kapa HiFi HotStart Ready Mix. 1 μL of USER enzyme 1 U/μL (NEB). 11. Place samples on ice and add 1 μL of Illumina Truseq 8 nt barcoded primer p5 (50 μM) and 1 μL of Illumina Truseq 8 nt barcoded primer p7 (50 μM). Then, add 38 μL of PCR master mix and transfer the total 60 μL of each sample to a PCR tube (PCR strip). PCR master mix: 18 μL H2O. 20 μL 2 Kapa HiFi HotStart Ready Mix. 12. Perform the PCR using the following thermocycling parameters: (a) 45 s at 98  C. (b) 16 cycles. 15 s at 98  C. 30 s at 63  C.

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30 s at 72  C. (c) 5 min at 72  C. (d) Hold at 4  C. 13. Place the PCR strip in a DynaMag-PCR Magnet and transfer the supernatant to a new 1.5 mL tube. 14. The PCR product is purified using 108 μL of AMPure XP beads following the manufacturer’s protocol. Elute AMPure XP beads in 20 μL of EB. 15. Add loading buffer and run purified PCR product on a 2% agarose gel for 40 min at 90 V. 16. The PCR product should be between 200 bp and 400 bp and is excised from the gel. Avoid primer-dimer bands that are typically observed around 140 pb. (Fig. 4). 17. DNA is extracted from the gel slice using QIAquick Gel Extraction Kit following the manufacturer’s protocol. The final column elution of DNA is done using 17 μL of prewarmed (55  C) QIAquick EB. 18. Illumina libraries are quantified using the KAPA Library Quantification Kit for Illumina platforms following the manufacturer’s protocol. 19. Typically 8–12 samples can be pooled per sequencing run with a NextSeq 500/550 High Output Kit v2.5 (75 Cycles) that yields between 35 and 50 million 75 bases single-end reads for each sample. We follow Illumina’s NextSeq System Denature

500bp 200bp -

Fig. 4 Size distribution of sequencing-ready PCR product. The purified PCR library product is loaded on a 2% agarose gel and run for 40 min at 90 V. This results in a tight smear (red box) with a size distribution between 200 to 400 bp. Following gel purification and quantification, the DNA is ready to load onto an Illumina Next-Generation Sequencing Platform

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and Dilute Libraries Guide and recommend barcode compatibility. Libraries are analyzed using a standard bioinformatics pipeline using Bowtie as an aligner and MACS as the peakcaller as previously described in [11–14].

4

Notes 1. If you suspect that there are dead cells, isolate live cells by Ficoll or Percoll. The following protocol is optimized for lymphocytes using Ficoll. For other cell types we recommend optimizing a Percoll gradient according to the density of the cell type. (a) In 15 or 50 mL tubes, add 5 mL of lymphocyte separation medium from Lonza (http://bio.lonza.com/uploads/tx_ mwaxmarketingmaterial/Lonza_ ManualsProductInstructions_Lymphocyte_Separation_ Medium.pdf). The lymphocyte separation medium is a mixture of Ficoll and sodium diatrizoate (Hypaque) with density adjusted to 1.077 g/mL, and it can be used with human and mouse lymphocytes. (b) Centrifuge cells, remove media, resuspend pellet in 1 mL of cell growth media with 10% of serum, add 6 mL of PBS, and with a P1000 micropipette tip very slowly transfer the cell suspension on top of the lymphocyte separation media Ficoll solution. Be careful to avoid breaking the interphase between the cell suspension and the Ficoll. The interphase can be maintained if one inclines the tube and adds the cell suspension along the sides of the wall. (c) Spin at 450  g for 25 min with the slowest acceleration and deceleration without any brakes. (d) Live cells fall in the interphase between Ficoll and PBS media while dead cells pellet in the Ficoll. Live cells are recovered using a P1000 micropipette tip and transferred to a new 50 mL tube with 40 mL complete media. Do not take more than 3 mL from the interphase. (e) Spin for 7 min at 450  g. Discard media and resuspend the cell pellet in fresh complete media. Cells are counted and viability assessed with trypan blue staining. (f) Aliquot the number of cells necessary to make agarose plugs. 2. When washing the cells in PBS in preparation for embedding in agarose plugs, add PBS to wash the cells without disturbing the pellet after the first centrifugation. Small cells like lymphocytes can become very sticky. This washing step of the pellet without resuspension helps to remove any residual media components while minimizing cell loss.

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3. The number of cells that fits in a plug depends on the cell size. The volume of cells and suspension buffer has to be between 62 and 70 μL, but the volume of suspension buffer added cannot be lower than 55 μL (meaning that the volume of the cellular pellet has to be between 7 and 15 μL). When adding 37.5 μL of 2% agarose to the cell suspension, the final agarose concentration should be between 0.75% and 0.8%. If the cell pellet size is too big, make two plugs. Lower volumes increase agarose concentrations and make the plugs so compacted that enzymes cannot enter, thereby dramatically lowering the efficiency of the entire END-seq process. Concentrations below 0.75% increase fragility of the plugs. Large numbers of cells in the plug increase mechanical resistance of the plugs in later steps while plugs with less than two million cells become very fragile (easily chipped) and necessitate special care. 4. When mixing agarose with the cells it is best to be quick to avoid solidification of the cell-agarose mixture before pipetting into the plug mold. 5. Monitor that plugs become clear during the first 30 min of the proteinase K reaction. After 10 min, you can observe the proteinase K advancing to the center of the plug and the borders of the plug become clearer. After 35–40 min the whole plug should be clear. If there are too many cells or the agarose concentration is too high, the center will remain undigested after 35–40 min. In this case, discard the plugs and start again. Although the effect of proteinase K during the first 30 min on plug clarity should be obvious, plugs from some cell types with a rich extracellular matrix can maintain cloudiness even after proteinase K treatment. 6. After the proteinase K reaction, plugs can be rinsed with wash buffer and stored until the next day at room temperature. Do not store them at 4  C as the SDS in the lysis buffer remains in the plug. At 4  C the SDS precipitates and will form crystals that could damage the DNA. If this happens, discard the plugs and start again. 7. Avoid plugs from being stranded in the screen cap or on the wall of the 50 mL tubes during reactions and washes. Always check how many plugs are in the buffer by stirring after every rinse and wash. If a plug is stranded, it will dry and enzymes will not be able to permeate the plug resulting in damaged DNA. In such a scenario, it is best to discard the plug and start again. We recommend discarding and adding washes in the same location on the screened cap. This will help prevent plugs from sticking to the cap after decanting. To facilitate the entrance of liquid through the screened cap, slightly tilt the 50 mL tube when pouring in fresh wash solution.

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8. For transferring plugs from the screen cap to Eppendorf tubes, we recommend using a disposable spatula to avoid DNA contamination between different samples. Also avoid mixing plugs from different samples in the same proteinase K or RNAse reactions and/or washes. In the case of transferring plugs from one 1.5 mL tube to another, avoid using spatulas, simply transfer by connecting the opening of Eppendorf tubes, and decant the plug into the new tube (Fig. 3c). 9. When removing liquid from the 1.5 mL tubes with a plug inside, tilt the tube and introduce the tip of a 1000 mL micropipette to the bottom of the tube and aspirate. Be careful to not pull the plug into the tip as this will break the plug. 10. During the spot dialysis step, use forceps to grip the membrane by its edge and gently place on the TE surface while simultaneously maintaining the membrane in a horizontal position to prevent dipping or sinking of membrane. 11. Depending on the cell type, some white debris can appear after resuspending the DNA pellet in TE after ethanol precipitation. In this scenario, centrifuge the solution and transfer the clear supernatant to a new tube while avoiding the debris. Sometimes the debris does not precipitate and remains on the surface of the solution. In this case, remove the solution from the bottom of the tube. 12. Estimations of DNA concentration after Covaris sonication and DNA precipitation: for 20 million B-cells, around 65 micrograms of DNA should be recovered. For four million B-cells, approximately 14 micrograms are obtained. 13. Make fresh bead binding and wash solutions each time an experiment is performed. Mix beads well by vortexing and pipetting up and down before every use. 14. Order END-seq adapters 1 and 2 with HPLC purification. Reconstitute them by adding 1 NEB T4 Ligase Buffer to a concentration of 10 μM. Vortex the oligos and incubate at 50  C for at least 5 min. Distribute the oligos in 500 μL aliquots in screwcap tubes and assemble them in a floater. Heat 1 L of water in a glass beaker in the microwave oven until it boils. Measure temperature with a thermometer and make sure it is higher than 95  C. Put the floater with screwcap tubes containing oligos on the surface of the hot water and wait until it reaches room temperature, typically around 3–4 h. Finally, pool all aliquots and make 100 μL working aliquots. Aliquoted oligos can be stored at 20  C indefinitely.

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References 1. Iacovoni JS et al (2010) High-resolution profiling of gammaH2AX around DNA double strand breaks in the mammalian genome. EMBO J 29(8):1446–1457 2. Szilard RK et al (2010) Systematic identification of fragile sites via genome-wide location analysis of gamma-H2AX. Nat Struct Mol Biol 17(3):299–305 3. Yamane A et al (2011) Deep-sequencing identification of the genomic targets of the cytidine deaminase AID and its cofactor RPA in B lymphocytes. Nat Immunol 12(1):62–69 4. Barlow JH et al (2013) Identification of early replicating fragile sites that contribute to genome instability. Cell 152(3):620–632 5. Frock RL et al (2015) Genome-wide detection of DNA double-stranded breaks induced by engineered nucleases. Nat Biotechnol 33 (2):179–186 6. Tsai SQ et al (2015) GUIDE-seq enables genome-wide profiling of off-target cleavage by CRISPR-Cas nucleases. Nat Biotechnol 33 (2):187–197 7. Wang X et al (2015) Unbiased detection of off-target cleavage by CRISPR-Cas9 and TALENs using integrase-defective lentiviral vectors. Nat Biotechnol 33(2):175–178

8. Crosetto N et al (2013) Nucleotide-resolution DNA double-strand break mapping by nextgeneration sequencing. Nat Methods 10 (4):361–365 9. Yan WX et al (2017) BLISS is a versatile and quantitative method for genome-wide profiling of DNA double-strand breaks. Nat Commun 8:15058 10. Longhese MP et al (2010) Mechanisms and regulation of DNA end resection. EMBO J 29 (17):2864–2874 11. Canela A et al (2016) DNA breaks and end resection measured genome-wide by end sequencing. Mol Cell 63(5):898–911 12. Canela A et al (2017) Genome organization drives chromosome fragility. Cell 170 (3):507–521. e18 13. Canela A et al (2019) Topoisomerase II-induced chromosome breakage and translocation is determined by chromosome architecture and transcriptional activity. Mol Cell 75 (2):252–266. e8 14. Tubbs A et al (2018) Dual roles of poly(dA:dT) tracts in replication initiation and fork collapse. Cell 174(5):1127–1142. e19

Chapter 3 Resection of a DNA Double-Strand Break by Alkaline Gel Electrophoresis and Southern Blotting Erika Casari, Elisa Gobbini, Michela Clerici, and Maria Pia Longhese Abstract Generation of 30 single-stranded DNA (ssDNA) at the ends of a double-strand break (DSB) is essential to initiate repair by homology-directed mechanisms. Here we describe a Southern blot-based method to visualize the generation of ssDNA at the ends of site-specific DSBs generated in the Saccharomyces cerevisiae genome. Key words DNA double-strand breaks, Resection, Single-stranded DNA, HO endonuclease, Southern blot, S. cerevisiae

1

Introduction DNA double-strand breaks (DSBs) are among the most cytotoxic DNA lesions, because failure to repair them can result in loss of genetic information or cell death, whereas inaccurate repair can lead to chromosome rearrangements [1, 2]. Eukaryotic cells can repair a DSB by either end-joining or homology-directed repair (HDR) mechanisms. The canonical end-joining mechanism, nonhomologous end joining (NHEJ), occurs via direct re-ligation of the broken DNA ends [3]. By contrast, HDR is a more complex process that uses DNA information stored in a homologous doublestranded DNA (dsDNA) as the template to reconstitute any missing genetic information at the break site [4, 5]. The first step of HDR is the nucleolytic degradation of the 50 DNA strands at both sides of the DSB to generate 30 -ended singlestranded DNA (ssDNA) through a process termed DNA end resection [6]. The ssDNA tails are first coated by the ssDNA binding complex replication protein A (RPA), which is then replaced by the recombinase Rad51 to form a right-handed helical filament that searches and catalyzes the invasion of duplex homologous DNA molecules [4, 5]. In vegetatively growing cells, HDR uses the sister

Andre´s Aguilera and Aura Carreira (eds.), Homologous Recombination: Methods and Protocols, Methods in Molecular Biology, vol. 2153, https://doi.org/10.1007/978-1-0716-0644-5_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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chromatid as the repair template, and this restricts recombination to the S and G2 phases of the cell cycle when the sister chromatid is available. This cell-cycle control of recombination is based on activation of key resection proteins by cyclin-dependent kinase (CDK)catalyzed phosphorylation events [7]. Much of our knowledge about DNA end resection has come from genetic and biochemical studies in the budding yeast Saccharomyces cerevisiae, in which expression of the HO endonuclease creates a site-specific DSB located adjacent to the Y region at the MAT locus [8]. The HO-induced DSB is normally repaired by gene conversion using one of the two silent HML or HMR donors of mating-type information resulting in mating-type switching [8]. By using a galactose-inducible HO endonuclease gene [9], galactose addition leads the majority of cells to generate a DSB at the MAT locus, whose repair occurs synchronously so that the kinetics of DSB resection and repair can be followed and defined in detail. Therefore, S. cerevisiae has been considered an ideal model organism to study the response to DSBs, and different methods have been developed to monitor DSB resection in this organism. Here we illustrate a Southern blot approach to visualize the kinetics of resection at an HO-induced DSB during a time course experiment in a single gel blot. This assay relies on the ability of restriction enzymes to cut dsDNA but not ssDNA. Therefore, the ssDNA generated during resection of a DSB end is resistant to endonucleolytic cleavage and can be visualized as appearance of slower migrating bands that can be detected by Southern blot under denaturing conditions with a probe that anneals to the unresected 30 end at one side of the DSB (Fig. 1). This approach requires yeast strains (i.e., JKM139) expressing the HO gene under the control of a galactose-inducible promoter [10]. Galactose addition results in high levels of HO production, which recognizes and cleaves a unique 24-bp cleavage sequence at the MAT locus. To allow the visualization of the resection products, the DSB generated upon HO induction in the JKM139 strain is irreparable, as this strain carries the deletion of both the HML and HMR cassettes that are used to repair the HO-induced DSB at the MAT locus by gene conversion [10]. The same method can be used to monitor resection of DSBs located at different genomic loci [11] or generated by other nucleases [12].

2

Materials Prepare all solutions using ultrapure water (ddH2O, prepared by purifying deionized water to obtain a sensitivity of 18 MΩ cm at 25  C) and analytical-grade reagents. Prepare and store all solutions at room temperature unless otherwise specified.

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Fig. 1 Schematic representation of the system used to detect resection of the HO-induced DSB at the MAT locus. HO induction generates a DSB at the MAT locus. SspI (S)-digested genomic DNAs run under alkaline conditions are hybridized with the indicated RNA probe that anneals to the unresected strand at one side of the DSB. The probe reveals a 1.1 kb fragment representing the uncut MAT locus. When HO catalyzes the cleavage, a HO-cut fragment of 0.9 kb is produced. 50 –30 nucleolytic degradation progressively eliminates SspI sites, generating longer SspI fragments (r1–r7) detected by the probe 2.1 Induction of a DNA Double-Strand Break in S. cerevisiae Cells

1. S. cerevisiae JKM139 strain (hoΔ hmlΔ::ADE1 MATa hmrΔ:: ADE1 ura3-52 leu2-3112 trp1::hisG lys5 ade1-100 ade3::GAL:: HO) (J. Haber, Waltham University, USA) [10]. 2. Yeast extract peptone dextrose (YEPD): 2% (w/v) Bacto peptone, 1% (w/v) yeast extract, 2% (w/v) dextrose, 0.005% adenine hemisulfate salt. Dissolve in ddH2O. Autoclave. 3. YEP + raffinose (YEPR). 2% (w/v) Bacto peptone, 1% (w/v) yeast extract, and 0.005% adenine hemisulfate salt. Dissolve in ddH2O. Autoclave. Add sterilized raffinose from 30% solution to 2%. 4. 30% raffinose. Dissolve raffinose pentahydrate ddH2O. Autoclave or sterilize by filtration.

in

5. 30% galactose. Dissolve galactose 99.0% in ddH2O. Sterilize by filtration. Do not autoclave (see Note 1). 2.2 Extraction of Genomic DNA

1. Spheroplasting solution: 0.9 M sorbitol, 0.1 M ethylenediaminetetraacetic acid (EDTA), pH 7.5. 2. Zymolyase solution. Dissolve 2 mg/mL Zymolyase 20T® (see Note 2) from Arthrobacter luteus (Nacalai Tesque or similar) in spheroplasting solution containing 14 mM β-mercaptoethanol.

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3. 1 TE: 10 mM Tris–HCl pH 7.5, 1 mM EDTA pH 7.5. Autoclave. 4. Lysis solution: 2.2% sodium dodecyl sulfate (SDS), 278 mM EDTA, 445 mM Tris-base pH 8.5. Prepare the lysis solution just before use. 5. 5 M potassium acetate. Dissolve potassium acetate in ddH2O. Autoclave. 6. 96% and 70% ice-cold ethanol. Store at 20  C. 7. RNase solution. Dissolve 10 mg/mL RNase A, Dnase free, in 10 mL of 10 mM Tris–HCl pH 7.5, 15 mM NaCl. Heat to 100  C for 5 min and cool down at room temperature. Prepare small aliquots and store them at 20  C. 8. 2-propanol anhydrous 99.5%. 9. 1 TAE (Tris acetate–EDTA) buffer: 40 mM Tris, 20 mM acetic acid, 1 mM EDTA. For 1 L of 50 TAE buffer stock, dissolve 242 g of Tris base in approximately 600 mL ddH2O. Add 57.1 mL glacial acetic acid and 100 mL 0.5 M EDTA and bring the final volume to 1 L with ddH2O. Before use, dilute in ddH2O to a final concentration of 1. 10. 6DNA loading bromophenol blue.

buffer:

30%

glycerol,

0.25%

11. Ethidium bromide solution: prepare a stock of 10 mg/mL ethidium bromide in ddH2O. 12. Agarose gel: melt 0.8% agarose in 1 TAE buffer. Cool at approximately 60  C and add 10 mg/mL ethidium bromide solution to a final concentration of 1 μg/mL. Pour the gel into a gel tank and insert a comb. 13. UV lamp with a camera. 2.3 DNA Digestion and Denaturation

1. 3 M sodium acetate pH 5.2. Dissolve 3 M sodium acetate in ddH2O. Adjust pH to 5.2 with glacial acetic acid. Autoclave. 2. SspI restriction enzyme and 10 buffer supplied by the distributor. 3. 0.5 M EDTA pH 8.0. Dissolve 0.5 M EDTA in ddH2O. Adjust pH to 8.0 with sodium hydroxide (NaOH). Autoclave. 4. 1 alkaline loading buffer: 50 mM NaOH, 1 mM EDTA pH 8.0, 2.5% Ficoll (type 400) in ddH2O, 0.025% bromophenol blue.

2.4 Alkaline Gel Electrophoresis and Transfer

1. Horizontal electrophoresis system with a gel running chamber (gel size 25  20 cm) and 32-tooth comb (thickness 1.0 mm and width of teeth 4.0 mm). 2. 1 alkaline electrophoresis buffer: 50 mM NaOH, 1 mM EDTA pH 8.5.

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3. A glass plate that fits the gel. 4. 0.25 N hydrochloric acid (HCl). Dilute HCl in ddH2O just before use. 5. 0.5 N NaOH, 1.5 M NaCl. Dissolve in ddH2O just before use. 6. Nylon hybridization transfer membrane (GeneScreen® from PerkinElmer or equivalent). 7. 20 SSC buffer: 3 M NaCl, 300 mM sodium citrate. Adjust pH to 7.0 with HCl. Autoclave. 8. Paper towels. 9. Parafilm. 10. Whatman 3 MM paper. 11. Neutralization solution: 0.5 M Tris–HCl pH 7.5, 1 M NaCl. 12. UV crosslinker. 2.5

Probe Labeling

1. Plasmid pML514 (available upon request) carrying a 900-bp fragment of the MAT locus cloned downstream to the T7 bacteriophage promoter. This plasmid was constructed by inserting in pGEM®-7Zf() part of the MAT DNA sequence obtained by PCR using yeast genomic DNA as a template and PRP643 (50 -CGG AAT TCC CTG GTT TTG GTT TTG TAG AGT GG-30 ) and PRP644 (50 -CGG AAT TCG AAA CAC CAA GGG AGA GAA GAC-30 ) as primers. 2. In vitro Riboprobe System-T7 (purchased from Promega or equivalent) containing recombinant RNasin® RNase inhibitor, 10 mM rATP, 10 mM rCTP, 10 mM rGTP, 10 mM rUTP, 100 mM dithiothreitol (DTT), 5 transcription optimized buffer, T7 RNA polymerase, RQ1 Rnase-free DNase, nuclease-free water. 3. BamHI restriction enzyme and appropriate buffer. 4. rUTP–α32P (800 Ci/mmol specific activity). 5. Sephadex G-50 chromatography column (GE Healthcare or equivalent).

2.6 Filter Hybridization

1. Hybridization tubes and hybridization oven. 2. Formamide hybridization buffer: 5 SSPE, 50% formamide, 6% dextran sulfate sodium salt, 4 Denhardt’s solution (dilute 50 Denhardt’s solution from Sigma Aldrich or equivalent), 100 μg/mL deoxyribonucleic acid sodium salt from salmon testes, 200 μg/mL yeast tRNA. 3. 20 SSPE buffer: 3 M NaCl, 0.2 M NaH2PO4, 20 mM EDTA. Adjust pH to 7.4 with NaOH. Autoclave. 4. 1 SSPE, 0.1% SDS. 5. 0.1 SSPE, 0.1% SDS.

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6. 0.2 SSPE, 0.1% SDS. 7. Autoradiography cassette with intensifying screens. 8. Autoradiography films or imaging plates.

3

Methods

3.1 Induction of DNA Double-Strand Break

1. Inoculate cells in 50–100 mL YEPD medium. 2. Grow cells at 26  C to a concentration of 105–106 cells/mL. 3. Harvest cells by centrifugation. 4. Wash cells with YEPR to remove glucose and resuspend the cell pellet in an equal volume of YEPR. 5. Grow cells overnight at 26  C. 6. When cells reach a concentration of 8  106 to 107 cells/mL, collect a 50 mL sample by centrifugation for the uninduced control. Then, add galactose to 3% final concentration to the remaining cell culture to induce expression of the HO endonuclease (see Note 3). 7. Grow cells at 26  C and collect 50 mL samples at each time point by centrifugation. During the experiment the efficiency of DSB formation can be determined (see Note 4). 8. Wash cells with 1 mL of 0.9 M sorbitol, 0.1 M EDTA. 9. Freeze and store the pellets at 20  C.

3.2 Extraction of Genomic DNA

1. Thaw the cell pellet at room temperature and resuspend the pellet in 400 μL spheroplasting solution, 14 mM β-mercaptoethanol (freshly made). 2. Add 100 μL zymolyase solution and invert the tube 4–6 times. Incubate the tube at 37  C. Check the formation of spheroplasts after 30–60 min under a light microscope (see Note 5). 3. When >95% cells become spheroplasts, centrifuge at 15,000  g for 1 min, and carefully remove the supernatant with a pipetteman. 4. Carefully resuspend spheroplasts in 400 μL 1 TE. 5. Add 90 μL lysis solution and incubate in a water bath for 30 min at 65  C. 6. Add 80 μL 5 M potassium acetate and invert several times. Place the tube in a cold room for approximately 1 h. 7. Spin down at 15,000  g, 4  C for 30 min. Transfer the supernatant to a 1.5 mL tube. Discard the pellet. 8. Add 1 mL ice-cold 96% ethanol and invert the tube several times. A white cloudy precipitate should form. Place the tube at 80  C for 30 min to facilitate precipitation.

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9. Spin down at 15,000  g, 4  C for 10 min and remove the supernatant. 10. Wash the pellet with 1 mL ice-cold 70% ethanol. Remove the ethanol. 11. Leave the tube open at room temperature for the ethanol to evaporate and the pellet to dry. 12. Resuspend the dried pellet in 500 μL 1 TE and gently dissolve it (do not vortex). 13. Add 2.5 μL RNase solution and incubate for 1 h at 37  C. 14. Add 500 μL 2-propanol and invert the tube several times. Place the tube at 80  C for 30 min (or overnight) to facilitate the precipitation. 15. Spin at 15,000  g, 4 supernatant.



C for 30 min and remove the

16. Wash the pellet with 1 mL ice-cold 70% ethanol. Remove the ethanol. 17. Leave the tube open at room temperature for the ethanol to evaporate and the pellet to dry. 18. Add 30 μL 1 TE and gently dissolve the pellet (do not vortex). 19. Load 1 μL of each sample (added to 10 μL of 1 DNA loading buffer) on a 0.8% agarose gel with ethidium bromide and run in 1 TAE buffer. Check the quality of the extracted DNA under an UV lamp (see Note 6). 3.3 DNA Digestion and Denaturation

1. Digest 10–15 μg of genomic DNA with 10 U of SspI or other restriction enzymes that cut dsDNA but not ssDNA (see Note 7). Digest with 1 enzyme buffer in a total volume of 70 μL for 5–6 h at 37  C. 2. Test 1 μL of each digestion reaction on a 0.8% agarose gel with ethidium bromide and run in 1 TAE buffer to check that all samples are digested. 3. Precipitate the digested DNA with 2 volumes of 96% ethanol, 5 mM EDTA pH 8.5, 0.3 M sodium acetate pH 5.2. Place the tubes overnight at 80  C to facilitate precipitation. 4. Spin at 15,000  g, 4 supernatant.



C for 30 min and remove the

5. Wash the pellet with 1 mL ice-cold 70% ethanol. Remove the ethanol. 6. Air dry until the pellet appears glassy. 7. Add 25 μL 1 alkaline loading buffer. Leave the tube at room temperature for 1.5–2 h and gently mix them every 15–30 min (do not vortex). A 1 kb DNA ladder can also be treated with

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1 alkaline loading buffer and then used as a marker for the size estimation of the DNA resection fragments. 8. Apply 1 μL denatured DNA to 10 μL alkaline loading buffer and load on a 0.8% agarose gel with ethidium bromide. Run in 1 TAE buffer to check whether the DNA is completely denatured (see Note 8) and to evaluate the amount of DNA in each digestion. Equilibrate the DNA concentration in different tubes by adding an appropriate amount of 1 alkaline loading buffer. 3.4 Alkaline Gel Electrophoresis and Transfer

1. Melt 0.8% agarose in 450 mL ddH2O and pour into a gel tray. 2. Put the gel tray in a large gel box in 1 alkaline electrophoresis buffer. Allow the gel to equilibrate for 30 min or longer (see Note 9). 3. Load 24 μL of each sample dissolved and equilibrated in alkaline loading buffer. 4. Run the gel overnight at voltages 95% cells appear large budded, add galactose to induce HO expression. Nocodazole-mediated arrest can be maintained for 8–10 h. To monitor resection in the G1 phase, MATa haploid cells can be treated with the α-factor pheromone. As these cells can recover from the α-factor block after 1.5–2 h due to α-factor degradation by the Bar1 protease, strains carrying the deletion of the BAR1 gene need to be used to obtain a persistent G1 arrest [13]. Dissolve 1 mg/mL α-factor in ddH2O and add this solution to exponentially growing YEPR cells to a final α-factor concentration of 0.5 μg/mL. After 2 h at 26  C, when >95% of cells appear unbudded, add galactose to induce HO expression. α-actor-mediated arrest in bar1Δ cells can be maintained for at least 8 h. 4. Generation of a single DSB activates the DNA damage checkpoint that arrests the metaphase to anaphase transition [14]. Therefore, the effectiveness of DSB induction in checkpoint-proficient strains can be determined by checking

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under a microscope that cells become dumbbell shaped (the bud is roughly the same size as the mother cell) after 4–6 h from galactose induction. 5. To check the efficiency of zymolyase digestion, prepare two drops (5 μL) of the same sample on a microscope slide and add 1–2 μL of 10% SDS to one of them. Spheroplasts lyse in SDS solution. >95% spheroplasts can be obtained in 40–50 min. 6. If high-molecular weight DNA molecules are extracted and completely resuspended in 1 TE buffer, a single band should be detectable in the gel without smears or signals retained in the well. 7. Restriction enzymes digest dsDNA by cleaving two phosphodiester bonds, one within each strand of the duplex DNA. Only few restriction enzymes cleave ssDNA, although at low efficiency. 8. When DNA is completely denatured, a smear is detectable in the agarose gel. 9. The agarose gel is equilibrated in alkaline electrophoresis buffer after solidification because the addition of NaOH to a warm agarose solution causes polysaccharide hydrolysis. Alternatively, the agarose gel can be melted in ddH2O and then cooled down to 60  C, so that NaOH to 50 mM and EDTA pH 8.5 to 1 mM can be added just before pouring the gel. 10. Ethidium bromide is omitted from alkaline agarose gels because it does not bind DNA at high pH. DNA can be stained with ethidium bromide after the electrophoresis. However, DNA will be faint because the ethidium bromide does not bind very well to ssDNA. 11. DNA can be transferred onto either neutrally or positively charged nylon membranes. 12. In vitro transcription of templates containing 30 overhangs could give rise to erroneous transcripts [15]. Therefore, plasmids should not be linearized with restriction enzymes that leave 30 overhangs. 13. Plasmid DNA must be completely cleaved because uncut plasmid DNA can give rise to long transcripts including vector sequences that will incorporate radiolabeled rNTPs. 14. The linearized plasmid DNA should be concentrated because the in vitro transcription reaction requires a small volume of template. 15. A good signal can be obtained after an overnigth exposure to an autoradiography film with freshly labeled rUTP–α32P.

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Acknowledgments We thank members of the Longhese lab for discussion. This article was supported by the Fondazione AIRC under IG 2017—ID. 19783 project—P.I. Longhese Maria Pia and Progetti di Ricerca di Interesse Nazionale (PRIN) 2015 to M.P.L. E.C. was supported by a fellowship from the Italian Ministry of University and Research (MIUR) through grant “Dipartimenti di Eccellenza—2017.” References 1. Jackson SP, Bartek J (2009) The DNA-damage response in human biology and disease. Nature 461:1071–1078. https://doi.org/10.1038/ nature08467 2. Liu P, Carvalho CM, Hastings PJ, Lupski JR (2012) Mechanisms for recurrent and complex human genomic rearrangements. Curr Opin Genet Dev 22:211–220. https://doi.org/10. 1016/j.gde.2012.02.012 3. Chang HHY, Pannunzio NR, Adachi N, Lieber MR (2017) Non-homologous DNA end joining and alternative pathways to doublestrand break repair. Nat Rev Mol Cell Biol 18:495–506. https://doi.org/10.1038/nrm. 2017.48 4. Mehta A, Haber JE (2014) Sources of DNA double-strand breaks and models of recombinational DNA repair. Cold Spring Harb Perspect Biol 6(9):a016428. https://doi.org/10. 1101/cshperspect.a016428 5. Kowalczykowski SC (2015) An overview of the molecular mechanisms of recombinational DNA repair. Cold Spring Harb Perspect Biol 7:a016410. https://doi.org/10.1101/ cshperspect.a016410 6. Bonetti D, Colombo CV, Clerici M, Longhese MP (2018) Processing of DNA ends in the maintenance of genome stability. Front Genet 9:390. https://doi.org/10.3389/fgene.2018. 00390 7. Trovesi C, Manfrini N, Falcettoni M, Longhese MP (2013) Regulation of the DNA damage response by cyclin-dependent kinases. J Mol Biol 425:4756–4766. https://doi.org/10. 1016/j.jmb.2013.04.013 8. Lee CS, Haber JE (2015) Mating-type gene switching in Saccharomyces cerevisiae. Microbiol Spectr 3:MDNA3-0013-2014. https:// doi.org/10.1128/microbiolspec.MDNA30013-2014

9. Jensen R, Sprague GF Jr, Herskowitz I (1983) Regulation of yeast mating-type interconversion: feedback control of HO gene expression by the mating-type locus. Proc Natl Acad Sci U S A 80:3035–3039. https://doi.org/10. 1073/pnas.80.10.3035 10. Lee SE, Moore JK, Holmes A, Umezu K, Kolodner RD, Haber JE (1998) Saccharomyces Ku70, mre11/rad50 and RPA proteins regulate adaptation to G2/M arrest after DNA damage. Cell 94:399–409. https://doi.org/ 10.1016/s0092-8674(00)81482-8 11. Clerici M, Mantiero D, Lucchini G, Longhese MP (2005) The Saccharomyces cerevisiae Sae2 protein promotes resection and bridging of double strand break ends. J Biol Chem 280:38631–38638. https://doi.org/10. 1074/jbc.M508339200 12. Manfrini N, Guerini I, Citterio A, Lucchini G, Longhese MP (2010) Processing of meiotic DNA double strand breaks requires cyclindependent kinase and multiple nucleases. J Biol Chem 285:11628–11637. https://doi. org/10.1074/jbc.M110.104083 13. Chan RK, Otte CA (1982) Physiological characterization of Saccharomyces cerevisiae mutants supersensitive to G1 arrest by a factor and alpha factor pheromones. Mol Cell Biol 2:21–29. https://doi.org/10.1128/mcb.2.1.21 14. Pellicioli A, Lee SE, Lucca C, Foiani M, Haber JE (2001) Regulation of Saccharomyces Rad53 checkpoint kinase during adaptation from DNA damage-induced G2/M arrest. Mol Cell 7:293–300. https://doi.org/10.1016/ S1097-2765(01)00177-0 15. Schenborn ET, Mierendorf RC (1985) A novel transcription property of SP6 and T7 RNA polymerases: dependence on template structure. Nucleic Acids Res 13:6223–6236. https://doi.org/10.1093/nar/13.17.6223

Chapter 4 Analysis of DNA Double-Strand Break End Resection and Single-Strand Annealing in S. pombe Zhenxin Yan, Sandeep Kumar, and Grzegorz Ira Abstract DNA double-strand break (DSB) end resection is an essential step for homologous recombination. It generates 30 single-stranded DNA needed for the loading of the strand exchange proteins and DNA damage checkpoint proteins. To study the mechanism of end resection in fission yeast, we apply a robust, quantitative and inducible assay. Resection is followed at a single per genome DSB synchronously generated by the tet-inducible I-PpoI endonuclease. An additional assay to follow resection involves recombination between two direct repeats by single-strand annealing (SSA), since SSA requires extensive resection to expose two single-strand repeats for annealing. The kinetics of resection and SSA repair are then measured using Southern blots. Key words DNA double-strand break, End resection, Single-strand annealing, Southern blotting

1

Introduction DNA double-strand breaks are repaired by either nonhomologous end joining (NHEJ) or by homologous recombination (HR). The first step of HR called resection is the degradation of the 50 strands that generates 30 single-stranded DNA (ssDNA) [1]. The 30 ssDNA is essential for loading of the strand exchange protein Rad51 as well as DNA damage checkpoint proteins. DSB ends are initially resected by the MRN complex (MRX in budding yeast) assisted by Ctp1 (Sae2 in budding yeast) to generate short 30 ssDNA (initial resection) followed by long-range resection by either Exo1 or Rhq1-Dna2 (Sgs1-Dna2 in budding yeast) to generate longer 30 ssDNA (extensive resection). Here we describe a method to follow the resection kinetics at an inducible DSB in fission yeast (Schizosaccharomyces pombe). The gene encoding I-PpoI endonuclease controlled by the tetO7 promoter was inserted at the leu1 locus [2]. The endogenous I-PpoI cleavage sites within rDNA repeats were eliminated by growing cells in the presence of constant I-PpoI expression where survivor

Andre´s Aguilera and Aura Carreira (eds.), Homologous Recombination: Methods and Protocols, Methods in Molecular Biology, vol. 2153, https://doi.org/10.1007/978-1-0716-0644-5_4, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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cells emerged carrying altered sequences at I-PpoI cleavage sites [2] (see Note 1). Subsequently, a new cleavage site of I-PpoI was inserted in the lys1 locus on chromosome I or the arg1 locus on chromosome III. Synchronous DSB induction is achieved by the addition of anhydrotetracycline (ahTET) to the growth medium. The kinetics of resection is measured by Southern blotting with a series of probes detecting the restriction fragments generated at different distances from the DSB. An additional way to follow resection in fission yeast uses a repairable system where a DSB between two identical and direct lambda DNA repeats (755 bp long) is repaired by single-strand annealing (SSA) [3]. Extensive resection exposes the single-strand repeats that can anneal to each other forming an SSA product. Sequences between the repeats and one of the repeats are lost as a consequence of SSA. The kinetics of SSA product formation is monitored by Southern blotting. To eliminate alternative DSB repair pathways, SSA is followed in the absence of Rad51, which is dispensable for SSA. This SSA assay can only be used to monitor the kinetics of repair, not viability (see Note 2). These are robust and quantitative assays to study the mechanism of DSB end resection in fission yeast. These assays were used to define the function of helicases, nucleases, chromatin remodeling factors, checkpoint proteins, and Rad52 in resection [3]. Materials and methods are nearly the same for the assays designed to study resection kinetics or SSA kinetics except that different DNA probes, different restriction enzymes, and different quantification methods are used.

2

Materials Prepare all solutions using ultrapure water. Prepare and store reagents at room temperature unless indicated otherwise. Experiments using isotopes should follow radiation safety regulations.

2.1 Yeast Strains, Cell Culture, and DSB Induction

1. The genotype of the fission yeast strain used to study the kinetics of DSB end resection is leu1-32::pDUAL-TETp-IPpoI ura4-D18 rDNA-I-Ppolmt ade6-216 lys1::I-PpoI-hph (strain ySK113).The genotype of the fission yeast strain used to study the kinetics of SSA is leu1-32::pDUAL-TETp-I-PpoI ura4-D18 rDNA-I-Ppolmt ade6-216 arg1::λ1-I-PpoI-λ2hph λ2-ura4 (strain yZY003). These strains are available upon request. 2. EMMG medium: for 1 l of liquid EMMG medium, dissolve 28.2 g EMMG powder (United States Biological, E2205-15) and six supplemental chemicals (Adenine, arginine, leucine,

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lysine, histidine, and uracil, 250 mg each) in 1 l of H2O. The medium should be filter sterilized (0.22 μm). 3. EMMG agar plates: add 20 g agar to 1 l EMMG liquid medium, autoclave at 121  C for 20 min, and pour the medium into Petri dishes. 4. YES agar plates: for 1 l of YES agar medium, dissolve 5 g yeast extract, 30 g glucose, and six supplemental chemicals (Adenine, arginine, leucine, lysine, histidine, and uracil, 250 mg each) in 1 l of H2O. Add 20 g agar, autoclave at 121  C for 20 min, and pour the medium into Petri dishes. 5. ahTET: dilute anhydrotetracycline hydrochloride (Acros Organics, AC23313-1000) in H2O at 10 mM concentration. Store 1 ml aliquots frozen at 80  C or  20  C. Cover the tubes with aluminum foil to protect from light. 2.2

DNA Processing

1. Glass disruptor beads, 0.5 mm (Sigma, acid washed). 2. STES buffer: 0.2 M Tris–HCl [pH 7.6], 0.5 M NaCl, 0.1% SDS, 0.01 M EDTA. 3. Phenol/chloroform/isoamyl alcohol (25:24:1, v/v). 4. Chloroform. 5. TE buffer: 10 mM Tris–HCl [pH 8.0], 1 mM EDTA. 6. RNase, 10 mg/ml. 7. NaAc, 3 M. 8. EcoRI and EcoRI buffer from New England Biolabs (NEB). 9. HpaI, StuI, BspHI, and CutSmart buffer from NEB.

2.3 DNA Electrophoresis

1. TBE buffer: For 4 l of 5 TBE buffer, add 242.2 g Tris base, 123.6 g boric acid, and 13.4 g Na2-EDTA to 3 l of H2O. When dissolved completely, bring up to a final volume of 4 l with H2O. Dilute with H2O to 1 TBE before use. 2. 0.8% agarose gel: add 3.2 g of agarose to 200 ml 1 TBE buffer, and microwave for ~3 min. Add another 200 ml 1 TBE buffer and swirl to mix. Let it sit at room temperature (RT) for 5 min to cool. Add 8 μl 1% ethidium bromide and swirl to mix. Pour the liquid into a gel box to cast the gel and leave it at RT for 30 min (small gel) or 60 min (large gel) before use.

2.4

Southern Blotting

1. Denaturing buffer/transfer buffer: 0.4 M NaOH. 2. Filter paper (46  57 cm, Whatman, 3030-917). 3. Blotting pad (20  20 cm, VWR, North American 28298032).

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4. Transfer membrane: positively charged nylon hybridization membrane (PerkinElmer, NEF988001PK). 5. 20% SDS: prepare in a chemical fume hood. For 2 l 20% SDS, add 400 g SDS to 1.5 l of warm H2O (warmed for ~4 min in microwave). When dissolved completely, bring up to a final volume of 2 l with H2O. 6. 20 SSC: for 4 l 20 SSC, add 689.3 g NaCl and 353 g Na-citrate to 3 l of H2O. When dissolved completely, bring up to a final volume of 4 l with H2O. 7. Wash buffer: 2 SSC, 0.2% SDS. 8. Hybridization buffer: for 500 ml hybridization buffer, add 250 ml 0.5 M Na2HPO4 [pH 7.2], 175 ml 20% SDS, 1 ml 0.5 M EDTA [pH 8.0], and 74 ml of H2O. 9. RadPrime DNA labeling system (Invitrogen, 18,428-011). Mix the 0.5 mM dCTP, dGTP, and dTTP together in a 1:1:1 ratio to make the “CGT mix.” 10. [α-32P] dATP, 3000 Ci/mmol 10 mCi/ml (PerkinElmer, BLU512H250UC). 11. Isotope purification column: Micro Bio-Spin chromatography column, P-30 (Bio-Rad, 732-6223). 12. Strip buffer: 1% SDS, 0.1% SSC.

3 3.1

Methods Cell Culture

1. Grow the fission yeast strain at 30  C on YES or EMMG agar plates. 2. Day 1: inoculate cells in 50 ml EMMG medium and culture overnight to saturation on a shaker at 30  C. Day 2: inoculate with 5–10 ml of the saturated culture into 1000 ml EMMG in a 2 l flask and culture overnight on a shaker at 30  C (usually 12–16 h). For mutant strains that grow slower than wild type, inoculate with 10–20 ml of the saturated preculture.

3.2 Induction of DSBs and Collection of Cells

1. Day 3: when the cell density reaches 2–4  106/ml, collect 200 ml of cells and mark as the “time 0” sample. Then add ahTET to the rest of the culture to a final concentration of 4 μM. Turn the light off or cover the flask with aluminum foil to protect the ahTET from degradation (see Note 2). 2. Spin down the cells at >3,500  g for 2 min. Transfer the cell pellet to a 1.5 ml EP tube, spin down at >4000  g for 30 s, remove the remaining liquid, and store the pellets at 20  C. 3. Collect and store the cells at time points indicated in Table 1.

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Table 1 Time and volume of cell culture collected Extensive resection

Time (h) Volume (ml)

0 200

1 180

2 160

4 135

6 120

8 90

Initial resection

Time (min) Volume (ml)

0 200

30 200

60 180

90 160

120 135

150 135

3.3

DNA Extraction

10 80

12 70

– –

– –

1. Add glass beads on top of the cell pellets to a total volume of 500 μl. Add 120 μl TE buffer, 300 μl STES buffer, and 350 μl chloroform/phenol/isoamyl alcohol. 2. Break the cells by vortexing them for 6 min at 4  C (this will break the chromosomes into ~20 kb DNA fragments). Centrifuge the mixture at >16,000  g for 5 min, and then transfer the aqueous top layer (~400 μl) to a new 1.5 ml EP tube. 3. Add 350 μl chloroform/phenol/isoamyl alcohol, mix, and centrifuge the samples at >16,000  g for 5 min, and then transfer the top aqueous layer (~400 μl) to a new 1.5 ml EP tube. 4. Add 350 μl chloroform (do this in a chemical fume hood), mix, and centrifuge the samples at >16,000  g for 5 min, and then transfer the top aqueous layer (~400 μl) to a new 1.5 ml EP tube. 5. Precipitate the DNA by adding 40 μl 3 M NaAC and 1 ml ethanol (100%) and mixing. 6. Centrifuge the mixture at >16,000  g for 20 min. Remove the supernatant by aspiration (avoid disrupting the pellet). Then wash the pellet with 600 μl 70% ethanol, centrifuge for 5 min at >16,000  g, and remove the supernatant by aspiration. Spin down the tube again briefly and remove the remaining ethanol by aspiration. Dry the pellet at RT for 30–60 min. 7. Resuspend the pellet in 300 μl of TE. Add 2 μl RNase (10 mg/ ml) and incubate at 37  C for 2 h up to overnight, and then precipitate the DNA by adding 30 μl NaAC and 900 μl ethanol. Centrifuge and wash as mentioned in step 6. Then resuspend the pellet in 150 μl H2O for ~30 min at RT.

3.4 Restriction Enzyme Digestion and DNA Separation

1. Digest all DNA (150 μl) by adding 30 μl 10x reaction buffer and 2–5 μl restriction enzyme(s) and H2O to a total reaction volume of 300 μl. Digest genomic DNA overnight (16–20 h) at the temperature indicated by the vendor. After digestion, precipitate the DNA by adding and mixing 30 μl NaAc and 900 μl ethanol. Centrifuge and wash as mentioned in Subheading 3.3, step 6. Then resuspend the pellet in 30–40 μl of TE buffer. Restriction enzymes specific for monitoring resection at

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Table 2 Primers for preparing Southern blot probes by PCR Primer name

Restriction enzyme (s)

Sequence

0 kb

EcoRI

50 -CAGAACAGCGGGCAGTTCGG-30 50 -GGTTGTAACGAGACTTGGGGTAAGG-30

3.2 kb

EcoRI

50 -CCATGTTTAGAATGTGAACGAGATGC-30 50 -GGACGTGGCATGAAGAATGTC-30

17.3 kb

HpaI/StuI

50 -GTATACGTTACTTGACATGTGCACACC-30 50 -CGAGCTAAAATACTTAGTGCCTCTGGAC-30

27.0 kb

HpaI/StuI

50 -CCTACGCAACTGAGTCTAATAAGTC-30 50 -CCTTTGCTCCAGTTTGTTAAGC-30

lys1 load control HpaI/StuI

HpaI/StuI

50 -CGTGAGGTTGCTAGGAGAATCAAACTTC-30 50 -CTCTTCAAGCTCACCACTCCCG-30

lys1 load control EcoRI

EcoRI

50 -GGACTTCAAGGGCCCGTTAAC-30 50 -CGTGGCAGCTTATAGTTGTGAAGGAG-30

SSA lambda

BspHI

50 -CATTGAGCAGTGCAGCGAAC-30 50 -CCCGCCAGATGATAAGCATC-30

SSA load control

BspHI

Same as lys1_load_control_HpaI/StuI

nonrepairable breaks and for monitoring SSA kinetics are specified in Table 2. 2. Check the quality and completeness of DNA digestion by loading and separating 2 μl of each sample in a small 0.8% agarose gel for ~1 h at 80–120 V. 3. Take a picture of the gel using the UV gel imaging system to make sure that the DNA is fully digested. Measure the amount of DNA in each sample using a fluorometer (e.g., Qubit) and equalize the amount of DNA in samples with TE buffer. 4. Cast a large 0.8% agarose gel in 1 TBE buffer with a comb appropriate for the sample number and volume. For a large gel box, a 300 ml gel is used. 5. Add 4 μl DNA gel loading dye (6) to 20 μl DNA, mix, and centrifuge for a few seconds. Load 18–24 μl of each sample. 6. Run the DNA gel at 47–60 V for 14–19 h until the DNA fragments between 2 kb and 10 kb are well separated. 7. Take a picture of the gel using a UV gel-imaging system to make sure the samples are well equalized and separated. 3.5 Alkaline Gel Transfer

1. Cut the DNA gel into the size of the blotting paper (20  20 cm) and place in a glass tray on an orbital shaker at ~30 rpm. Denature the DNA with denaturing buffer for 20 min (buffer should cover the gel), and then replace with

Analysis of Resection in Fission Yeast (9)

Weight (6)

(5)

(4) (3)

(1)

(4) (7)

(7)

(8) (9)

(2)

(7)

(5)

(8)

53

(7) (7)

(7) (8) Front view

Top view

Fig. 1 Scheme of the Southern blot “transfer sandwich.” (1) Two packs of paper towels. (2) Six layers of blotting pads, 20  20 cm. (3) Two layers of Whatman filter paper, 20  20 cm. (4) Positively charged membrane. (5) Agarose gel with DNA. (6) Two layers of Whatman filter paper cut into the size of the gel. (7) Four cups with denature/transfer buffer, ~250 ml per cup. (8) Two strips of bridge filter papers, 46  4 cm. (9) One layer of Whatman filter paper, 16  18 cm

new denaturing buffer, and continue to shake for another 20 min. 2. Cut the filter paper and the transfer membrane to the size of the gel. Cut four filter papers to 20  20 cm, four filter paper strips to 46  4 cm, and one filter paper to 16  18 cm. Cut the transfer membrane to 21  21 cm (1 cm longer and 1 cm wider than the gel). Mark the corner of the membrane with the date and experiment number. 3. Transfer the gel to the transfer membrane by assembling a “transfer sandwich” as shown in Fig. 1 (also known as a capillary blotting apparatus). In detail, place two packs of paper towels on the table top (5–10 cm high for each pack), and then place 5–6 blotting pads on top of the paper towels. Wet two layers of Whatman filter paper (20  20 cm) and place on top of the blotting pad avoiding any bubbles. Wet the transfer membrane and put it beneath the gel. Hold the bottom of the membrane and put the membrane and gel together on top of the filter/blotting paper stack. Wet two other filter papers and put them on top of the gel. Roll out all the bubbles between each layer of the “sandwich” using a long plastic pipette. Put two bi-layer “buffer bridge” wide strips on top of the stack, and put the two ends of each bridge into the transfer buffer (the bridge acts as a wick for buffer transfer). Put another filter paper (16  18 cm) on top of the bridges to connect them together. Cover the stack with a piece of plastic wrap and put a weight (about 0.5 kg) on top of the stack. Cover the transfer sandwich with plastic wrap to prevent buffer evaporation. 4. Let the transfer of DNA to the membrane occur for ~20 h. Disassemble the “transfer sandwich,” place the membrane in a

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clean glass tray, and wash the membrane with wash buffer for 20 min on an orbital shaker at ~30 rpm. Dry the membrane on a paper towel at RT for at least 20 min. 3.6 Probe Labeling and Hybridization

1. DNA probes (500–800 bp) for Southern blotting are prepared by PCR, purified by regular gel extraction (NucleoSpin Gel and PCR Clean-up, MACHEREY-NAGEL, 740609.250) and diluted to a concentration of ~50 ng/μl. The probes next to the DSB are used to test initial DSB end resection and the probes at over 10 kb away from the DSB are used to test extensive resection. The DNA probes specific for monitoring resection at nonrepairable breaks and for monitoring the kinetics of SSA are specified in Table 2. 2. For one Southern blot, add 30–50 ng of the desired probe in a 1.5 ml EP tube, and H2O to a total volume of 20 μl. Denature the DNA probe in a 100  C Dri-Bath block for 5 min (hold the cap by a CapLock tube clip to keep the cap closed), and then cool it down on ice for 3 min. 3. Add 20 μl 2.5 RadPrime buffer (Invitrogen), 3 μl CGT mix, 1 μl Klenow Fragment, and 3 μl [α-32P] dATP to the tube, mix gently, and centrifuge for 15 s. Incubate at 37  C for 0.5–2 h. 4. Purify the probe using a Micro Bio-Spin chromatography column according to the manufacturer’s instructions (Bio-Rad). Heat denature the purified sample at 100  C for 5 min and cool it down on ice for 5 min. 5. Put the membrane in the hybridization tube with 50 ml hybridization buffer. Put the hybridization tube in the hybridization incubator. Remove any bubbles between the membrane and hybridization glass tube. Set the temperature to 65  C. Incubate for at least 30 min for prehybridization. 6. Pipette ~40 μl of the purified probe mix into the hybridization tube and hybridize overnight. 7. Discard the hybridization buffer, add 50 ml wash buffer, and let it rotate in the hybridization incubator for 30 min. Repeat the washing two additional times. 8. Remove the membrane from the hybridization tube, place on a paper towel to remove the remaining buffer, and quickly wrap with plastic wrap (do not let it dry). Place the membrane face up inside the exposure cassette and put the phosphor screen on top of the membrane, phosphor side down. Close the cassette and leave it at room temperature for several hours up to overnight depending on the strength of the signal. (Note: Bleach the phosphor screen on a white light transilluminator for 30 min every time before use.) 9. Put the phosphor screen on the glass surface of the Typhoon Trio scanner. Align the screen to the top left.

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10. Open the Typhoon Trio Imager software, select the region to be scanned, and follow the instructions to scan the screen at the desired resolution. After scanning, the wrapped blots can be stored at 4  C for a few months. 11. To reprobe the same membrane remove the plastic wrap and put the blot in a plastic tray. Heat 400 ml strip buffer in the microwave for 3 min and pour the hot buffer on top of the blot. Gently shake the tray for 3 min then discard the strip buffer. Repeat the wash three times to remove the hybridized probe. Dry the blot by leaving it on paper towels at RT. Rehybridize the blot with additional probes as needed. 3.7

Data Analysis

1. Use ImageQuant TL (GE Healthcare) or ImageJ (NIH) software to quantify the intensity of the bands on a Southern blot. Export the band intensity values to Excel for analysis.

time (min) 0 parental

Wild-type cells 30 60 90 120 150

I-PpoI cut I-PpoI cut 3.2 kb load control

b.

0 kb from DSB 100 80 60 40 20 0 30

60

90

120 150

% of unresected DSB ends

a.

% of unresected DSB ends

2. For resection kinetics, normalize the intensity of the bands by the intensity of the loading control. Then divide the normalized intensity of each band by the normalized band intensity at time 0 to obtain the percentage of unresected DSB ends (Fig. 2). For single-strand annealing, normalize the intensity

3.2 kb from DSB 100 80 60 40 20 0 0

c. time (hr) 0 parental I-PpoI cut 17.3 kb 27.0 kb load control

1

Wild-type cells 2 4 6 8

d. 10 12

% of unresected DSB ends

time (min)

30 60 90 120 150 time (min)

100

17.3 kb 27.0 kb

80 60 40 20 0 0

2

4

6

8 10 12

time (hr)

Fig. 2 Analysis of resection by Southern blots and the resection kinetics. (a) Southern blots showing resection at DSB ends and 3.2 kb from DSB ends. Loading control is shown. (b) Plots showing initial resection kinetics. The 0 kb plot shows the relative band signal intensities compared to the band at 30 min. The 3.2 kb plot shows the relative band intensities compared to the band at 0 min. (c) Southern blot showing extensive resection at 17 and 27 kb from DSB ends. (d) Plots showing extensive resection kinetics. The plots show the relative band intensities compared to the band at 0 h. Band signal intensities were normalized by the intensities of the loading control bands. Error bars denote standard deviation (n ¼ 3)

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a.

b.

DSB 21 kb Chr III λ2 BspHI

ura4

λ1

EG

hygR

λ2

BspHI

BspHI

BspHI

6.8 kb

1.6 kb SSA product

2

rad51∆ 3 4

5

6

7

λ2 (6.8 kb)

BspHI

3.2 kb % of SSA product formation

1

λ2 hygR BspHI

c.

time (hr) 0

SSA product (3.2 kb)

80 rad51∆ cells

I-PpoI cut

60 40 λ2 (1.6 kb)

20

load control

0 0

1

2

3

4

5

6

7

time (hr)

Fig. 3 SSA assay. (a) Schematic of the SSA assay between λ sequences. λ DNA sequence harboring a I-PpoI cleavage site in the middle was inserted at the arg1 locus on chromosome III and an identical λ sequence was inserted 21 kb upstream of the I-PpoI cleavage site. Boxes marked “EG” represent two essential genes. (b) Southern blot analysis of the kinetics of SSA product formation in rad51Δ cells. (c) Plots showing SSA kinetics. The intensity of each band was normalized to the loading control

of the SSA product band by the intensity of the loading control. Then divide the normalized intensity of the SSA product band by the normalized intensity of the 1.6 kb parental band at time 0 to obtain the percentage of SSA product formation (Fig. 3).

4

Notes 1. A drawback of this system is that each of the rDNA sequences contains a cleavage site of I-PpoI, so only the I-PpoI-resistant rDNA mutation strains can be used. Other fission yeast systems using HO endonuclease or the I-SceI endonuclease to induce site-specific recombination were developed [4, 5]. However, I-PpoI provides the most synchronous induction of DSBs. 2. I-PpoI is leaky, leading to some DSB induction even before the addition of anhydrotetracycline to the media. To eliminate cells carrying the SSA assay that induced DSBs prior to anhydrotetracycline addition, SSA was designed to have two essential genes between the lambda repeats. Thus, cells that induced I-PpoI enzyme and SSA repair before anhydrotetracycline addition die and are eliminated from the population due to loss of essential genes (Fig. 2).

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Acknowledgements This work was supported by NIH grants GM080600 and GM125650. References 1. Symington LS, Gautier J (2012) Double-strand break end resection and repair pathway choice. Annu Rev Genet 45:247–271 2. Sunder S et al (2012) A new method to efficiently induce a site-specific double-strand break in the fission yeast Schizosaccharomyces pombe. Yeast 29(7):275–291 3. Yan Z et al (2019) Rad52 restrains resection at DNA double-strand break ends in yeast. Mol Cell 76(5):699–711

4. Wang J et al (2018) A heterochromatin domain forms gradually at a new telomere and is dynamic at stable telomeres. Mol Cell Biol 38 (15):e00393–e00317 5. Watson AT, Werler P, Carr AM (2011) Regulation of gene expression at the fission yeast Schizosaccharomyces pombe urg1 locus. Gene 484 (1–2):75–85

Chapter 5 Quantifying DNA End Resection in Human Cells Yi Zhou and Tanya T. Paull Abstract DNA double-strand break (DSB) end resection initiates homologous recombination (HR) and is critical for genomic stability. DSB resection has been monitored indirectly in mammalian cells using detection of protein foci or BrdU foci formation, which is dependent on single-stranded DNA (ssDNA) products of resection. Here we describe a quantitative PCR (qPCR)-based assay to directly measure levels of ssDNA intermediates generated by resection at specific DSB sites in human cells, which is more quantitative and precise with respect to the extent and efficiency of resection compared with previous methods. This assay, excluding the time for making the stable cell line expressing the restriction enzyme AsiSI fused to the estrogen receptor hormone-binding domain (ER-AsiSI), can be completed within 3 days. Key words DNA repair, DNA end resection, DNA damage, Single-stranded DNA quantitation

1

Introduction DNA double-strand breaks (DSBs) are one of the most deleterious DNA lesions in eukaryotic cells and are caused by a variety of external and internal factors including ionizing radiation, chemotherapeutic drugs, reactive oxygen products of metabolism, and replication fork collapse. Upon induction of DSBs, DNA repair is rapidly initiated along with cell cycle checkpoint arrest. If not repaired correctly, DSBs can lead to chromosome rearrangements, genomic instability, and tumorigenesis [1]. Eukaryotic cells have developed two major pathways to repair DSBs: nonhomologous end joining (NHEJ) and homologous recombination (HR). NHEJ is the predominant repair pathway for DSBs and is utilized throughout the cell cycle while the activity of HR is limited to S and G2 phases where sister chromatids are available as repair templates and cyclin-dependent kinase (CDK) is active [2]. NHEJ is “error prone” because it is often associated with short range of end processing before rejoining of the DNA ends. In contrast, HR is relatively “error free” since the break is repaired using homologous DNA as the template. HR is initiated with the

Andre´s Aguilera and Aura Carreira (eds.), Homologous Recombination: Methods and Protocols, Methods in Molecular Biology, vol. 2153, https://doi.org/10.1007/978-1-0716-0644-5_5, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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resection of the 50 strands to generate 30 single-stranded DNA (ssDNA), which is required for Rad51 binding and strand invasion. The ssDNA intermediates of resection serve to inhibit NHEJ but are required for Rad51 filament formation and strand invasion during HR [3, 4]. Therefore, the initiation of resection is thought to be a critical control point for the choice between HR and NHEJ, which has potentially important consequences for genome stability since NHEJ is error prone and also implicated in the formation of chromosomal translocations that can be critical events in cancer cells [5]. DSB resection in mammalian cells is often assessed indirectly by monitoring the formation of RPA foci, Rad51 foci, or BrdU foci. Although these methods can provide a global estimate for the level of resection, they are subjectively dependent on the immunofluorescence protocol and antibody used and cannot determine the length of resection from a DSB site. Previously, we developed a quantitative PCR (qPCR)-based assay to directly measure levels of ssDNA intermediates generated by resection at specific DSB sites in human cells [6]. This assay is more quantitative and more precise with respect to the extent and efficiency of resection compared with existing foci-based methods. Using this assay, we have quantitatively investigated the roles of several known DNA repair factors, including CtIP, Mre11, SOSS1, Exo1, Ku, DNA-PKcs, 53BP1, and BRCA1, in DSB end resection in human cells, which has demonstrated the validity of our method [6]. In several other studies, we have examined how DNA-PKcs kinase activity, ATM kinase activity, CtIP nuclease activity, and RPA/Dna2 affect DSB end resection in human cells by taking advantage of this assay [7– 11]. Here we provide a detailed description of the procedure for the assay. We utilize the ER-AsiSI system [12] to induce DSBs at specific sites of genomic DNA. In this system, the restriction enzyme AsiSI is fused to the estrogen receptor hormone-binding domain and can be induced to enter the nucleus where it generates DSBs at sequence-specific sites (50 -GCGATCGC-30 ) upon 4-hydroxytamoxifen (4-OHT) treatment [12]. There are about 1000 AsiSI restriction sites in the human genome although not every site is cleaved upon 4-OHT exposure and nuclear translocation of AsiSI. It has been shown that approximately 150 DSBs can be actually induced by the AsiSI enzyme in human cells [13]. Since the ER-AsiSI cassette is constructed in a retroviral vector containing a puromycin selection marker [12], the ER-AsiSI system can be efficiently introduced into target cells by retrovirus infection and puromycin selection [6]. After induction of DSBs in target cells, the genomic DNA is extracted using a method in which cells are embedded in low-gelling point agar that protects the DNA from shearing and damage during extraction. This method is modified from a previously described strategy [14], which involves

Quantitation of DNA Resection Intermediates in Human Cells Restriction site

5’ primer

DNA without end resection

3’ primer

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Taqman probe

Restriction digestion qPCR

No PCR products

DNA with end resection

Restriction digestion qPCR PCR products

Fig. 1 The principle for quantitation of DNA end resection in human cells. To measure the amount of ssDNA, select a restriction site in the interested DNA region and digest or mock digest the DNA sample using the corresponding restriction enzyme, followed by qPCR analysis of the digested DNA sample using primers across the restriction site. If the DNA remains double stranded at the interested region (upper panel), it will be cut by the restriction enzyme and there would be no PCR amplification. In contrast, if the interested DNA region becomes single stranded due to resection (bottom panel), it will not be cut by the restriction enzyme, and thus one strand of DNA remains intact, which will generate a lot of PCR products. For each sample, a ΔCt is calculated by subtracting the Ct value of the mock-digested sample from the Ct value of the digested sample.  The percentage  of ssDNA is calculated using this equation: ssDNA% ¼ 1= 2ðΔC t1 Þ þ 0:5  100

(1) embedding the cells in agar; (2) lysing the cells by ESP buffer containing proteinase K and detergent; (3) stripping the genomic DNA from histones and other proteins by a high salt (HS) buffer; (4) washing the agar adequately with phosphate buffer; and (5) melting the agar containing genomic DNA and diluting the sample with ddH2O to prevent it from gelling again. Almost all restriction enzymes cut double-stranded DNA (dsDNA) but not single-stranded DNA (ssDNA). We exploit this feature of restriction enzymes to distinguish original dsDNA from ssDNA products of resection at specific sites (Fig. 1). To quantitate ssDNA generated by resection at specific DSBs, we choose two AsiSI sites on Chromosome 1 (“DSB1,” Chr 1: 89231183; “DSB2,” Chr 1: 109838221) that have been shown to be cleaved with high efficiency [15]. We design three pairs of qPCR primers across BsrGI or BamHI restriction sites, which are used to distinguish between ssDNA and dsDNA, at various distances from each AsiSI site. In addition, we design another two pairs of primers across the two AsiSI sites to monitor the percentage of double-

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DSB I AsiSI

BsrGI

BsrGI

BsrGI

(Chr 1: 89231183)

(335 nt)

(1618 nt)

(3500 nt)

DSB II BamHI

BamHI

BamHI

(Chr 1: 109838221) (364 nt)

(1754 nt)

(3564 nt)

AsiSI

No DSB HindIII (Chr. 22: No AsiSI site within 2 Mb)

Fig. 2 Design of Taqman qPCR primers and probes for quantitation of ssDNA intermediates of resection. Schematic diagram of Taqman qPCR primers and probe designs for measurement of DSB% at two selected AsiSI sites (“DSB1” and “DSB2”) located on Chromosome 1 and measurement of resection at sites adjacent to the two AsiSI sites. The primer pairs for measurement of DSB% at two selected AsiSI sites are marked by blue arrows. The primer pair (red arrows) on Chromosome 22 (“No DSB”), which is designed across a HindIII restriction site, is used as negative control. The primer pairs for measurement of resection at “DSB1” and “DSB2” are across BsrGI and BamHI restriction sites and are marked by orange arrows (335 nt, 1618 nt, and 3500 nt from the AsiSI site) and purple arrows (364 nt, 1754 nt, and 3564 nt from the AsiSI site), respectively. All Taqman probes, which are not shown in the diagram, contain a 50 reporter group 6FAM and a 30 quencher group TAMRA and are designed at either side of the restriction site

strand breaks (DSB%) present at the two sites as well as a pair of primers at a site where there is no nearby AsiSI sequence on Chromosome 22 (“No DSB”) which is used as a negative control (Fig. 2). The percentage of ssDNA generated by resection at various sites is measured by qPCR as previously described [6]. Briefly, for each sample, a ΔCt was calculated by subtracting the Ct value of the mock-digested sample from the Ct value of the digested sample. The percentage of ssDNA was calculated with the following equa  tion: ssDNA% ¼ 1= 2ðΔC t1 Þ þ 0:5  100 [16]. It is notable that the accumulation of DSBs at selected AsiSI sites may be affected by various treatments such as siRNA-mediated depletion of proteins [6] and treatment with small-molecular inhibitors [17]. Thus, it is essential to measure the percentage of double-strand breaks (DSB %) present at selected AsiSI sites in order to accurately interpret the effects of these treatments on resection. We have used the assay in the context of site-specific DSBs in order to measure end resection, but in theory this method can be used to quantitate any ssDNA intermediates in any genomic context. Direct measurement of ssDNA intermediates of resection has obvious advantages over foci-based methods in that quantitative analysis of the actual resection products can be performed with respect to the extent and efficiency of resection. However, this

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assay is limited by the number of DSBs that can be actually studied and the requirement for sequence specificity of the cut site. Other methods such as RPA ChIP-Seq have been utilized to address the need for resection assays at random or unknown DNA damage sites in the mammalian genome [18, 19]. A combination of these methods may be necessary to quantitatively measure ssDNA at nuclease accessible as well as inaccessible sites.

2

Materials 1. Agarose (0.6% (wt/vol) in PBS) solution: Mix 0.15 g agarose powder with 25 ml PBS in a 100 ml glass conical flask, heat the solution for 40 s in a microwave to completely dissolve the agarose, and place the flask in a 37  C shaking incubator until use (see Note 1). 2. 50 mg/ml proteinase K: Take a vial of proteinase K (100 mg), add 2 ml ddH2O, mix carefully, and make 200 μl aliquots. Store the solution at 20  C (see Note 2). 3. 0.5 M EDTA, pH 8.0: Mix 93.1 g EDTA with 400 ml ddH2O, add NaOH pellet to the solution while stirring vigorously on a magnetic stirrer until pH 8.0. Adjust the volume to 500 ml with ddH2O. 4. 1 M CaCl2: Dissolve 1.1 g CaCl2 in 10 ml ddH2O. Store the solution at 4  C. 5. ESP buffer: Dissolve 10 g N-lauroylsarcosine in 500 ml 0.5 M EDTA solution; adjust the pH to 8.0. Filter the solution and store it at 4  C. Before use, mix appropriate amount of this solution with proteinase K stock and CaCl2 stock to get a final concentration of 1 mg/ml for proteinase K and 1 mM for CaCl2 (see Note 3). 6. HS buffer: Prepare 500 ml solution containing 1.85 M NaCl, 0.15 M KCl, 5 mM MgCl2, 2 mM EDTA, and 4 mM Tris; adjust the pH to 7.5. Filter the solution and store it at 4  C. Before use, mix appropriate amount of this solution with Triton X-100 to get a final concentration of 0.5% (wt/vol) for Triton X-100 (see Note 4). 7. Phosphate buffer: Prepare 1 l phosphate buffer containing 8 mM Na2HPO4, 1.5 mM KH2PO4, 133 mM KCl, and 0.8 mM MgCl2; adjust the pH to 7.4. Filter and store the solution at 4  C. 8. Plasmid extraction: Plasmids used in this protocol are extracted using plasmid miniprep kit following the manufacturer’s instructions. Plasmid concentration is measured by NanoDrop 2000.

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Methods

3.1 HA-ER-AsiSI Retrovirus or AID-ER-AsiSI Lentivirus Packing

1. Seed 293T cells in a 6-well plate at a density of 6–8  105 cells/ well the day before transfection (see Note 5). 2. When cell confluency is about 70–80%, transfect 2 μg retroviral vector pBABE-HA-ER-AsiSI together with two helper plasmids (1.8 μg CSZ-MGP and 0.2 μg pMD2G) into 293T cells in each well of the 6-well plate using Lipofectamine 2000 following the manufacturer’s instructions (see Note 6). Alternatively, we use the lentivirus construct pTP3991, containing AID-AsiSI-ER, derived from the degron-containing construct pAID-AsiSI [20] with the helper plasmids Δ8.9 and VsVg. 3. 24 h after transfection, split the cells in each well of the 6-well plate into a 25 cm2 tissue culture flask containing 4 ml medium. 4. 48 h later, harvest the culture medium containing HA-ERAsiSI retrovirus; immediately add 4 ml fresh warm medium to the flask without disturbing the cell layer (see Note 7). 5. Filter the harvested medium with a 0.45 μm syringe filter. If there are a lot of floating dead cells in the culture medium, use more than one filter for each flask. 6. Aliquot and freeze the medium with liquid nitrogen and store it at 80  C for future use (see Note 8). 7. Collect and freeze the culture medium again 24 h later as in steps 4–6.

3.2 Infection of Target Cells

1. To infect target cells, grow the cells in 25 cm2 tissue culture flask to ~50% confluency and infect the cells with 4 ml viruscontaining culture medium supplemented with 0.1% polybrene overnight (see Note 9). 2. Replace the medium with fresh medium the next day.

3.3 Selection of Target Cells

1. 48 h after infection, begin to select the cells with 2 μg/ml puromycin for 2–3 weeks to generate a stable cell line. Change the medium every 3 days. 2. Verify the expression of HA-ER-AsiSI in the survival cells by Western blotting using anti-HA antibody.

3.4 Induction of DSBs at Specific Sites in Target Cells

1. Induce nucleus translocation of AsiSI by adding 4-OHT. 2. (Optional) Verify the induction of DSBs by AsiSI enzyme 4 h after treating the cells with 4-OHT by observing γH2AX foci formation through immunofluorescent staining.

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3.5 Genomic DNA Extraction

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1. 1–2 h before the cells are ready for genomic DNA extraction, dissolve 0.15 g low-gelling point agarose powder (BD Biosciences) in 25 ml PBS (Gibco) by microwaving for 40 s to make a 0.6% agarose solution (see Note 10). 2. Place the flask with the agarose solution in a shaking (~200 rpm) 37  C incubator to cool down the solution and keep it at 37  C until use (see Note 11). 3. Trypsinize the cells with 0.25% trypsin-EDTA, spin down the cells at 500  g for 4 min, and resuspend the cells with 37  C agarose solution at a density of 1.5–6.0  106 cells/ml (do a titration for different cell types to determine the optimal density) (see Note 12). 4. Drop 50 μl cell suspension on a piece of Parafilm to generate a solidified agar ball at 25  C (room temperature). The agar ball looks a little turbid as there are a lot of cells in it (see Note 13). 5. Transfer the agar ball to a 1.5 ml centrifuge tube carefully. 6. Treat the agar ball with 1 ml ESP buffer for 20 h at 16  C with rotation (20 rotations/min) in an incubator rotator (see Note 14). 7. Carefully aspirate ESP buffer from the tube without puncturing the agar ball. The agar ball turns completely clear and is hard to be observed (see Note 15). 8. Treat the agar ball with 1 ml HS buffer for 20 h at 16  C with rotation as in step 19 (see Note 16). 9. Carefully aspirate HS buffer from the tube without puncturing the agar ball. 10. Wash the agar ball with 1 ml cold phosphate buffer for 1 h at 4  C with rotation (40 rotations/min) in the incubator rotator (see Note 17). 11. Carefully aspirate phosphate buffer from the tube without puncturing the agar ball. 12. Repeat steps 23 and 24 for five more times. 13. Carefully aspirate phosphate buffer from the tube without puncturing the agar ball. Try to aspirate ALL of the buffer. 14. Melt the agar ball by placing the tube (lid closed) in a 70  C heat block for 10 min. Flick the tube a few times after the gel is melted to get the solution to the bottom of the tube. 15. Dilute the solution 15-fold with 70  C ddH2O, mix it well and cool it down to room temperature (see Note 18). 16. Mix the solution with equal volume of appropriate 2 NEB restriction enzyme buffer. The gDNA sample can be used immediately or stored at 4  C for future use. Choose the buffer corresponding to the restriction enzyme for gDNA digestion in the next step (see Note 19).

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3.6 Restriction Enzyme Digestion

1. Digest or mock digest 20 μl genomic DNA sample (~140 ng in appropriate 1 NEB restriction enzyme buffer) with 20 units of selected restriction enzyme at 37  C overnight (see Note 20).

3.7 Quantitation of ssDNA Intermediate of Resection by qPCR

1. Use 3 μl digested or mock-digested gDNA sample as a template in a 25 μl qPCR reaction containing 12.5 μl 2 Taqman Universal PCR Master Mix, 0.5 μM of each primer, and 0.2 μM probe (see Note 21). Each 96 well contains: (a) 0.5 μl 50 primer (25 μM). (b) 0.5 μl 30 primer (25 μM). (c) 0.5 μl probe (10 μM). (d) 12.5 μl 2 Taqman Universal PCR Master Mix (ABI). (e) 8 μl ddH2O. (f) 3 μl digested or mock-digested gDNA sample. The PCR program is as follows (1.6  C/s ramp rate): Hold stage: (a) 50  C 2 min. (b) 95  C 10 min. PCR stage: 40–45 cycles (a) 95  C 15 s. (b) 60  C 1 min (signal recording step). 2. Calculate the percentage of ssDNA generated by resection at selected sites. For each sample, a ΔCt is calculated by subtracting the Ct value of the mock-digested sample from the Ct value of the digested sample. The percentage of ssDNA is calculated with the equation: ssDNA% ¼   following 1= 2ðΔC t1 Þ þ 0:5  100.

3.8 Measurement of % DSB at Selected AsiSI Site

4

1. Measure the amount of DSBs present at the selected AsiSI site using a pair of primers across this site. Use the “No DSB” primers to normalize the amount of gDNA in the qPCR reaction. DSB percentage in 4-OHT mock-treated cells is set to zero.

Notes 1. The flask should be capped to prevent the evaporation of water. We favor to heat the solution for 30 s, swirl the flask for a few seconds, and then heat the solution for another 10 s. Just prepare the agarose solution 1–2 h before the cells are ready for genomic DNA extraction. Make sure the temperature of the solution is not higher than 37  C when used.

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2. Avoid thawing and freezing the solution for multiple times as this will decrease the activity of proteinase K. 3. N-lauroylsarcosine powder is very irritating to the respiratory system and eyes. Working with appropriate protective equipment is recommended. Proteinase K should be added to the ESP buffer right before use. 4. Triton X-100 should be added to the HS buffer right before use. Triton X-100 is quite viscous and should be pipetted very slowly in order to get the accurate amount. 5. The growing status of 293T cells is critical for virus packing; avoid using 293T cells that are too old or look unhealthy. 6. The cells should not be over-confluent; otherwise, they will come off easily during transfection and virus harvest. Replacing the medium with 2 ml fresh 37  C medium without antibiotics before the transfection is essential. 7. The culture medium looks bright yellow. It does not matter if the cells are not good looking because the retrovirus is harmful to the cells. 8. Store the supernatant in small aliquots to avoid repetitive freeze-thawing that will reduce the viability of the virus. 9. Culture medium collected at 48 h after splitting the 293T cells generally contains more virus than culture medium collected at 72 h and is more efficient for infecting target cells. 10. The flask in which the agarose solution is made should be capped to prevent the evaporation and spill of water during heating. Make sure the agarose powder has low-gelling point as regular agarose solution may be gelling at 37  C. 11. Shaking the flask containing agarose solution in a 37  C incubator shaker is the best way to keep the solution at 37  C and prevent it from gelling. We favor to put a stir bar in the solution. 12. Make sure the cells are fully trypsinized in order to obtain a single-cell suspension in agarose solution, which is important for good extraction of genomic DNA. Make sure the temperature of the agarose solution is not higher than 37  C. 13. This step should be performed quickly as the agarose solution undergoes gelling fast. Avoid getting a lot of air bubbles into the agar ball. It does not matter if the agar ball is flat. 14. Steps 16–19 should be performed as fast as possible to avoid repair of AsiSI-induced breaks. Make sure that the agar ball is floating around in the buffer rather than being attached to the wall or the cap of the tube.

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15. A good way to avoid puncturing the agar ball is to invert the tubes for a few times, making sure the agar ball is not attached to the bottom, followed by inserting the pipette tip to the bottom along the tube wall and aspirating the buffer carefully. 16. The agar ball can be left in the HS buffer for another 20 h before proceeding to the next step. 17. Note that the temperature is turned to 4  C to prevent the DNA from degradation as there is no EDTA in the phosphate buffer. 18. It is important to preheat the water to 70  C in order to prevent the agarose solution from gelling. 19. To avoid shearing the gDNA, mix the solution gently with a pipette. Do not vortex the solution. The gDNA samples may be stored at 4  C for up to 1 month. 20. The genomic DNA should be digested for at least 12 h. We favor performing the digestion in a 37  C incubator to avoid evaporation and accumulation of water on the lids. The digested gDNA samples may be stored at 4  C for up to 1 week before proceeding to qPCR analysis. 21. Prepare the qPCR mixture as fast as possible. We favor making a mixture containing Taqman Master Mix, primers and probe first, and then add 22 μl of the mixture to the assigned wells, followed by the addition of 3 μl digested or mock-digested gDNA sample to each well.

Acknowledgments Work in the Paull laboratory is supported by the Cancer Prevention and Research Institute of Texas grant RP110465-P4. References 1. Ciccia A, Elledge SJ (2010) The DNA damage response: making it safe to play with knives. Mol Cell 40:179–204. https://doi.org/10. 1016/j.molcel.2010.09.019 2. Trovesi C, Manfrini N, Falcettoni M, Longhese MP (2013) Regulation of the DNA damage response by Cyclin-dependent kinases. J Mol Biol 425(23):4756–4766. https://doi.org/ 10.1016/j.jmb.2013.04.013 3. Huertas P (2010) DNA resection in eukaryotes: deciding how to fix the break. Nat Struct Mol Biol 17:11–16. https://doi.org/10. 1038/nsmb.1710 4. Symington LS, Gautier J (2011) Doublestrand break end resection and repair pathway

choice. Annu Rev Genet 45:247–271. https:// doi.org/10.1146/annurev-genet-110410132435 5. Lieber MR, Gu J, Lu H, Shimazaki N, Tsai AG (2010) Nonhomologous DNA end joining (NHEJ) and chromosomal translocations in humans. Subcell Biochem 50:279–296. https://doi.org/10.1007/978-90-481-34717_14 6. Zhou Y, Caron P, Legube G, Paull TT (2013) Quantitation of DNA double-strand break resection intermediates in human cells. Nucleic Acids Res 42(3):e19. https://doi.org/10. 1093/nar/gkt1309

Quantitation of DNA Resection Intermediates in Human Cells 7. Makharashvili N, Tubbs AT, Yang SH, Wang H, Barton O, Zhou Y, Deshpande RA, Lee JH, Lobrich M, Sleckman BP, Wu X, Paull TT (2014) Catalytic and noncatalytic roles of the CtIP endonuclease in double-strand break end resection. Mol Cell 54:1022–1033. https://doi.org/10.1016/j.molcel.2014.04. 011 8. Zhou Y, Lee JH, Jiang W, Crowe JL, Zha S, Paull TT (2017) Regulation of the DNA damage response by DNA-PKcs inhibitory phosphorylation of ATM. Mol Cell 65:91–104. https://doi.org/10.1016/j.molcel.2016.11. 004 9. Lee JH, Mand MR, Kao CH, Zhou Y, Ryu SW, Richards AL, Coon JJ, Paull TT (2018) ATM directs DNA damage responses and proteostasis via genetically separable pathways. Sci Signal 11:512. https://doi.org/10.1126/scisignal. aan5598 10. Myler LR, Gallardo IF, Zhou Y, Gong F, Yang SH, Wold MS, Miller KM, Paull TT, Finkelstein IJ (2016) Single-molecule imaging reveals the mechanism of Exo1 regulation by single-stranded DNA binding proteins. Proc Natl Acad Sci U S A 113:E1170–E1179. https://doi.org/10.1073/pnas.1516674113 11. Zhou Y, Paull TT (2013) DNA-dependent protein kinase regulates DNA end resection in concert with Mre11-Rad50-Nbs1 (MRN) and ataxia telangiectasia-mutated (ATM). J Biol Chem 288:37112–37125. https://doi.org/ 10.1074/jbc.M113.514398 12. Iacovoni JS, Caron P, Lassadi I, Nicolas E, Massip L, Trouche D, Legube G (2010) High-resolution profiling of gammaH2AX around DNA double strand breaks in the mammalian genome. EMBO J 29:1446–1457. https://doi.org/10.1038/emboj.2010.38 13. Massip L, Caron P, Iacovoni JS, Trouche D, Legube G (2010) Deciphering the chromatin landscape induced around DNA double strand breaks. Cell Cycle 9:2963–2972. https://doi. org/10.4161/cc.9.15.12412 14. Stenerlow B, Karlsson KH, Cooper B, Rydberg B (2003) Measurement of prompt DNA

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double-strand breaks in mammalian cells without including heat-labile sites: results for cells deficient in nonhomologous end joining. Radiat Res 159:502–510 15. Miller KM, Tjeertes JV, Coates J, Legube G, Polo SE, Britton S, Jackson SP (2010) Human HDAC1 and HDAC2 function in the DNA-damage response to promote DNA nonhomologous end-joining. Nat Struct Mol Biol 17:1144–1151. https://doi.org/10.1038/ nsmb.1899 16. Zierhut C, Diffley JF (2008) Break dosage, cell cycle stage and DNA replication influence DNA double strand break response. EMBO J 27:1875–1885. https://doi.org/10.1038/ emboj.2008.111 17. Zhou Y, Paull TT (2013) DNA-dependent protein kinase regulates DNA end resection in concert with the Mre11-Rad50-Nbs1 (MRN) complex and ataxia-telangiectasia-mutated (ATM). J Biol Chem 288(52):37112–37125. https://doi.org/10.1074/jbc.M113.514398 18. Yamane A, Resch W, Kuo N, Kuchen S, Li Z, Sun HW, Robbiani DF, McBride K, Nussenzweig MC, Casellas R (2011) Deep-sequencing identification of the genomic targets of the cytidine deaminase AID and its cofactor RPA in B lymphocytes. Nat Immunol 12:62–69. https://doi.org/10.1038/ni.1964 19. Yamane A, Robbiani DF, Resch W, Bothmer A, Nakahashi H, Oliveira T, Rommel PC, Brown EJ, Nussenzweig A, Nussenzweig MC, Casellas R (2013) RPA accumulation during class switch recombination represents 50 –3’ DNA-end resection during the S-G2/M phase of the cell cycle. Cell Rep 3:138–147. https://doi.org/10.1016/j.celrep.2012.12. 006 20. Aymard F, Bugler B, Schmidt CK, Guillou E, Caron P, Briois S, Iacovoni JS, Daburon V, Miller KM, Jackson SP, Legube G (2014) Transcriptionally active chromatin recruits homologous recombination at DNA doublestrand breaks. Nat Struct Mol Biol 21:366–374. https://doi.org/10.1038/ nsmb.2796

Chapter 6 Genetic and Molecular Approaches to Study Chromosomal Breakage at Secondary Structure–Forming Repeats Anissia Ait Saada, Alex B. Costa, and Kirill S. Lobachev Abstract DNA repeats capable of adopting stable secondary structures are hotspots for double-strand break (DSB) formation and, hence, for homologous recombination and gross chromosomal rearrangements (GCR) in many prokaryotic and eukaryotic organisms, including humans. Here, we provide protocols for studying chromosomal instability triggered by hairpin- and cruciform-forming palindromic sequences in the budding yeast, Saccharomyces cerevisiae. First, we describe two sensitive genetic assays aimed to determine the recombinogenic potential of inverted repeats and their ability to induce GCRs. Then, we detail an approach to monitor chromosomal DSBs by Southern blot hybridization. Finally, we describe how to define the molecular structure of DSBs. We provide, as an example, the analysis of chromosomal fragility at a reporter system containing unstable Alu-inverted repeats. By using these approaches, any DNA sequence motif can be assessed for its breakage potential and ability to drive genome instability. Key words Inverted repeats, Secondary structures, Genome instability, DSB detection, Gross chromosomal rearrangements

1

Introduction DSBs are harmful DNA lesions that cells experience during each cell cycle. Their detrimental outcomes include a variety of chromosomal rearrangements, mutagenesis, and even cell lethality. DNA can acquire DSBs upon exposure of cells to extracellular damaging agents, during replication fork collapse, and due to activation of programmable, site-specific DNA endonucleases [1]. Importantly, chromosomes, especially in eukaryotes, often carry an intrinsic potential for breakage due to the presence of repeats with an internal symmetry that allows DNA to fold into non-B-form secondary structures or accumulate noncanonical intermediates. Stable hairpins and cruciforms, G4 quadruplexes, triplexes, and R-loops can trigger DSBs and rearrangements because (1) they are targets for structure-specific nucleases and (2) they represent strong barriers for DNA replication [2]. Bioinformatic algorithms

Andre´s Aguilera and Aura Carreira (eds.), Homologous Recombination: Methods and Protocols, Methods in Molecular Biology, vol. 2153, https://doi.org/10.1007/978-1-0716-0644-5_6, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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designed to identify secondary structure–forming motifs are not always accurate and are only suggestive. Hence, analysis of the potential of particular regions of DNA to be fragile often requires experimental evaluation. Here, we present methods to study the ability of these sequence motifs to induce genetic instability in the chromosomal context by sensitive genetic assays and by direct detection and assessment of the molecular structure of mitotic DSBs. Overall, these methods allow two main questions to be addressed. First, does a particular DNA sequence motif induce recombination and GCRs and to what degree? Second, what is the mechanism of DSB formation? The latter question requires identification of proteins that maintain the stability of the repeats or trigger their fragility. Below, we describe methods to analyze fragility promoted by inverted repeats. We have applied similar approaches to study the instability of GAA/TTC triplet repeats [3, 4]. We believe that any sequence motif suspected to induce breakage and rearrangements can be studied using these methods. As a rule of thumb, we insert inverted repeats into the counterselectable marker LYS2 located on URA3- or TRP1-containing integrative vectors [5]. These vectors allow replacement of chromosomal LYS2 at the endogenous locus on chromosome II (ChrII) or at LYS2 relocated to the left arm of chromosome V (ChrV) next to the CAN1 gene (in the GCR assay, Fig. 1) with a lys2::Alu-IRs allele using the “pop-in, pop-out” technique, selecting Lys
Ura
or Lys
Trp
isolates with a replacement on alpha-aminoadipatecontaining medium. In the recombination assay, we integrate a second mutant lys2 allele (e.g., lys2-8 or lys2-del5’) into the LEU2 locus on chromosome III (ChrIII) (Fig. 1, strain A) [3, 6]. The spontaneous rate of interchromosomal ectopic recombination between lys2 alleles that do not contain fragile motifs is ~2  10
7 cells/division. Insertion of a 320 bp long Alu quasipalindrome (Alu-IRs with a 12 bp asymmetrical spacer), Alu-QP, into the LYS2 gene leads to a nearly 1,000-fold increase in the recombination rate. These inverted repeats were found to induce chromosomal DSBs with a frequency of ~2%. This system was instrumental in identifying hyporecombination mutants deficient in the endonuclease activity of the Mre11/Rad50/Xrs2 (MRX) complex that is required to open and process hairpin-capped breaks formed at the location of Alu-QP [6]. The simplicity of the assay makes it attractive to assess the fragility potential of any analyzed sequence motif. However, it has several disadvantages. First, the frequency of spontaneous interchromosomal ectopic recombination is relatively high. Therefore, only repeats that induce breaks leading to recombination with frequencies higher than the background level can be assessed. Second, the Alu-QP-induced recombination frequency is very high. A print from replica-plated colonies on media lacking lysine is completely covered with Lys+ papillae making a screen for hyperfragility mutants that predispose Alu

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Fig. 1 Experimental systems to study chromosomal instability induced by secondary-structure forming sequences. Breaks occurring at fragile motifs can lead to ectopic recombination. These events can be scored using strain A in which, in addition to the fragile motif inserted into the LYS2 gene, a truncated version of LYS2 (lys2-del50 ) is inserted into ChrIII. Interchromosomal recombination between the two nonfunctional lys2 genes restores a wild type LYS2, allowing cells to grow on a medium lacking lysine. Strain B contains only the GCR cassette inserted into the nonessential, telomere-proximal part of ChrV. This strain is used in the GCR assay and for DSB detection. A break occurring at the fragile motif leads to formation of a 43 kb fragment, loss of which results in canavanine-resistant and adenine auxotroph (CanR Ade
) red colonies. Chromosome breaks can be directly detected by separating chromosomes on CHEF gels and using probe P1. Breaks detected after restriction digestion of the DNA are detected using probe P2

repeats for breakage problematic. Third, a screen for mutants deficient in the DSB formation and that are expected to have the hyporecombination phenotype is also difficult because a similar phenotype is exhibited by mutants that affect general DSB repair and recombination (such as rad52, rad51, rad54, etc.). In the GCR assay (Fig. 1, strain B), lys2-carrying inverted repeats is located next to the counter-selectable CAN1 marker and ADE2 on the left arm of ChrV [4, 7]. The ~43 kb region between the left telomere and the lys2 gene does not contain essential genes and can be lost. These events can be scored by isolating canavanine-resistant, red ade2 auxotrophs. A detailed description of the assay and selective media used are presented in Fig. 1 and “Methods” below. The spontaneous rate of ChrV arm loss is extremely low: ~2  10
9 cells/division. Therefore, it makes

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this assay very sensitive in identifying repeats even with a minor potential for instability. Insertion of Alu-QP leads to a ~10,000fold increase in the GCR rate. The deduced mechanism for the GCRs is the formation of hairpin-capped breaks due to cruciform resolution, formation of a dicentric chromosome, breakage during anaphase, resection, and repair of broken forks via break-induced replication involving Ty or delta elements that leads to acquisition of a telomere from a nonhomologous chromosome [4]. This assay was instrumental in identifying 17 proteins involved in the maintenance of repeat stability [7, 8]. However, similar to the limitations of previously described recombination assay, caution should be taken when interpreting results leading to identification of mutants with low frequency of GCRs. For example, deficiency in one of the steps during repair of the broken chromosomes (e.g., the removal of the non-homologous tails and break-induced recombination) would be manifested as a decrease in GCR levels. However, these hypo-GCR mutants are proficient in break formation. Overall, the genetic assays described above are powerful in providing insights into mechanisms promoting, preventing, and repairing DSBs occurring at fragile motifs and can be used for genetic screens. Their sensitivity and ease of use make them a good primary approach to study inverted repeats and fragile motifs in general. Nevertheless, these genetic assays, if possible, should be accompanied by molecular techniques allowing one to physically determine the frequency of the DSB formation at a molecular level and validate the effects of the mutations. We present here a straightforward method for detection of DSBs occurring at the location of inverted repeats by Southern blot hybridization with probes designed for detection of the broken chromosome arm. We found that the minimum level of mitotic DSBs reliably detected by this method is ~1% ([7] and data not shown). One challenging issue when using this technique is to avoid breaking chromosomal DNA during the purification step. DNA purification protocols that rely on the precipitation of DNA following enzymatic or mechanical treatments inevitably yield mechanically sheared DNA and generate a high background of nonspecific bands of intensity comparable to the expected DSB bands. To overcome this problem, we embed cells directly into agarose plugs and all of the following treatments of the cells take place “in-plug” in order to preserve DNA integrity. Genomic DNA in-plug can be analyzed directly by CHEF (contour-clamped homogenous electric field) gel electrophoresis to separate broken arms (43 kb in the example we give) from intact chromosomes (585 kb). If broken fragments are long (>40 kb), they can be visualized even in resection-proficient cells. We used this approach to quantify DSB formation in wild type and sae2 mutants carrying a perfect Alu-palindrome (Fig. 2). Whereas this technique is versatile for the direct quantification of DSBs occurring at fragile motifs, it

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Fig. 2 Example of DSB detection by CHEF. (a) Schematic depiction of break formation at a secondary structure. Cruciform formation at inverted repeats can be targeted by nucleases leading to hairpin-capped broken ends (1). Replication of hairpin-capped chromosomal fragments leads to dimer formation, i.e., dicentric and acentric chromosomes (2). Of note, only replication of the 43 kb fragment is shown. Acentric dimers and broken arms are detected using probe P1 (the probe is specific to the HPA3 gene). (b) Example of DSB formation at an Alu-palindrome. In this example, chromosomes were separated on a 1% pulsed field agarose gel for 26 h at 6 V/cm using a two-state mode with a 120 included angle and the following electrical parameters: initial switch time ¼ 3.14 s, final switch time ¼ 7.68 s, ramping factor a ¼ linear. An HPA3specific probe was used to reveal a 585 kb band corresponding to unbroken ChrV, an 86 kb band corresponding to the acentric dimer, and a 43 kb band corresponding to the broken arm. The broken arm fragment has different migration properties depending on its resection state: in wild-type strains, opening of hairpin-capped broken ends by the MRX complex leads to resection. The resulting ssDNA-containing molecules run with delay [10] and present a more smeared profile compared to nonresected fragments observed in sae2Δ strains where MRX attack is prevented

does not allow the molecular structure of the break to be deduced. Another limit of the technique is that replicating chromosomes are resistant to entering the gel due to the presence of the DNA branched structures and, therefore, only DSBs occurring outside of S phase can be detected. A modification of this technique is to digest agarose-embedded DNA by appropriate restriction enzymes, which allows mapping of rare DSBs in mitotic cells accurately. It is important to keep in mind that, depending on the resection range at DSBs, the result can be biased since ssDNA is refractory to restriction digestion. This makes the technique unsuitable for quantitative analysis in resection-proficient strains but rather powerful for revealing the molecular structure of the breaks. In the case of inverted repeats, deficiencies in the MRX complex or Sae2 almost completely block resection of the DSBs because termini of the broken molecules are covalently closed hairpins that require the nuclease activity of Mre11 for processing. Therefore, fine mapping of the hairpincapped breaks using restriction digestion is greatly facilitated in

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Fig. 3 Analysis of the structure of DSB formed at inverted repeats. (a) Representation of restriction digestion (vertical grey arrows) of unbroken chromosome, acentric dimer, and broken arm. “n” represents the distance between the fragile motif and the telomere-proximal restriction site (RS1). In this example, the distance between the fragile motif and the centromere-proximal restriction site (RS2) is larger than n. Therefore, the unbroken fragment runs slower than the dimer and DSB fragments (unbroken fragment > 2n, acentric dimer ¼ 2n, broken arm ¼ n). (b) Schematic illustration of the analysis of hairpin-capped breaks by 2DGE and migration profile of DNA molecules in (a) In neutral-neutral 2DGE, DNA molecules are separated according to their native sizes during the first and second dimension (left panel). In neutral-alkaline 2DGE, DNA molecules are separated according to their native sizes only during the first dimension. Then, DNA is denatured and the second dimension is run in alkaline conditions. Denaturation of the hairpin-capped molecules generates a linearized fragment of 2n size similar to the denatured acentric dimer. Thus, in alkaline 2DGE, the n-sized hairpin-capped break should be detected at the same position as a 2n fragment after running DNA in the second dimension

mrx and sae2 mutants [6, 7]. Separation of digested, unresected DNA by alkaline two-dimensional gel electrophoresis (2DGE) in the protocol described below allows one to gain insight into the molecular structure of a DSB (Fig. 3).

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Materials

2.1 Recombination Assay

1. Strains containing a secondary-forming motif (e.g., Alu-IR) inserted into the LYS2 gene (on ChrII or V) and a truncated version of LYS2 (lys2-del50 allele cloned into pRS305 vector, p305L3) integrated into leu2 on ChrIII: MATα, ade5-1, his72, leu2-3112:: p305L3 (LEU2), trp1-289, ura3-Δ, lys2::Alu-IR. Control strain containing a sequence without any potential to form non-B-DNA (e.g., Alu direct repeats) inserted into LYS2. 2. Sterile, 96-well plates to dilute cells. 3. YPD plates: 10 g yeast extract (BD), 20 g peptone (BD), 20 g glucose (BD), 17 g agar (Sigma). 4. Synthetic medium plates without lysine: 1.4 g/l -Lys dropout (20 mg Adenine sulfate, 20 mg L-Arginine HCl, 100 mg LAspartic acid, 20 mg L-Histidine HCl,100 mg L-Glutamic acid, 30 mg L-Isoleucine, 100 mg L-Leucine, 20 mg L-Methionine, 400 mg L-Serine, 50 mg L-Phenylalanine, 200 mg L-Threonine, 20 mg L-Tryptophan, 30 mg L-Tyrosine, 20 mg Uracil, 150 mg L-Valine), 20 g/l glucose, 1.7 g/l Difco yeast nitrogen base without amino acids and bases, 5 g/l ammonium sulfate, 17 g/l agar. 5. Colony counter.

2.2

GCRs Assay

1. Strains containing the GCR cassette: MATa, bar1Δ, trp1Δ, his3Δ, ura3Δ, leu2Δ, ade2Δ, lys2Δ, V34205::ADE2, lys2::AluIRs. 2. Sterile, 96-well plates to dilute cells. 3. YPD plates. 4. Synthetic medium plates containing canavanine and low concentration of adenine: 1.4 g/l -Arg dropout (5 mg Adenine sulfate, 20 mg L-Arginine HCl, 100 mg L-Aspartic acid, 20 mg L-Histidine HCl, 100 mg L-Glutamic acid, 30 mg L-Isoleucine, 100 mg L-Leucine, 30 mg L-Lysine HCL, 20 mg L-Methionine, 400 mg L-Serine, 50 mg L-Phenylalanine, 200 mg LThreonine, 20 mg L-Tryptophan, 30 mg L-Tyrosine, 20 mg Uracil, 150 mg L-Valine.), 20 g/l glucose, 1.7 g/l Difco yeast nitrogen base without amino acids and bases, 5 g/l ammonium sulfate, 17 g/l agar, 60 mg/l of canavanine (20 mg/ml canavanine stock solution in water). 5. Colony counter.

2.3

Agarose Plugs

1. YPD liquid media. 2. 0.5 M EDTA pH 7.5. 3. Plug solution: 0.5 M EDTA pH 7.5, 10 mM Tris–HCl pH 7.5. Autoclave and store at room temperature.

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4. SCE solution: 1 M sorbitol, 0.1 M sodium citrate, 0.05 M EDTA, pH 7.5. 5. Zymolyase solution: 50 mg/ml Zymolyase 20T, 10% beta mercaptoethanol in SCE (store at
20  C). 6. Low-melt (LM) agarose (Lonza) to make plugs. 7. Plug mold (see Note 1). 8. Proteinase K solution: 5 mg/ml proteinase K, 5% laurylsarcosine in 0.5 M EDTA (store at
20  C). 2.4

CHEF

1. MidRange PFG ladder (NEB). 2. Pulsed-field certified chromosomes.

agarose

(Bio-Rad)

to

separate

3. 0.5 TBE: 45 mM Tris-base, 45 mM boric acid, 1 mM EDTA. 4. Bio-Rad CHEF mapper XA apparatus. 5. 10 mg/ml ethidium bromide. 6. Genescreen nylon membrane (Perkin Elmer). 7. Whatman filter paper. 8. Apparatus for electric transfer (Idea Scientific). 9. 0.4 N NaOH. 10. 2 SSC. 11. PerfecHyb Plus Hybridization Buffer (Sigma). 12. BcaBEST Labeling Kit (Takara) for probe labeling (see Note 2). 13. α-32P dCTP (Perkin Elmer). 14. G-50 Microspin columns (GE Healthcare). 15. Salmon sperm DNA. 16. Wash solution: 0.1 SSC, 0.1% SDS. 17. Phosphor screen and a suitable cassette. 2.5 Neutral-Neutral and Neutral-Alkaline 2D Gels

1. 1 TE: 10 mM Tris–HCl, 1 mM EDTA, pH 7.5. 2. Prepare freshly made 1 TE containing 1 mM PMSF (from 100 mM PMSF stock solution in isopropanol). 3. Restriction enzyme and its appropriate restriction buffer. 4. 1 kb DNA ladder. 5. 1 TBE: 89 mM Tris-base, 89 mM boric acid, 2 mM EDTA. 6. UltraPure Agarose to run DNA in first and second dimension. 7. 10 mg/ml ethidium bromide. 8. Ruler and razor.

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9. Horizontal electrophoresis tanks: Thermo Owl A1 large gel electrophoresis system for first dimension and Thermo Owl A5 with Built-In Recirculation for second dimension. 10. Alkaline buffer I: 250 mM NaOH, 5 mM EDTA pH 8. 11. Alkaline buffer II: 50 mM NaOH, 1 mM EDTA pH 8. 12. Genescreen nylon membrane. 13. Whatman filter paper. 14. Depurination solution: 0.25 N HCl. 15. Denaturation solution: 0.5 N NaOH, 1.5 M NaCl. 16. Neutralization solution: 1.5 M NaCl, 1 M Tris–HCl, pH adjusted to 7.4. 17. PerfecHyb Plus hybridization buffer. 18. 10 SSC. 19. BcaBEST Labeling Kit for probe labeling. 20. α-32P dCTP. 21. G-50 Microspin columns. 22. Salmon sperm DNA. 23. Wash solution: 0.1 SSC, 0.1% SDS. 24. Stratagene’s PosiBlot 30–30 pressure blotter and pressure control station (or custom-made posiblotter). 25. Phosphor screen and a suitable cassette.

3

Methods

3.1 Fluctuation Test to Score for Homologous Recombination Events (See Note 3)

1. Streak out the strains to be tested on YPD plates and culture at 30  C for 3 days to obtain single colonies. 2. Select 14 colonies per strain and resuspend each colony in 200 μl of water in a 96-well plate. 3. Perform a five-step 1:10 serial dilution of all suspensions. 4. Depending on the colony size, plate either the 1/10000 or 1/100000 dilution on YPD plates (2  100 μl). This will allow cell viability/plating efficiency to be defined. 5. Plate the 0 dilution for the control and 1/100 dilution for the strains with Alu-QP on -Lys plates (2  90 μl). 6. Incubate at 30  C until the colonies can be counted. 7. Determine the rate and confidence interval of recombination between lys2 alleles using the number of colonies on YPD and corresponding selective media plates.

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3.2 Fluctuation Test to Score for GCR Events (See Notes 3 and 4)

1. Streak out the strains to be tested on YPD plates and culture at 30  C for 3 days to obtain single colonies. 2. Select 14 colonies per strain and resuspend each colony in 200 μl of water on a 96-well plate. 3. Perform a five-step 1:10 serial dilution. 4. Depending on the colony size, plate either the 1/10000 or 1/100000 dilution on YPD plates (2  100 μl). This will allow cell viability/plating efficiency to be defined. 5. Plate the 1/100 or 1/1000 dilution on canavanine-containing plates (2  90 μl). 6. Incubate the plates at 30  C until the colonies grow and can be counted (see Note 5). 7. Determine the rate and confidence interval of GCRs using the number of colonies on YPD plates and the number of red colonies on corresponding selective media plates.

3.3 Cell Embedding in Agarose Plugs

A challenge in DSB detection is to yield intact purified DNA. Therefore, this protocol has been optimized to avoid mechanical shearing of DNA molecules as much as possible. By embedding cells into agarose plugs, spheroplasting cells in plugs, and keeping plugs at temperatures below 30  C, DNA integrity is maintained. 1. Select a single colony and grow cells overnight in 10 ml of YPD (see Note 6) until saturation is reached. 2. Determine cell density by counting the cells using a hemocytometer. 3. Pellet the cells and resuspend in 1 ml of water. 4. Transfer between 4 and 8  108 cells per plug to a 2 ml microcentrifuge tube, pellet, and remove supernatant. 5. Prepare 1.5% LM agarose in 0.1 M EDTA pH 7.5 and keep solution at 50  C (see Note 7). 6. Add plug solution to the pellet to reach a total volume of 70 μl. Vortex briefly. 7. Add 10 μl of zymolyase solution. Vortex briefly. 8. Warm the cells in 50  C water bath and add 80 μl of preheated, 1.5% LM agarose. Keep the cells at 50  C for the minimum time required to mix the cells with the LM agarose. 9. Load the suspension into the plug mold immediately using a 200 μl micropipette. 10. Repeat steps 6–9 for each sample. 11. Allow the plugs to solidify 5–10 min at 4  C. 12. Push the plugs into microcentrifuge tubes containing 800 μl of plug solution and 10 μl of zymolyase solution (see Note 8). 13. Incubate 2 h to overnight at 30  C.

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14. Add 200 μl of proteinase K solution and incubate several hours to overnight at 30  C. 15. Replace the proteinase K solution with 1 ml of fresh plug solution. 16. Store plugs at 4  C. 3.4

CHEF Analysis

1. Prepare 3 l of 0.5 TBE. 2. Slice the plugs to the desired length. 3. Equilibrate each sliced plug in running buffer (0.5 TBE) for 30 min. 4. Prepare 1% pulsed field certified agarose gel in running buffer and cool to 50–55  C (see Note 9). 5. Align each plug and a very thin slice of PFG marker along the top of the gel tray. 6. Immobilize the plugs by applying melted agarose with a 1 ml micropipette and allowing the agarose to partially solidify. 7. Pour the remaining gel and allow to solidify. 8. Add the remaining prepared 0.5 TBE to the CHEF mapper tank and chill to 14  C. 9. Run the gel using the appropriate parameters (see example in Fig. 2b). 10. Stain the gel with 0.5 μg/ml ethidium bromide in 0.5 TBE and cut the region to be transferred. 11. Transfer DNA in 0.5 TBE at 12 V for 2–4 h (see Note 10). 12. Crosslink DNA to the membrane by exposing it to 120 mJ of UV-C. 13. Soak the membrane in 0.4 N NaOH for 10 min to denature DNA. 14. Wash the membrane in 2 SSC for 10 min. 15. Proceed to hybridization as described in Subheading 3.6 (see Note 11).

3.5 Neutral-Neutral and Neutral-Alkaline 2DGE 3.5.1 In-Plug Digestion of Agarose-Embedded DNA

1. Slice the plugs to the desired length and transfer them to a 24-well plate. Prepare two slices per strain (see Note 12). 2. Wash the plugs twice in 1 TE for 30 min. 3. Wash in 1 TE containing 1 mM PMSF for 1 h. 4. Wash in H2O for 30 min. 5. Transfer the plugs to 2 ml microcentrifuge tubes. 6. Incubate in 200 μl of 2 restriction enzyme buffer for 1 h. 7. Incubate in 200 μl of 1 restriction enzyme buffer for 1 h. 8. Add 50 units of restriction enzyme.

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9. Incubate at 37  C overnight. 10. Add 25 units of restriction enzyme and incubate for 3 additional hours. 11. Remove digestion solution and equilibrate the plugs in 200 μl of 1 TBE. 3.5.2 Running DNA in the First Dimension

1. Prepare a plug containing a 1 kb ladder by mixing 15 μl of 1 kb ladder with 15 μl of LM agarose. 2. Align the plugs and the 1 kb ladder plug along the top of the gel tray. Leave ~0.5–1 cm or more between each plug. 3. Prepare 250 ml of 0.7–1% agarose gel in 1 TBE. Cool to 50–55  C. 4. Immobilize the plugs by applying melted agarose with a 1 ml micropipette. 5. Pour the remaining gel and allow it to solidify. 6. Add 1.5 l of 1 TBE to an Owl A1 electrophoresis tank. 7. Carefully place the gel tray in the tank ensuring it is not floating and it is submerged in running buffer. 8. Run at slow voltage (90%; reduced viability may result in excessive cell death upon transfection. The quality and concentration of the DNA is also extremely important for optimal transfection; we recommend to use maxiprep kit from Qiagen or similar to achieve a DNA stock concentration of ~1 mg/ml of endotoxin-free, high-quality DNA.

3.1 Generation of EGFPMBP-BRCA2 Stable Cell Lines in Human DLD1 BRCA2 Deficient Cells (See Workflow in Fig. 1)

The day before transfection, seed 2.5–3  106 cells (BRCA2
/
) per 10 cm-cell culture plate in order to obtain a cell confluency of 70% the following day (see Note 4). Use RPMI complete medium supplemented with 0.1 mg/ml hygromycin (see Note 5).

3.1.1 Day 0: Seeding the Cells 3.1.2 Day 1: Transfection

Transfect one 10 cm cell culture plate with 10 μg of EGFPMBPBRCA2 plasmid (Fig. 1a) using 20 μl TurboFect transfection reagent: 1. Remove the medium and rinse the cells twice with 5 ml of 1 PBS.

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2. Add 10 ml of RPMI complete medium (without selection antibiotics, see Note 5) to the cells, and put the plate back in the incubator until the transfection mix is ready. 3. Prepare the transfection mix: Add 10 μg of plasmid to 1 ml RPMI serum-free medium in a 1.5 ml eppendorf tube, vortex for a few seconds and spin down briefly. Add 20 μl of TurboFect transfection reagent to the DNA mix without touching the sides of the tube, vortex, and spin down briefly. 4. Incubate the DNA transfection mix for 20 min at room temperature. 5. Add the transfection mix dropwise to the cells and incubate the cells at 37  C in a humidified incubator with 5% CO2. 3.1.3 Day 2: Change of Medium

1. Remove the medium from the transfected cells and add 10 ml RPMI complete medium. Incubate the cells at 37  C in a humidified incubator with 5% CO2.

3.1.4 Day 3: Serial Dilution

1. Remove the medium and rinse the cells twice with 5 ml 1 PBS. 2. Add 1 ml trypsin to the plate and incubate at 37  C for 3–5 min. 3. Resuspend the de-attached cells in 8 ml RPMI complete medium supplemented with hygromycin (0.1 mg/ml) and G418 (1 mg/ml) selection antibiotics (see Note 5). 4. Make 1:2, 1:4, 1:8, 1:16, and 1:32 dilutions into 10 cm cell culture plates using complete medium supplemented with selection antibiotics (hygromycin and G418, see Note 5). 5. Change the medium every 3–4 days (RPMI complete medium supplemented with selection antibiotics, see Note 5). Cell colonies will start to appear 10 days after transfection.

3.1.5 Day 10–15: Isolation of Single-Cell Clones

Pick the clones once you observe cells forming white colonies under light by the naked eye. Make sure you pick well-separated colonies (see Subheading 3.1.4 step 4). 1. Prepare a 96-well cell culture plate by adding 200 μl/well of prewarmed RPMI complete medium supplemented with selection antibiotics (see Note 5). 2. Mark your colony at the bottom of the plate. 3. Remove the medium from the 10 cm cell culture plate and rinse the cells twice with 5 ml 1 PBS. 4. Use a P10 micropipette to carefully add a few μl of trypsin to the colony, pipette up and down until the cells have de-attached, and transfer it to a single well in the prepared 96-well plate. Pick around 30–50 single colonies.

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5. Incubate the 96-well plate at 37  C in a humidified incubator with 5% CO2. 3.1.6 Day 15–30: Expansion of Single-Cell Clones

3.1.7 Selection of Single Clones for HR Assay: Harvesting, Lysis, and Western Blotting

1. Maintain the clones in RPMI complete medium supplemented with selection antibiotics. 2. When the cells reach ~70% confluency expand them stepwise from the 96-well plate to 24- and 6-well plates to finally 6/10 cm plates. Harvest half of the cells for analysis of the protein expression level by western blot (see next Subheading 3.1.7) and continue to expand the rest of the cells to make a frozen stock. Harvest the cells for western blot analysis when the cells have reached 70% confluency in a 6/10 cm plate. 1. Remove the media and rinse the cells twice with 5 ml 1 PBS. 2. Add 1 ml trypsin to the plate and incubate at 37  C for 3–5 min. 3. Resuspend the trypsinized cells in 10 ml RPMI complete medium, take 10 μl of the cell suspension and mix with 10 μl of Trypan blue, and measure viable cells using a cell counter. 4. Re-seed 5 ml of cell suspension in a 10 cm plate for expansion of the clones to make a frozen stock, and incubate at 37  C. 5. Centrifuge the other 5 ml cell suspension in a 15 ml tube at 300  g for 5 min at 4  C. 6. Remove the medium, add 10 ml cold 1 PBS to the cell pellet, and centrifuge at 300  g for 5 min at 4  C. 7. Resuspend the cell pellet in cold lysis buffer H (1 ml buffer/ 1  106 cells) and incubate on ice for 30 min, and vortex every 10 min. 8. Sonicate and centrifuge at 10,000  g for 15 min, and collect the supernatant. 9. Determine the total protein concentration using Bradford assay. 10. Run 50–100 μg total protein lysate on a 4–15% SDS-PAGE followed by immunoblotting using BRCA2 (see Note 1) and GFP antibodies to detect EGFPMBP-BRCA2. 11. Select clones with similar expression levels for the recombination assay.

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3.2 Recombination Assay in Human DLD1 Cells (See Scheme of the Assay in Fig. 2a) 3.2.1 Day 0: Seeding the Cells 3.2.2 Day 1: Transfection

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The day before transfection (day 0), seed the cells (2.5–3  106 cells per 10 cm cell culture plate) such that they are subconfluent on the day of transfection (~70% confluency) (see Note 4 and Fig. 4b). Incubate the cells at 37  C in a humidified incubator with 5% CO2.

Co-transfect the cells with the two TALEN plasmids together with the promoter-less mCherry donor plasmid (Fig. 2b, c) using Amaxa Cell Line Nucleofector Kit V, following manufacturer specifications. For the negative-control reaction, transfect the cells with the mCherry donor plasmid alone.

Fig. 2 Schematic representation of the HR assay. (a) This reporter assay comprises the co-transfection of two plasmids coding for TALENs designed to generate a DSB at the AAVS1 locus in the genome, together with a promoter-less donor plasmid enconding the mCherry fluorescent protein bearing two ~800 bp regions of homology to the AAVS1 site at both sides of the mCherry gene (50 arm and 30 arm). Once transfected, the TALENs are expressed and generate a site-specific DSB in the PPP1R12C locus (black box, TALEN site). When the HR pathway is active, the promoter-less mCherry donor is integrated in the PPP1R12C locus downstream of the endogenous promoter of the same gene, driving the expression of mCherry that can be then detected by flow cytometry with the appropriate filter. SA, splicing acceptor; 2A, ribosome stuttering signal. Exons are depicted as blue boxes; promoter-less cherry is depicted in gray in the donor plasmid and in red in the modified locus. (b) Plasmid map of the of the AAVS1-2A-mCherry promoter-less donor plasmid. (c) Representation of the human chromosome 19 showing the AAVS1 locus (blue arrow) within the PPP1R12C gene and the TALEN target sequences used in this HR assay

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1. Prepare a DNA master mix containing 3 μg of the donor plasmid and 1 μg of each TALEN plasmid. For the negativecontrol reaction, prepare a DNA mix with the donor plasmid and replace TALEN with dH2O. 2. Prepare a 6-well plate with 3 ml of RPMI complete medium per well and preincubate the plate at 37  C in a humidified incubator with 5% CO2. 3. Prepare a master mix of the appropriate Cell Line Nucleofector solution (solution V for DLD1 cells) by adding supplement solution (included in the AMAXA kit) to the Nucleofector solution in a ratio of 4.5:1; 100 μl of the master mix is needed per transfection (see Note 6). 4. Select the appropriate program on the Nucleofector (L-024 for DLD1 cells) and calibrate with a blank (either 100 μl of dH2O or 1 PBS) added to the electroporation cuvette (see Notes 3 and 7). 5. Remove the medium from the culture plate and rinse the cells once with 2 ml 1 PBS. Add 0.5 ml trypsin per well and incubate at 37  C for 3–5 min. 6. Resuspend the trypsinized cells in 5 ml RPMI complete medium, take 10 μl of the cell suspension and mix with 10 μl of Trypan blue, and measure viable cells using a cell counter (see Note 8). 7. Centrifuge 1  106 cells per reaction in a microcentrifuge tube (one tube per transfection) at 300  g for 5 min at 20  C. 8. Remove the medium and resuspend the cell pellet in 1 ml 1 PBS by pipetting up and down, and centrifuge again at 300  g for 5 min at 20  C. 9. Remove the PBS and resuspend the cell pellet in 100 μl of Nucleofector Solution V (see step 3), add the DNA mix (see step 1), and transfer to an electroporation cuvette (see Note 9). 10. Right before electroporation of the cells, insert the blank cuvette into the Nucleofector cuvette holder and run the nucleofector program (L-024) (see Note 7). Remove the blank and insert the cuvette with the sample and electroporate the cells with the L-024 program (see Note 3). 11. Once the program is finished, transfer the sample from the cuvette to the prepared 6-well plate (see step 2) using the pipette provided by AMAXA kit. We recommend using ~500 μl of the prewarmed medium from the well to recover the sample from the cuvette (see Note 9). 12. Incubate the cells at 37  C in a humidified incubator with 5% CO2.

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3.2.3 Day 2: Change of Medium

Remove the medium from the cells and replace it with fresh RPMI complete medium with appropriate selection antibiotics (see Note 5).

3.2.4 Day 3–7: Cell Expansion

Once the cells have reached ~70% confluency passage them as required.

3.2.5 Day 8: Harvesting the Cells for Flow Cytometry

1. Remove the medium from the plates and rinse the cells once with 1 ml 1 PBS. Add 0.5 ml trypsin per well and incubate at 37  C for 3–5 min. 2. Resuspend the trypsinized cells in 2 ml RPMI complete medium and transfer the cells to a flow cytometry tube. 3. Centrifuge the cells at 300  g for 3 min at 20  C. 4. Remove the medium and resuspend the pellet in 2 ml 1 PBS, and centrifuge the cells at 300  g for 3 min at 20  C. 5. Remove PBS and resuspend the cells in an appropriate amount of 1 PBS (see Note 10).

3.2.6 Fluorescence-Activated Cell Sorting (FACS) Analysis

The percentage of mCherry-positive cells is analyzed by flow cytometry and data is analyzed using FlowJo or similar software (see Note 11). 1. Gate the viable cells from the Forward Scatter (FSC-A, x-axis) versus Side Scatter (SSC-A, y-axis) plot. 2. Display the gated viable cells in a FSC-A (x-axis) versus FSC-W (y-axis) versus plot or a SSC-A versus SSC-H plot to exclude doublets. 3. Display the gated singlet viable population in a mCherry-A (yaxis) versus FSC-A (x-axis) plot to monitor the number of mCherry-positive cells. An example of the results of the assay obtained by FACS is shown in Fig. 3. The quantification of the same experiments is shown in Fig. 4a.

4

Notes 1. For detection of EGFPMBP-BRCA2 with western blotting we use mouse anti-BRCA2 (1:1000, OP95, EMD Millipore). 2. We replaced the original GFP from the donor plasmid, promoter-less AAVS1-2A-GFP-PA plasmid (kind gift from Dr. C. Giovannangeli) with the mCherry tag from the pET28 mCherry plasmid using NEB Gibson Assembly (Gibson Assembly Master Mix, New England BioLabs) to avoid interreference from the GFP from EGFP-MBP-BRCA2. However, the GFP works equally well for these experiments.

Fig. 3 Example of the flow cytometry gating and analysis resulting from a HR assay at 8 days posttransfection. Results for (a) DLD1 BRCA2+/+ parental cells, (b) DLD1 BRCA2 deficient cells (BRCA2
/
), and (c) DLD1 BRCA2
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cells complemented with GFPMBP-BRCA2 (BRCA2 WT). The plots on the right show the analysis of the negative control (cells transfected with mCherry donor alone (
TALEN)). Viable cells were gated from the Forward Scatter (FSC-A) versus Side Scatter (SSC-S) plot and displayed in a FSC-A versus FSC-W plot to exclude doublets. The gated singlet population was displayed in a FSC-A versus mCherry-A plot to detect the mCherry-positive population. 10,000 singlet events were collected for each experiment

B mCherry + cells (%)

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Fig. 4 HR activity assay. (a) HR activity of DLD1 parental cells (BRCA2+/+), DLD1 BRCA2 deficient cells (BRCA2
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TALEN) or together with the two TALEN-bearing plasmids (+TALEN). Results represent the mean from three independent experiments; error bars (SD). (b) The effect of cell seeding density on the HR efficiency was analyzed by measuring the HR activity of DLD1 BRCA2+/+ cells seeded at three different densities the day before AMAXA transfection (the numbers indicate seeding density/10 cm culture plate). For DLD1 BRCA2+/+ cells, a seeding density of 1  106/10 cm plate generates a cell confluency around 60–70% the day after, which gives the highest amount of mCherry+ cells (black bar)

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3. For the co-transfection of the cells with the two TALEN plasmids together with the promoter-less mCherry donor plasmid we use Amaxa Cell Line Nucleofector Kit (Lonza). For DLD1 cells we use Kit V (Lonza VCA-1003) and the program L-024 on the AMAXA Nucleofector Device (Lonza). 4. Cell confluence >70% at the time of transfection may result in lower transfection efficiency and therefore lower HR levels (Fig. 4b). Selection antibiotics in the medium during transfection may also result in lower transfection efficiency. 5. DLD1 BRCA2 deficient (BRCA2
/
) cells require hygromycin selection whereas DLD1 BRCA2
/
stably expressing EGFPMBP-BRCA2 (BRCA2 WT) cells require hygromycin plus G418 disulfate salt. Make sure to supplement RPMI complete medium with the appropriate antibiotic selection. 6. The ratio of Nucleofector solution to supplement is 4.5:1; for a single reaction use 82 μl of Nucleofector solution and 18 μl of the supplement for a total of 100 μl reaction volume. Prepare the master mix of Nucleofector Solution V corresponding to the number of reactions you are planning to include in the experiment. 7. Perform blank calibration before each electroporation. 8. Ensure that there are at least 2  106 viable cells in the plate. 9. Cells should not stay long in neither the Nucleofector solution nor the electroporation cuvette; we recommend you to prepare and electroporate one sample at a time. The transfection sample must cover the entire bottom of the cuvette; remove any air bubbles by gently taping the cuvette on the bench. 10. Depending on the cell number on the day of analysis, resuspend the cells in 200 μl to 1 ml 1 PBS. Cells resuspended in PBS should not be too confluent nor too diluted for FACS analysis (~1  106 cells/ml is optimal). 11. Allow the cells to grow 7–8 days post transfection before performing the FACS analysis to avoid autofluorescence and maximize the signal/noise ratio.

Acknowledgments We thank all members of Carreira lab for fruitful comments on this manuscript. The work in Carreira lab is supported by ANR grant ANR-17-CE12-0016-01 and French Breast Cancer Foundation Cancer du Sein: Parlons-en! DV is supported by a Fellowship from the French National Ministry of Education and Science.

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References 1. Pierce AJ, Johnson RD, Thompson LH, Jasin M (1999) XRCC3 promotes homologydirected repair of DNA damage in mammalian cells. Genes Dev 13:2633–2638 2. Rouet P, Smih F, Jasin M (1994) Expression of a site-specific endonuclease stimulates homologous recombination in mammalian cells. Proc Natl Acad Sci U S A 91:6064–6068 3. Moynahan ME, Pierce AJ, Jasin M (2001) BRCA2 is required for homology-directed repair of chromosomal breaks. Mol Cell 7:263–272 4. Guidugli L, Pankratz VS, Singh N et al (2013) A classification model for BRCA2 DNA binding domain missense variants based on homology-directed repair activity. Cancer Res 73:265–275. https://doi.org/10.1158/ 0008-5472.CAN-12-2081 5. Shimelis H, Mesman RLS, von Nicolai C et al (2017) BRCA2 Hypomorphic missense variants confer moderate risks of breast cancer. Cancer Res 77:2789–2799. https://doi.org/ 10.1158/0008-5472.CAN-16-2568 6. Hockemeyer D, Soldner F, Beard C et al (2009) Efficient targeting of expressed and

silent genes in human ESCs and iPSCs using zinc-finger nucleases. Nat Biotechnol 27:851–857. https://doi.org/10.1038/nbt. 1562 7. Brunet E, Simsek D, Tomishima M et al (2009) Chromosomal translocations induced at specified loci in human stem cells. Proc Natl Acad Sci U S A 106:10620–10625. https://doi. org/10.1073/pnas.0902076106 8. DeKelver RC, Choi VM, Moehle EA et al (2010) Functional genomics, proteomics, and regulatory DNA analysis in isogenic settings using zinc finger nuclease-driven transgenesis into a safe harbor locus in the human genome. Genome Res 20:1133–1142. https://doi.org/ 10.1101/gr.106773.110 9. Ehlen A, Martin C, Miron S, et al Proper chromosome alignment depends on BRCA2 phosphorylation by PLK1. biorxivorg. https://doi. org/10.1101/265934 10. von Nicolai C, Ehlen A, Martin C et al (2016) A second DNA binding site in human BRCA2 promotes homologous recombination. Nat Commun 7:12813. https://doi.org/10. 1038/ncomms12813

Chapter 10 Interhomolog Homologous Recombination in Mouse Embryonic Stem Cells Fabio Vanoli, Rohit Prakash, Travis White, and Maria Jasin Abstract Homologous recombination is a critical mechanism for the repair of DNA double-strand breaks (DSBs). It occurs predominantly between identical sister chromatids and at lower frequency can also occur between homologs. Interhomolog homologous recombination (IH-HR) has the potential lead to substantial loss of genetic information, i.e., loss of heterozygosity (LOH), when it is accompanied by crossing over. In this chapter, we describe a system to study IH-HR induced by a defined DSB in mouse embryonic stem cells derived from F1 hybrid mice. This system is based on the placement of mutant selectable marker genes, one of which contains an I-SceI endonuclease cleavage site, on the two homologs such that repair of the I-SceIgenerated DSB from the homolog leads to drug resistance. Loss of heterozygosity arising during IH-HR is analyzed using a PCR-based approach. Finally, we present a strategy to analyze the role of BLM helicase in this system. Key words Interhomolog homologous recombination (IH-HR), Homology-directed repair (HDR), Loss of heterozygosity (LOH), I-SceI, Mouse embryonic stem cells, Bloom helicase (BLM)

1

Introduction Homologous recombination (HR), also called homology-directed repair (HDR), is an important double-strand break (DSB) repair mechanism in mammalian cells. HR relies on an identical or highly homologous sequence that can act as a template for repair synthesis primed by a DNA end. One source of homology is the identical sister chromatid, which is present following DNA replication in the S and G2 phases of the cell cycle. The identical sister chromatid is the preferred template for repair, likely in part due to its proximity [1]. The homologous chromosome can also be a template for repair, i.e., interhomolog homologous recombination (IH-HR), although it appears to be used less frequently [2]. This contrasts with DSB repair during meiotic prophase I where IH-HR is

Fabio Vanoli and Rohit Prakash contributed equally to this work. Andre´s Aguilera and Aura Carreira (eds.), Homologous Recombination: Methods and Protocols, Methods in Molecular Biology, vol. 2153, https://doi.org/10.1007/978-1-0716-0644-5_10, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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A IH-HR reporter 129 (S)

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CO-LOH Fig. 1 S2neo/Pneo (S/P) reporter to detect IH-HR and LOH. (A) The S/P reporter contains two non-functional copies of the neomycin resistance gene (neo). The S2neo gene is mutated by the insertion of an I-SceI site (red box) at an NcoI site; the Pneo gene is mutated by the insertion of a PacI site at an EagI site. A DSB induced by

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frequent as it is crucial for the reductional division to generate gametes [3]. Except in cases such as inbred lab strains and consanguinity in humans, homologs will contain polymorphisms along their lengths. One outcome of IH-HR is gene conversion such that one or more polymorphisms from one homolog are copied into the other homolog at the site of the DSB. Thus, IH-HR can lead to a small genomic region that has undergone loss of heterozygosity (LOH) [2]. However, a much larger region of LOH results if IH-HR occurs with crossing over: The whole chromosomal region distal to the crossover site will become homozygous if the two exchange chromosomes segregate to opposite daughter cells. Although not a genomic rearrangement per se, LOH can have important genetic consequences, in particular, leading to tumorigenesis if it results in loss of a functional tumor suppressor gene. For example, in the case of hereditary retinoblastoma, loss of the wild-type RB1 allele occurs by IH-HR in 40% of cases [4]. To study mechanisms and the genetic dependencies of IH-HR, a reporter has been developed in mouse embryonic stem (ES) cells (Fig. 1A) [2]. The S/P reporter consists of two non-functional copies of the neomycin-resistance gene (neo), each of which is integrated at the same locus (an Rb1 intron) on chromosome 14 homologs: One neo copy is interrupted by an I-SceI endonuclease recognition site in the 30 portion of the gene (S2neo, S allele) while the second copy is interrupted by insertion of a PacI restriction site in the 50 portion of the gene (Pneo, P allele). An I-SceIgenerated DSB in S2neo induces IH-HR with the homologous sequence in Pneo. DNA synthesis during IH-HR using Pneo as a template restores a functional neomycin-resistance gene (neo+) as long as synthesis does not proceed through the PacI site, such that cells are resistant to media containing the neomycin analog G418. The S/P reporter has been introduced into F1 hybrid-derived ES cell lines of Balb/c  129/SC (129/Balb) [5] and C57BL/ 6  129/Sv (129/B6) origin [6]. The frequency of IH-HR using the 129/Balb strain is ~5  105 [2, 7], similar to the frequency measured in isogenic ES cell lines [8] but ~100 times less frequent than HR between direct repeat neo sequences which can use the sister chromatid as a template [9]. This relatively low frequency of ä Fig. 1 (continued) I-SceI in the S2neo gene (red arrowhead) can be repaired by IH-HR using the Pneo gene as template to restore a functional gene (neo+). (B) IH-HR involving gene conversion without crossing over (noncrossover). LOH is limited to the region of gene conversion surrounding the DSB site. (C) IH-HR involving gene conversion with crossing over. Left: Segregation of a non-exchange chromatid with an exchange chromatid results in LOH from the site of the crossover to the distal end of the chromosome (CO-LOH; light purple shaded box) with daughter cells that can be either neo+ or not. Right: If the exchange chromatids segregate together to the same daughter cell, LOH remains localized to the site of gene conversion

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IH-HR as compared with HR involving direct repeats has also been observed in human B-lymphoblastoid cell lines [10]. Most IH-HR events involve gene conversion at the DSB site without exchange of flanking markers (noncrossover, Fig. 1B) such that the region of LOH is localized, typically only involving the I-SceI recognition site. Gene conversion tracts can extend longer distances although the frequency decreases as the distance between the DSB site and the markers increase, i.e., from 10.5% for a marker 0.3 kb from the DSB to 1.9% for one 6 kb away. Crossing over is infrequent such that only a small fraction of clones analyzed (1.3%) have LOH extending to the end of the chromosome (Fig. 1C). We have termed this crossover-derived LOH, or CO-LOH [2, 7], to distinguish it from other LOH mechanisms such as nondisjunction that would not give rise to a neo+ gene. Crossing over, and hence CO-LOH, is suppressed by BLM, a RECQ helicase family member which is deficient in Bloom syndrome, a pan cancer syndrome also associated with growth retardation [11, 12]. Biochemical experiments have demonstrated that BLM can “dissolve” recombination intermediates which otherwise would be resolved as crossovers [13]. The F1 hybrid 129/B6 ES cell line contains tetracycline-off Blm alleles (Blmtet/tet) which allows titration and abrogation of BLM expression by the addition of doxycycline to the media [6]. Blmtet/tet cells incubated with doxycycline have a small reduction in the overall rate of IH-HR as compared with BLM-proficient cells, but a ~5-fold increase in CO-LOH [7], consistent with the role of BLM in suppressing crossing over. In this chapter, we describe experimental procedures for measuring IH-HR and LOH using our reporter. We also present a detailed protocol for introducing the IH-HR reporter into other cell lines. Finally, a strategy to evaluate the impact of mutant BLM proteins on IH-HR is described, involving ectopic integration of mutant cDNAs at the Rosa26 locus.

2 2.1

Materials Cell Culture

2.1.1 Cell Culture Media for Mouse Embryonic Stem (ES) Cells

1. DME-HG (500 ml). 2. 75 ml fetal bovine serum, ES cell grade. 3. 6 ml penicillin/streptomycin solution (100). 4. 6 ml nonessential amino acids. 5. 6 ml L-glutamine. 6. 50 μl leukemia inhibitory factor (LIF) (Thermo Fisher). 7. 4.3 μl 2-mercaptoethanol. 8. Geneticin (G418), powder.

Inter-Homolog Recombination in Mouse Embryonic Stem Cells 2.1.2 Coating Tissue Culture Dishes or Flasks

1. Gelatin (0.1%).

2.1.3 Passaging Cells

1. Trypsin.

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Gene Targeting

1. Bio-Rad Gene Pulser II. 2. 0.4-cm cuvettes. 3. Sodium acetate (3 M). 4. Ethanol. 5. Bi-distilled water (ddH2O). 6. Hygromycin (50 mg/ml stock).

2.3

Southern Blot

1. Tris/borate/EDTA (TBE). 2. Hydrochloric acid (HCl). 3. Sodium hydroxide (NaOH) powder. 4. Sodium chloride (NaCl) powder. 5. Tris (hydroxymethyl) aminomethane hydrochloride (1.5 M). 6. Sodium dodecyl sulfate (SDS) (10%). 7. Saline-sodium citrate (SSC) (20). 8. Whatman paper. 9. GenScreen Plus membrane. 10. Denhardt’s solution (50): 5 g Ficoll, 5 g polyvinylpyrrolidone, 5 g Fraction V BSA, ddH2O up to 500 ml. Filter and store at 4  C. 11. Salmon sperm DNA (10 mg/ml stock). 12. Random primer labeling kit. 13. ATP [α-32P]. 14. Exposure cassette and film for radioactive detection.

2.4

Staining

1. Methanol 100%. 2. Giemsa stain.

2.5

PCR Analysis

1. Lysis buffer (10): 100 mM Tris pH 8.0, 4.5% NP40, 4.5% Tween 20. Filter and store at 4  C. Make fresh 2 dilution and add proteinase K 100 μg/ml, every time. 2. Jeffrey’s buffer (10): 450 mM Tris HCl pH 8.8, 110 nM (NH4)2SO4, 45 mM MgCl2, 67 mM 2-mercaptoethanol, 44 μM EDTA, 10 mM dATP, 10 mM dCTP, 10 mM dGTP, 10 mM dTTP, 1.13 mg/ml BSA, 12.5 mM TRIS. Store at 20  C. 3. Taq DNA polymerase.

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2.6 Western Blot Analysis

1. TEGN buffer (10 mM Tris, pH 8.0, 1 mM EDTA, 10% glycerol, 0.5% Nonidet P-40, 400 mM NaCl). 2. Protease inhibitor (tablet). 3. DTT (0.1 M stock). 4. Nitrocellulose membrane. 5. Milk powder. 6. Tween-20. 7. PBS.

3

Methods

3.1 Targeting the Rb1 Locus for Generation of S2neo/Pneo (S/P) Cell Lines [2]

S/P cell lines are generated by targeting the SH and PH vectors to the Rb1 locus on mouse chromosome 14. The vectors contain the S2neo and Pneo reporter genes, respectively, followed by a hygromycin-resistance gene (hyg) that is flanked by LoxP sites. The two reporter genes have been inserted into genomic DNA fragments of the Rb1 locus derived from 129 (S2neo) and Balb (Pneo) mouse strains (Fig. 1A) [14]. The resulting constructs can be used for targeting mouse ES cell lines derived from an F1 hybrid of Balb (or B6) and 129 such that the two chromosomes contain polymorphisms along their length. The method presented here describes targeting in the 129/B6 Blmtet/tet cell line [6]. 1. Split ES cells 24 h before transfection from a 60 mm plate (~90% confluent) to 2  10 cm plates. 2. Linearize the SH targeting vector with HpaI. After 2 h incubation at 37  C, precipitate with sodium acetate and ethanol and resuspend in 30 μl of ddH2O. 3. Trypsinize and collect 5  106 cells and wash with PBS. 4. Resuspend cells in 700 μl PBS and combine with the purified targeting vector. 5. Transfect cells with Bio-Rad gene pulser II (800 V and 3 μF in a 0.4-cm cuvette) and plate in 1  10 cm plate. 6. 24 h after transfection, change media and start selection with hygromycin at 150 μg/ml final concentration. 7. After 10 days pick up and expand colonies. Analyze SH targeting at the Rb1 locus by Southern blot (Subheading 3.2). 8. Collect 5  106 cells from correctly targeted clones, wash with PBS, resuspend in 700 μl PBS, and combine with 50 μg Cre expression vector [15] to remove hyg, thereby generating the S allele. 9. Pulse cells (250 V and 950 μF in a 0.4-cm cuvette) and plate cells at low density.

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10. Confirm loss of hyg marker by Southern blot using the neo gene probe. 11. Repeat the steps 1–10 on the S allele clones with the PH targeting vector to then derive the P allele, and thus the S/P reporter cell line. 3.2

Southern Blot

1. Prepare genomic DNA using a commercial kit. 2. Use 5 μg genomic DNA per digestion. In case of low-quality DNA, increase up to 10 μg.

3.2.1 Sample Preparation, Digestion, and Gel Run

3. Digest genomic DNA with EcoRI restriction enzyme to confirm correct integration at Rb1 locus (Fig. 2A) and with StuI or HindIII/StuI (Fig. 2B) for the removal of hyg. Incubate overnight at 37  C. 4. Run the digestion on a 0.8% agarose gel containing dye in 0.5 TBE enough to obtain good separation. Conditions may vary depending on the dimensions of the gel box. 1. Cut the wells off the gel and agitate the gel on a rocker for 1  15 min in depurination buffer: 12 N HCl.

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Fig. 2 Constructing ES cells with the S/P reporter. (A) Scheme for Southern blot strategy to confirm the correct integration of S2neo and Pneo genes at the Rb1 locus. Untargeted and targeted Rb1 alleles give two fragments of different sizes after EcoRI digestion. The probe for Rb1 gene is indicated in yellow and is located outside the targeting homology arms. (B) The SH and PH targeting vectors, containing the S2neo and Pneo genes, respectively, are sequentially targeted to the two Rb1 alleles on chromosome 14. After each round of targeting, the hyg gene is removed by Cre-mediated recombination, and its removal is confirmed by Southern analysis using StuI/HindIII digestion of genomic DNA and a neo probe

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2. Rinse gel twice with water to completely remove HCl and shake gel twice (1  15 min and 1  20 min) in alkaline transfer buffer: 0.5 M NaOH, 1 M NaCl. 3. Prepare three pieces of whatman paper to fit the gel and one piece longer so that it can hang over into the transfer buffer. Cut one piece of GeneScreen hybridization transfer membrane the size of the gel and soak in alkaline transfer buffer for 10 min. 4. Fill glass dish with alkaline buffer and put a glass plate on the top. Place the longer piece of filter paper across the glass with the edges dipping into the transfer buffer. 5. Place the gel on the paper and the membrane on top of the gel. Mark the orientation of the membrane with a pencil. 6. Wet one piece of filter paper in the transfer buffer and place it on top of the membrane. Remove bubbles between membrane and gel. 7. Add the remaining two pieces of filter paper and stack 10 cm of paper towels on the top. Cover with another glass dish and a small weight (empty 500 ml bottle). 8. Transfer overnight, disassemble the transfer to remove the membrane, and store the membrane dry. 3.2.3 Hybridization

1. Wet the membrane for 15 min in neutralization buffer: 0.5 M Tris–HCl pH 7.5, 1 M NaCl. 2. Dry the membrane on filter paper for at least 2 h. 3. In the meanwhile, prewarm the tube at 67  C and prepare hybridization solution: 6 ml SSC 20 (final 6), 2 ml Denhardt’s solution 50 (5 final), 1 ml 10% SDS (0.5% final). Warm at 67  C. Boil 100 μl salmon sperm DNA for 5 min and keep it on ice. 4. Rewet membrane with SSC 2. 5. Put the membrane into the tube with 10 ml hybridization solution and the boiled salmon sperm. Incubate in oven for at least 1 h at 67  C.

3.2.4 Probe Preparation

1. For the neo probe, digest 5 μg of the pMC1neo plasmid with HindIII and XhoI for 2 h at 37  C. Run the digestion on 0.8% agarose and purify the 1.1 kb band. Quantify and store at 20  C. Probes are stable in the purification buffer. The Rb1 probe is a 440 bp PstI-PvuII fragment from Rb1 intron 18 (GRCm38.p4, chr 14:73216633-73216194). 2. For one reaction, use 50 ng of probe and label accordingly to the kit used. In case of a random labeling kit from Stratagene, add 50 ng probe, 10 μl random 9-mer, and ddH2O up to 34 μl. If required, add 1000 natural diverged S. cerevisiae isolates has revealed a broad spectrum of nucleotide variation between haploids and heterozygosity among diploid strains ranging from ~337 heterozygous single-nucleotide polymorphisms (referred to in this chapter as HetSNPs) in the laboratory strain Sigma1278b to >40,000 HetSNPs in the clinical isolate YJM311 [11–13]. Natural heterozygosity can offer valuable insight into the life history of a strain. For example, our group has extensively characterized the genome of a wild isolate called JAY270, originally isolated from a bioethanol production facility in Brazil [14]. JAY270 is only moderately heterozygous (~12,000 HetSNPs) [15, 16]. These HetSNPs, however, are not evenly distributed across the genome (Fig. 1a). Rather, there are extended tracts of homozygosity (Fig. 1a), which span whole chromosomes (e.g., Chr1) and large sections of chromosome arms (Fig. 1a, distal left and right arms of Chr4). While we cannot directly infer the pattern and timing of recombination events that have shaped the genome of JAY270, it is tempting to speculate that the original heterozygosity of JAY270 has eroded over its life history due to LOH events resulting from mitotic recombination and meiotic recombination followed by mating between sibling spores [17]. The JAY270 genome illustrates the premise that in the context of a heterozygous background, we can “see” the outcomes of interhomolog recombination on a genome-

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A. JAY270 - industrial bioethanol strain: ~12,000 HetSNPs Chr 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 B. S288c/YJM789 - lab/clinical hybrid: ~60,000 HetSNPs Chr 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 Fig. 1 Levels and distribution of heterozygosity in natural and artificially created hybrid diploid S. cerevisiae strains. (a) Linear representation of all nuclear chromosomes in the JAY270 diploid strain [16] displaying the specific positions of HetSNPs. Each red/blue line indicates a HetSNP between the two homologous chromosomes. Regions that lack HetSNP lines are entirely homozygous. Black circles indicate the positions of centromeres. JAY270 contains ~12,000 HetSNPs. (b) Linear representation of each chromosome in the S288c/YJM789 hybrid diploid showing the high density and uniform distribution of its ~60,000 HetSNPs [3]

wide scale. However, because heterozygous markers are absent in several portions of the genome, this natural diploid only supports high-resolution characterization of new recombination in the genomic regions where it remains heterozygous.

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For many years now, homologous recombination (HR) and specifically LOH events have been studied using hybrid diploid strains that are highly heterozygous because they are formed by mating two evolutionarily diverged haploid parents [3, 5, 18]. These genetic backgrounds enable researchers to map recombination breakpoints and tracts of homozygosity with high resolution and have been instrumental in expanding our understanding of HR-based DNA repair [8]. Currently, our group uses a wellcharacterized hybrid diploid formed by mating the reference strain S288c [19, 20] to YJM789, a clinical isolate [21, 22]. The S288c/ YJM789 diploid has ~60,000 HetSNPs distributed evenly throughout the genome (Fig. 1b, [23], making it an excellent background for genome-wide characterization of recombination events. 1.2 Genome-Wide Assessment of Recombination Outcomes

The ability to characterize the outcomes of recombination on a genome-wide basis was made possible by the introduction of single-nucleotide polymorphism (SNP) genotyping microarrays and WGS technologies [8, 11, 13]. SNP microarrays were used to produce the first high-resolution maps of both mitotic and meiotic recombination events in the yeast genome [3, 5]. These arrays can map recombination events and other chromosomal rearrangements to a resolution of ~1 kb [8] depending on the array design. More recently, WGS has been used to map recombination events with higher resolution than is practical with SNP microarrays. When would one want to consider mitotic recombination events on a genome-wide scale? SNP microarrays and WGS analysis have been used to assess the global responses to DNA damaging agents such as ultraviolet radiation as well as to identify recombination-prone regions (e.g., fragile sites) that are stimulated by replication stress [24–29]. In addition, WGS analysis can be used to characterize multiple mitotic recombination and mutational events that co-occur in the genome simultaneously [30, 31]. In this chapter, we describe a general procedure used by our group to detect genome-wide LOH associated with mitotic recombination in budding yeast. In the specific type of assay described here, we start with a parent diploid strain that contains wellcharacterized HetSNPs distributed across the genome and at least one hemizygous counter-selectable marker. Selection for loss of the marker typically yields clones that contain an LOH tract that resulted from allelic interhomolog recombination in the chromosome arm where the marker was inserted. While LOH of the marker may also arise via other events such as point mutation, segmental deletion, and chromosome loss (see Note 1), the spontaneous rates of allelic interhomolog mitotic recombination are substantially higher than the rates of those rarer alternative events; so the majority (sometimes all) of the clones recovered by such selection regime are caused by this most frequent genetic rearrangement class. The procedure outlined below is able to distinguish between the causal mechanisms.

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Materials Into the S288c/YJM789 hybrid strain background, we have inserted the counter-selectable URA3 gene onto a distal position in the right arm of the S288c chromosome IV (Chr4) (Fig. 2, red homolog). Both endogenous copies of the URA3 gene were mutated or fully deleted from the native locus on Chr5. When a DSB lesion occurs at a proximal position on the S288c Chr4 homolog containing the URA3 insertion, it may be repaired by HR using the allelic position on the YJM789 homolog as template and be resolved as a crossover (Fig. 2). In this scenario, depending on the segregation of recombinant chromatids in the subsequent mitosis, one of the resulting daughter cells will lose the URA3 insertion, become resistant to 5-fluoroorotic acid (5-FOA), and form a colony on selective media (Fig. 2, FOAr) [32, 33]. Again, we note that this procedure may also yield at low frequency clones that became resistant to 5-FOA through mechanisms other that mitotic recombination (see Note 1). The same general experimental approach could also be pursued using other counter-selectable markers such as CAN1 and AmdS [34] integrated at other positions of the yeast genome.

2.1

Yeast Strains

1. Experimental hybrid diploid JAY2357: This diploid was obtained by mating the JAY2355 and JAY308 described below. 2. JAY2355: MATa, ura3-52, leu2Δ1, trp1Δ63, SSF2::CORE3, can1Δ::NatMX4. This strain was derived from FY23 [35], which is isogenic to the S288c strain background. 3. JAY308: MATα, ho::hisG, ura3, gal2. This strain is isogenic with the YJM789 strain background [23].

2.2

Culture Media

1. Permissive medium: YPD (1% yeast extract, 2% peptone, 2% dextrose, 2% bacteriological agar). 2. Selective medium: Synthetic complete medium (0.17% yeast nitrogen base without amino acids, 0.14% complete drop out mix, 0.5% ammonium sulfate, 2% dextrose, 2% bacteriological agar) supplemented with 1 g/L 5-FOA).

3

Methods

3.1 Isolation of Yeast Clones Carrying LOH Tracts

1. To prepare a culture from which to select clones that have experienced LOH of URA3, we streak cells of the above strain to a rich YPD plate at a low density that enables single colonies to grow in isolation. 2. Colonies are grown for 2 days at which point they are individually picked up with a sterile toothpick and inoculated into a

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DSB

Crossover

Chromosome Segregation

URA3+/- URA3+/FOAS FOAS

URA3-/- URA3+/+ FOAR FOAS

Fig. 2 Interhomolog allelic mitotic recombination leading to loss-ofheterozygosity (LOH) of a counter-selectable marker. The sequence of panels (from top to bottom) illustrates the steps of a DSB repair event by HR. Other possible scenarios such as gene conversion and break-induced replication are not shown. A pair of duplicated homologous chromosomes represent the S288c/ YJM789 hybrid background: S288c in red, YJM789 in blue. Black circles represent centromeres; a “U” labeled yellow circle represents the counterselectable URA3 marker which has been inserted at a distal region of the chromosome. One of the chromatids in the S288c homolog sustains a DSB. Repair of the DSB occurs through HR using as template the allelic position from a chromatid in the YJM789 homolog. The recombination intermediate is resolved as a crossover resulting in a reciprocal exchange of genetic information. In the subsequent mitosis, two possible outcomes of segregation of the parental and recombinant chromatids produce genetically distinct sets of daughter cells. Left: one daughter cell receives two parental chromatids while the other receives two recombinant chromatids. Both daughter cells remain heterozygous for the URA3 marker insertion and are thus 5-FOA sensitive although HetSNP phasing is

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tube with 5 mL of YPD to expand the culture during overnight incubation. 3. The following morning, each culture is appropriately diluted and plated on synthetic complete media supplemented with 5-FOA such that individual 5-FOA resistant colonies can be selected. 4. Plates are incubated at 30C until colonies are visible (~2–3 days). To ensure independence between clones, a single 5-FOA-resistant colony from each plated culture is then picked using a sterile toothpick and patched to a new 5-FOA plate in order to expand the clone under selective conditions. 5. Cells from this patch are inoculated into a tube with 5 mL of liquid YPD and grown overnight. 6. From the saturated culture above, an aliquot is frozen in glycerol, and the remaining cell pellet is used for genomic DNA isolation and subsequent library preparation for WGS. 1. Prepare libraries for sequencing. Most small labs do not have a next-generation sequencer, but the materials and equipment needed for library preparation are relatively simple and generally available. Thus, many users will prepare their own libraries and then ship them out for sequencing at a core facility or private vendor. Preparing libraries in-house significantly decreases the overall sequencing costs. However, initially, outsourcing both library preparation and sequencing may produce more consistent sequencing data output until the user becomes confident enough to prepare their own high-quality libraries. Protocols for genomic DNA isolation and library preparation of yeast genomes can be found at the following references 36, 37.

3.2 Whole-Genome Illumina Sequencing

2. Sequence data generation. Once libraries are prepared, they should be sequenced using an Illumina sequencing platform to generate reads of approximately 150 bp in length. The reads do not necessarily need to be pair-ended for the analysis described here although doing so can provide information with which to characterize structural genomic variation (see Note 2).

ä Fig. 2 (continued) altered in one of them. Right: each daughter cell receives one parental and one recombinant chromatid. One cell becomes homozygous for the S288c sequences at positions distal to the crossover site, including the URA3 marker insertion and remains sensitive to 5-FOA. The other daughter cell (bold) becomes homozygous for the YJM789 sequences, thus losing the URA3 marker and becoming resistant to 5-FOA

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3.3 Computational Analysis of Sequencing Data to Identify LOH Tracts

Although the availability and affordability of WGS technologies has greatly increased, the computational analysis of sequencing data remains a limiting step and is often intimidating to classically trained geneticists, biochemists, and cell and molecular biologists. We admit sharing this hesitancy and consider ourselves to be in that generation of colleagues who are not yet fully versed in computational methods. With this in mind, we describe a generalized workflow, using primarily commercial bioinformatics software packages that allow researchers with limited command line programming skills to process Illumina sequencing data in order to extract single-nucleotide variation information and ultimately identify LOH tracts in yeast clones. We describe each step of the analysis in broad terms as many of the specific actions will depend on the software used. We use the CLC Genomics Workbench proprietary software (Qiagen) in our laboratory due to its intuitive graphical user interface and broad sequencing-based applicability. However, comparable analyses can be achieved using other commercial packages and a number of open-source sequence read mappers such as BWA (https://sourceforge.net/projects/bio-bwa/) [38, 39] and Bowtie2 (http://bowtie-bio.sourceforge.net/ bowtie2/index.shtml) [40]. This workflow is designed to interrogate a specific list of genomic positions known to be heterozygous (HetSNPs) in the parent hybrid diploid strain from which a FOA-resistant LOH clone is derived. While the raw sequencing data also contains the information required to identify de novo point mutations, we do not discuss them here. In order to interrogate the specific sites mentioned above, the analysis requires that the user generate or have access to an accurate list of the reference genome coordinates and base variants of the majority of HetSNPs present in the parent diploid strain (in this protocol, we refer to this as the HetSNP map). For instance, we use a refinement of the HetSNP map for the S288c/YJM789 hybrid diploid reported previously [3]. In the case of experiments using the JAY270 strain, we generated and reported the HetSNPs map ourselves [16]. These HetSNP maps are then used to interrogate the heterozygous status of known positions in the genome and to filter out base variants detected at uncharacterized genomic positions. To detect, analyze, and identify LOH tracts in selected clones, sequencing reads are processed in the following manner (for schematic, see Fig. 3).

3.3.1 Importing Sequencing Reads Derived from a LOH Clone

Illumina reads are stored in FASTQ format, a file type that describes both the nucleotide sequence and the corresponding base calling quality scores. These files can be imported into any of the above read mapping programs. During import, information about the availability of paired ends is included. Paired end reads should be imported as such even if the matching read information will not be used in the downstream steps.

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Map Reads

very high coverage

very low coverage

Detect all variants

Interrogate HetSNP map

Call regions of LOH

homozygous ‘non-reference’

homozygous ‘reference’

Fig. 3 Summary of the approach used to identify tracts of LOH. Sequencing reads are mapped to the S288c reference genome (red) and regions that are excessively over- or underrepresented are identified and excluded from further analysis. Next, sequence variants are detected with relaxed stringency. Variants that remained heterozygous are shown as red/blue dashed lines. Variants that have become homozygous are shown as either solid red or solid blue lines. Following low-stringency variant detection, specific allele frequency data are interrogated and retrieved for only HetSNP positions known to exist in the parent diploid. The remaining variants are filtered out and not considered further. Finally, LOH calls are made based on the allele frequencies at the HetSNP positions

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3.3.2 Mapping Reads to the Reference Genome

Once imported, sequencing reads from a LOH clone can be mapped to the reference genome. This process assigns each read to a corresponding location in the reference genome. Optional inclusion of paired-end read information may be taken into consideration at this step although it will not influence the primary LOH analysis described here. The reference S. cerevisiae genome from the haploid strain S288c is available for download on the Saccharomyces Genome Database (SGD, https://www.yeastgenome.org) [9]. We use the most recent version (released 01-31-2015) although it is essential to keep the reference genome version specifically coordinated with the strain’s HetSNPs map (see Note 3). Sequencing reads spanning repetitive regions may map to more than one genomic site. Due to this ambiguity, these reads should be ignored during the downstream analysis. In addition, some reads may fail to map to any portion of the S288c reference genome. This may be because they are unique to the nonreference parent of the hybrid strain (i.e., YJM789) because they are of poor quality or because they originate from a contaminating source. These data are ignored in the analysis described here because the HetSNP map does not contain any bases present in the mitochondrial chromosome or at repetitive elements or within regions of structural variation.

3.3.3 Identification of All Sequence Variants

Once reads derived from a LOH clone have been mapped onto the reference genome, a software function is run to detect the presence of sequence variants between the reads and the reference genome. In our case, we use CLC Genomics Workbench to detect these variants, and we keep the following variables in mind and adjust detection parameters accordingly. In general, the parameters that we set for variant detection are quite liberal, in the sense that even minor frequency differences (resulting from sequencing errors or mis-mapped reads) will be tolerated and populated into a long variant detection output table (VDOT) containing all detected variants. This is not a concern because we ultimately filter this table to extract data only for the highly specific HetSNPs map. The use of such liberal parameters for variant detection ensures that positions in the HetSNP map that remained heterozygous in the LOH clone have a high probability of being detected and populated in the VDOT. Our liberal settings enable us to detect variants at these positions even in cases where coverage was relatively low, or if the measured allele frequency deviated too far from 50% due to stochasticity in the sequencing library. These are the parameters we typically use for variant detection: 1. Fixed ploidy SNP detection: This parameter enables variant detection with the assumption that the genome of the sequenced LOH clone is diploid. This is useful for initial automated prediction of LOH in CLC but is not strictly necessary

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for the interrogation of HetSNP positions described below. Furthermore, and as we discuss below, the assumption that the experimentally derived LOH clone is in fact diploid is not always correct. Copy number analysis should be conducted to validate and supplement the sequence-based genotype calls. 2. Discard positions with excessively high coverage: As stated above, some genomic segments are present at multiple copies in the genome or share high similarity to other sequences (e.g., transposable elements, rDNA, telomeric and subtelomeric sequences). Reads can and will often be erroneously mapped, resulting in artificially high coverage at these repetitive regions. Because it is difficult to confidently call variants in these regions, these over-represented sequences should be ignored from the subsequent analysis. To determine the upper coverage limit value, we first examine the read mapping file and evaluate the median coverage for the whole genome and for specific nonrepetitive regions. We then set the high coverage cutoff at 5–8
the median coverage. 3. Set the minimum read coverage: Just like over-represented sequences, under-represented sequences can also challenge the confidence of variant detection. Thus, we typically exclude sequences/regions that are present at less than ~20% the median coverage. 4. Set the minimum variant count for detection at 1: When performing sequence analyses of clonal derivatives of heterozygous diploids (e.g., a clone that was selected for LOH), three classes of genotypes need to be identified for each HetSNP position in the parent strain: A/B, loci that remained heterozygous in the experimentally derived LOH clone (frequency of nonreference nucleotide [i.e., YJM789] ~50%); B/B, loci that became homozygous for the nonreference nucleotide (frequency of nonreference nucleotide >95%); and A/A, loci that became homozygous for the reference nucleotide (frequency of nonreference nucleotide 50%) of reads differ specifically from the reference and are rarely missed during the SNP variant detection. A/A tracts of homozygosity are more challenging to call because the majority of reads and, occasionally all of them, will match to the reference genome and will not be detected as a variant position and thus will not be populated into the VDOT. In order to boost our ability to identify A/A tracts of homozygosity, we set an extremely relaxed stringency parameter for SNP variant detection (minimum variant count ¼ 1). As discussed above, this parameter will allow many genomic positions to be included in the VDOT, including variant nucleotides that arose from amplification or base calling errors during

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sequencing. However, because we have a specific HetSNP map for all heterozygous positions present in the parental genome, we are able to recognize these artifacts and interrogate the genomic positions of interest to determine that they have become homozygous for the reference. For instance, consider a heterozygous genomic position in which the reference base is T and the nonreference base is A. If this site is heterozygous, then ~50% of the reads mapped to that position will contain T and the other 50% will contain A. However, if this site was mapped with similar coverage as the rest of the genome, and 97% of the reads contained T and 3% contained a C (which is neither the reference nor the nonreference base), we would conclude that this position had become homozygous for the reference base. If instead of 97%, 100% of the reads matched the reference base at that site, it would not be included in the VDOT. While we can indirectly infer that this type of site had likely become homozygous by examining the genotypes at neighboring HetSNPs, we also take the following additional measures to ensure that we correctly call the homozygous reference region. First, we always conduct a simple visual inspection of the read mapping file at the HetSNP positions that were called homozygous reference by an indirect inference. Second, we can map the same sequencing reads to the nonreference genome (e.g., to YJM789 instead of S288c; see Note 4). In doing so, genotype classes A/B and A/A are called directly whereas genotype B/B is called through indirect inference. Combining and cross-referencing the analyses based on the two independent read mappings to each haploid parent reference genome remove any ambiguity and enhance the resolution of genotype calls to equal all positions on the HetSNP map. 5. Run the variant detection function: Once the above parameters are specified, the software will analyze the whole read mapping file and create a VDOT containing each detected polymorphic position, the chromosome and nucleotide coordinate of the reference base, the identity of the reference base itself, the identity of the variant bases detected at that position, and the total coverage and specific number of reads displaying each of the base variants. This table also contains lines that show small insertions and deletion polymorphisms (indels) of 1 or a few bases between the two haploid genomes that make up the hybrid diploid. In some cases, this information can be used later to refine the resolution of LOH tract breakpoints but will not be used initially because the HetSNP map contains only single base variants.

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1. The variant detection procedure described above outputs a long VDOT containing a low stringency list of variants between the sequencing reads and the reference genome. This table can be exported as a Microsoft Excel spreadsheet where it is most convenient to perform the final formatting and LOH tract identification using built-in Excel functions. First, we refine the VDOT to isolate only the genomic positions present in the HetSNP map for the parental strain. Specifically, we use Excel’s VLOOKUP function to retrieve only data from the VDOT rows corresponding to the coordinates in the HetSNP map specific to the hybrid diploid used. To identify VDOT rows containing known HetSNPs, we use a unique search term called a MarkerID that consists of the chromosome number, the underscore symbol, and the nucleotide coordinate within the reference chromosome (e.g., Chr03_157963; Chr14_75492). As discussed above, genomic positions that have become 100% homozygous for the reference base will not be detected and will not be present in the VDOT. Thus, the VLOOKUP approach will not be able to identify a row specifying nucleotide information for that genomic position. Those positions are relatively uncommon when coverage is higher than 50–60
. The genotype calls in those cases are made initially through indirect inference, followed by independent positive validation as described above. 2. Once retrieved, the base count data for the HetSNP positions are used to calculate overall coverage and individual base frequencies in the reads. Those data are used to make the genotype calls at each HetSNP. 3. Positions that remained heterozygous (A/B) in the sequenced clone have 40–60% of reads that match the reference genome. Higher sequencing coverage will narrow this distribution closer to ~50%. Cases where the frequency of the reference base is close to 33% or 66% indicate a possible copy number gain from two copies in the parent strain to three copies in the sequenced clone (see Subheading 3.3.5 below). 4. Positions that became homozygous for the nonreference base (B/B) or that simply lost the reference base through a deletion (B/) have 95% that match the nonreference base. 5. Positions that became homozygous for the reference base (A/A) or that simply lost the nonreference base through deletion (A/) have >95% of reads that match the reference genome. In cases where 100% of reads match the reference base, the call needs to be made by indirect inference (see Note 5).

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3.3.5 Copy Number Analysis

The HetSNP analysis described above is not sufficient to differentiate between copy-neutral LOH (e.g., crossover type; Fig. 2) and LOH caused by segmental deletion or chromosome loss. It is essential to also take into consideration the depth of read coverage at the regions of genotypic change in order to unambiguously distinguish LOH mechanisms. For example, in cases where the above HetSNP analysis identified genotypes A/A or A/, the copy-neutral A/A call would likely be made initially because the rate of mitotic recombination leading to copy-neutral LOH is usually 10–100
higher than the rates of either segmental deletions or chromosome loss (Fig. 4b; [1, 10]. However, these two possible genotype alternatives can be distinguished through the analysis of the read depth coverage information. The example shown in Fig. 4b, which illustrates how an initial interpretation of copy neutral gene conversion (A/A), can be refined to a segmental deletion call after copy number analysis has been integrated into the inference. In this example, CNV analysis shows a ~50% reduction in depth coverage coinciding with the LOH tract. Similarly, cases where LOH of an entire chromosome is identified using HetSNP analysis might be initially interpreted as a case of chromosome loss (Fig. 4c). However, CNV analysis may show that coverage for that chromosome is similar to the coverage for the whole diploid genome, indicating that there are in fact two copies of the remaining homolog. Thus, instead of calling LOH due to monosomy (single copy of a chromosome), the call is refined to uniparental disomy. Before the broad dissemination and cost reduction of next generation sequencing, genome-wide CNV analysis would be primarily obtained using microarray-based competitive genomic hybridization (aCGH). This technique, which is still useful and remains in use, exploits the relative annealing of genomic DNA from an experimental clone and from a reference strain to tiled arrays of oligonucleotides, each from a defined genomic position, to estimate the number of copies at which that sequence is present in the clone of interest [41, 42]. When WGS data is available, analogous relative copy number information can be extracted by examining trends in read depth coverage along the genome. For researchers like us with limited coding skills, proprietary software such as Nexus Copy Number (BioDiscovery) provide powerful yet accessible genome-wide CNV determination and visualization using BAM read mapping files as input. When using this approach, the sequencing coverage of the parental diploid strain is used as a reference for normalization. Pairwise coverage comparison between the read depth in the experimental LOH clone and in the parental strain identifies under-, neutral, or overrepresented gene dosage regions. The output of CNV calls from Nexus Copy Number includes specific coordinates where the copy number changes occurred and that information is then compared to the

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B. Example 1

C. Example 2

gene conversion

chromosome loss

segmental deletion

uniparental disomy

SNP analysis only

Initial Inference

CNV and karyotype analysis Complete Inference

Fig. 4 CNV analysis is used in conjunction with HetSNP analysis to refine inferences of genotype and genome structure. (a) First, use HetSNP analysis to identify LOH tracts. This alone can lead to an incomplete inference of genotype and genome structure. Next, perform CNV and karyotype analysis. With these additional data, refined inferences of genotype and genome rearrangement mechanisms can be made. (b) Example 1: HetSNP genotyping alone would suggest that this clone has become homozygous for a region of red homolog. However, CNV analysis detects a ~50% lower coverage in the region, indicating that that this LOH genotype is in fact caused by a segmental deletion on the blue homolog. (c) Example 2: HetSNP genotyping would suggest that this clone has become monosomic due to loss of the red homolog. CNV analysis detects coverage similar to the median for rest of the genome, indicating that the clone is actually disomic for the blue chromosome (uniparental disomy)

HetSNP genotype calls to refine the assessment of genomic changes in the experimental clones (Fig. 4). Supplementation of these analyses with physical chromosome length measurement by pulse-field gel electrophoresis (PFGE) karyotyping can also be used to further characterize changes in chromosome structure and aneuploidy.

4

Notes 1. Alternative mechanisms of 5-FOA resistance: Most of the clones resistant to 5-FOA isolated through the procedure described above lose the entire URA3 marker insertion through a mitotic recombination event leading to homozygosis at the corresponding region of the homolog lacking URA3. However, clones that became 5-FOA resistant through non-crossover mechanisms can also be recovered at low frequency. These can include segmental deletions and whole chromosome loss similar to the examples shown in Fig. 4 as well as

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point mutations that inactivate the URA3 gene without affecting heterozygosity or copy number in the region. Each of these possibilities can be specifically discerned through the combination of HetSNP genotyping and CNV analysis, supplemented by PFGE, PCR, and Sanger sequencing. 2. Whether one chooses to generate single or paired end reads, at least for the analysis described here, will come down to the cost of library preparation kits and sequencing. In order to reliably call LOH at HetSNPs, we aim to produce an overall read coverage of between 60
and 120
. In our experience, coverage of 90
supports optimal downstream analysis whereas coverage above 120
has diminishing returns for the type of LOH analysis described here. Multiplexing approaches appropriate for the specific sequencing instrument’s data generation capacity should be used in order to achieve these coverage targets while maximizing the number of LOH clones analyzed per sequencer lane and minimizing cost. 3. Strain-specific HetSNP map refinement and reference genome version: The analysis above relies on a map of HetSNPs between the two parent haploids used to create a hybrid diploid. These lists do not necessarily need to, and indeed should not, include every possible position where the two haploids differ. While such a complete HetSNP map would provide maximal resolution for the characterization of LOH tracts, it is best to use a conservative HetSNP map that contains only single-base variants within unique DNA sequences. One should avoid positions that are vulnerable to read mapping errors, such as repetitive elements and homopolymer runs. Such refinement of the HetSNPs map might reduce resolution by ~10% of the actual variation between the haploids, but it substantially improves the reliability and reproducibility of the genotype calls. A good way to optimize a HetSNP map is to generate control sequencing reads directly from each of two the haploid parents as well as from the hybrid diploid and then run those control data through multiple iterations of the procedure above. Haploid control reads should always return the expected single genotype calls, and hybrid diploid control reads should always return heterozygosity for all positions in the HetSNP map. The positions that do not always conform to these strict expectations should be removed from the HetSNP map to improve the reliability of genotype calls made from reads derived from the experimental clones. Once this high confidence HetSNP map is developed, or if it is obtained from prior publications, it is essential to have the genomic coordinates in the map match the reference genome version that is used in the read mapping step. Each time the reference genome is updated [9], there are usually minor changes in the

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coordinates for each nucleotide between versions. If the coordinates in the HetSNP map are from an older version of the reference genome, they can be corrected to match the latest genome release, or the corresponding older version of the reference genome should be downloaded and used for read mapping. 4. Mapping reads to the nonreference genome: A good way to resolve ambiguities that may emerge from indirect calling of regions of homozygosity for the reference genome base (e.g., the A/A genotype) is to use the orthogonal approach of mapping the sequencing reads to the nonreference genome. In doing so, the same positions (those that contain the reference base) will now be detected as variants relative to the nonreference genome. In principle, these two independent read mappings should be used. However, in practice, mapping reads to a nonreference genome is more challenging because the assembly of non-S288c genomes, including the YJM789 genome [23], is not as contiguous and complete. Instead, these diverged genomes are usually available for download as a collection of contigs that are much more numerous and smaller than the 16 nuclear chromosomes assembled for the S288c genome. Therefore, interpreting the mapping of reads to a nonreference genome requires the additional step of generating a good alignment between the reference and nonreference genome assemblies, and deriving a set of corresponding coordinates between the two. A reasonable, but effective, alternative approach is to conduct nonreference read mappings primarily for specific genomic segments that are assembled into longer contigs of the nonreference genome, and for situations where the homozygous reference LOH call (A/A) cannot be independently validated by direct visualization of mapped reads to the S288c reference genome assembly. 5. Mitotic recombination often affects continuous segments of the genome rather than individual bases independently. Because these events typically affect multiple neighboring HetSNPs within the same recombination tract, we are able to enhance the confidence of genotype calls by placing them in the context of a regional segment. When we make an indirect inference of the A/A genotype, it is helpful to assess the heterozygosity of HetSNPs surrounding that individual position. Ultimately, though, these regions should be reviewed by visually examining the mapped sequencing reads, and in some cases, confirmation of a genotype may require validation using PCR and Sanger sequencing.

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Acknowledgments Mitotic recombination and general genome stability research in the Argueso laboratory was supported by NIH grant R35GM119788 to JLA. References 1. Klein HL, Bacˇinskaja G, Che J et al (2019) Guidelines for DNA recombination and repair studies: cellular assays of DNA repair pathways. Microb Cell 6:1–64 2. Gerton JL, DeRisi J, Shroff R et al (2000) Global mapping of meiotic recombination hotspots and coldspots in the yeast Saccharomyces cerevisiae. Proc Natl Acad Sci U S A 97:11383–11390 3. Mancera E, Bourgon R, Brozzi A et al (2008) High-resolution mapping of meiotic crossovers and non-crossovers in yeast. Nature 454:479–485 4. Pan J, Sasaki M, Kniewel R et al (2011) A hierarchical combination of factors shapes the genome-wide topography of yeast meiotic recombination initiation. Cell 144:719–731 5. St Charles J, Petes TD (2013) High-resolution mapping of spontaneous mitotic recombination hotspots on the 1.1 Mb arm of yeast chromosome IV. PLoS Genet 9:e1003434 6. Laureau R, Loeillet S, Salinas F et al (2016) Extensive recombination of a yeast diploid hybrid through meiotic reversion. PLoS Genet 12:e1005781 7. McGinty RJ, Rubinstein RG, Neil AJ et al (2017) Nanopore sequencing of complex genomic rearrangements in yeast reveals mechanisms of repeat-mediated double-strand break repair. Genome Res 27:2072–2082 8. Zheng DQ, Petes TD (2018) Genome instability induced by low levels of replicative DNA polymerases in yeast. Genes (Basel) 9(11):539 9. Cherry JM, Hong EL, Amundsen C et al (2012) Saccharomyces genome database: the genomics resource of budding yeast. Nucleic Acids Res 40:D700–D705 10. Symington LS, Rothstein R, Lisby M (2014) Mechanisms and regulation of mitotic recombination in Saccharomyces cerevisiae. Genetics 198:795–835 ¨ , Granek JA et al 11. Magwene PM, Kayıkc¸ı O (2011) Outcrossing, mitotic recombination, and life-history trade-offs shape genome evolution in Saccharomyces cerevisiae. Proc Natl Acad Sci U S A 108:1987–1992

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Genome-wide Analysis of LOH 24. Lemoine FJ, Degtyareva NP, Lobachev K et al (2005) Chromosomal translocations in yeast induced by low levels of DNA polymerase a model for chromosome fragile sites. Cell 120:587–598 25. Lemoine FJ, Degtyareva NP, Kokoska RJ et al (2008) Reduced levels of DNA polymerase delta induce chromosome fragile site instability in yeast. Mol Cell Biol 28:5359–5368 26. St Charles J, Hazkani-Covo E, Yin Y et al (2012) High-resolution genome-wide analysis of irradiated (UV and γ-rays) diploid yeast cells reveals a high frequency of genomic loss of heterozygosity (LOH) events. Genetics 190:1267–1284 27. Song W, Dominska M, Greenwell PW et al (2014) Genome-wide high-resolution mapping of chromosome fragile sites in Saccharomyces cerevisiae. Proc Natl Acad Sci U S A 111:E2210–E2218 28. O’Connell K, Jinks-Robertson S, Petes TD (2015) Elevated genome-wide instability in yeast mutants lacking RNase H activity. Genetics 201:963–975 29. Zheng DQ, Zhang K, Wu XC et al (2016) Global analysis of genomic instability caused by DNA replication stress in Saccharomyces cerevisiae. Proc Natl Acad Sci U S A 113: E8114–E8121 30. Sampaio NMV, Rodrigues Prause A, Ajith VP et al. (2017) Mitotic systemic genomic instability in yeast. bioRxiv 161869 31. Sakofsky CJ, Saini N, Klimczak LJ et al (2019) Repair of multiple simultaneous double-strand breaks causes bursts of genome-wide clustered hypermutation. PLoS Biol 17:e3000464 32. Boeke JD, LaCroute F, Fink GR (1984) A positive selection for mutants lacking orotidine-50 -phosphate decarboxylase activity in

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yeast: 5-fluoro-orotic acid resistance. Mol Gen Genet MGG 197:345–346 33. Boeke JD, Trueheart J, Natsoulis G et al (1987) 5-Fluoroorotic acid as a selective agent in yeast molecular genetics. Methods Enzymol 154:164–175 34. Solis-Escalante D, Kuijpers NG, Bongaerts N et al (2013) amdSYM, a new dominant recyclable marker cassette for Saccharomyces cerevisiae. FEMS Yeast Res 13:126–139 35. Winston F, Dollard C, Ricupero-Hovasse SL (1995) Construction of a set of convenient Saccharomyces cerevisiae strains that are isogenic to S288C. Yeast 11:53–55 36. Wilkening S, Tekkedil MM, Lin G et al (2013) Genotyping 1000 yeast strains by nextgeneration sequencing. BMC Genomics 14:90 37. Baym M, Kryazhimskiy S, Lieberman TD et al (2015) Inexpensive multiplexed library preparation for megabase-sized genomes. PLoS One 10:e0128036 38. Li H, Durbin R (2009) Fast and accurate short read alignment with burrows-wheeler transform. Bioinformatics 25:1754–1760 39. Li H, Durbin R (2010) Fast and accurate longread alignment with burrows-wheeler transform. Bioinformatics 26:589–595 40. Langmead B, Salzberg SL (2012) Fast gappedread alignment with bowtie 2. Nat Methods 9:357–359 41. Argueso JL, Westmoreland J, Mieczkowski PA et al (2008) Double-strand breaks associated with repetitive DNA can reshape the genome. Proc Natl Acad Sci U S A 105:11845–11850 42. Zhang H, Zeidler AFB, Song W et al (2013) Gene copy-number variation in haploid and diploid strains of the yeast Saccharomyces cerevisiae. Genetics 193:785–801

Chapter 16 Monitoring Gene Conversion in Budding Yeast by Southern Blot Analysis Miyuki Yamaguchi and James E. Haber Abstract By using an inducible site-specific double-strand break (DSB) in budding yeast, it is possible to monitor—in real time—the repair of the break by homologous recombination. A method is described using an ectopic homologous donor sequence to repair an HO endonuclease-induced DSB. These gene conversion events can occur with or without crossing-over, the products of which are distinguished as different-sized restriction endonuclease fragments. The method of Southern blotting is described in detail. Key words Saccharomyces cerevisiae, Budding yeast, Double-strand break repair, Gene conversion, Crossing-over, Southern blot

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Introduction The kinetics and outcomes of double-strand break (DSB) repair can be monitored in S. cerevisiae strains in which synchronously induced site-specific cleavage is repaired by homologous recombination. Here we describe the analysis by Southern blotting of a DSB induced by a galactose-inducible HO endonuclease gene (GAL::HO) which can be carried on a replicating plasmid [1, 2] or integrated into the ADE3 locus [3]. This approach has been used to analyze several modes of homologous recombination: singlestrand annealing, break-induced replication, and (as shown here) gene conversions both with and without an accompanying crossover. The details of these mechanisms of repair are reviewed by Haber [4]. Cleavage of the 24-bp HO recognition site is >90% complete within 30 min of induction; so it is possible to monitor several intermediate steps—by PCR, chromatin immunoprecipitation, and other techniques [4, 5]. A similar approach is possible with other site-specific endonucleases such as I-SceI [6] or CRISPR/Cas9 [7].

Andre´s Aguilera and Aura Carreira (eds.), Homologous Recombination: Methods and Protocols, Methods in Molecular Biology, vol. 2153, https://doi.org/10.1007/978-1-0716-0644-5_16, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Materials Prepare all solutions using distilled water and analytical-grade reagents. All reagents may be prepared and stored at room temperature (unless otherwise indicated). All commercial reagents should be used and stored following manufacturer recommendations. All buffers and solutions used in standard molecular biology techniques should be prepared using autoclaved stock solutions unless otherwise noted.

2.1

Mating-type gene (MAT) switching is carried out in strain MY270 or MY385d (ho HMLα-inc MATa hmrΔ::ADE1 ade1 leu2,3-112 leu2-3,112 lys5 trp1::hisG ura3-52 bar1Δ::TRP1 ade3::GAL::HO) in which the HMLα-inc locus carries a single-base mutation that prevents the product of DSB repair—MATα-inc—from being cleaved by HO; consequently these strains switch once, from MATa to MATα-inc (Figs. 1a and 2a). Repair by ectopic recombination is shown (Figs. 1b and 2b) for strain tGI354 [8] (ho hmlΔ∷ADE1 MATa-inc hmrΔ∷ADE1 arg5,6∷MATa∷HPH ade1 leu2-3,112 lys5 trp1∷hisG ura3-52 ade3∷GAL∷HO in which both HML and HMR are deleted. A cloned segment carrying MATα is introduced on chromosome V and is the recipient of a gene conversion event in which the normal MAT locus carries an uncleavable MATa-inc mutation.

Strains

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Fig. 1 Southern blots of HO-induced recombination. (a) Southern blot of MAT switching using StyI digest and MATdistal probe. (b) Southern blot of ectopic gene conversion with crossover and noncrossover products of strain tGI354, digested with EcoRI and analyzed with a MAT-Ya/Z1 probe

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Fig. 2 Recombination events induced by HO endonuclease. (a) Diagram of MAT switching with map of StyI sites (S) and probe. The MATdistal probe is immediately distal to MAT-Z2. This MATdistal probe recognizes a 0.9 kb StyI restriction fragment in MATa cells. Upon galactose induction, HO endonuclease is expressed and creates a double-strand break at the border between Ya and Z1. This cleavage results in a 0.7 kb fragment recognized by the MATdistal probe. Upon MAT switching, the sequence at Ya is replaced with Yα, resulting in the loss of the original StyI site and the MATdistal probe now recognizes a 1.9 kb StyI restriction fragment. (b) Map of EcoRI sites in tGI354 around the MAT locus on chromosome III and the MAT locus inserted at ARG5,6 on chromosome V. HO endonuclease cleaves the HO cut site (HOcs) at chromosome V. Ectopic gene conversion will occur, using the MATa-inc locus on chromosome III as a template. Triangles indicate EcoRI sites. Sizes of EcoRI restriction fragments are labeled. Intact parent chromosomes before galactose induction are pictured on the left. Ectopic gene conversion products and crossover products are pictured on the right

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2.2 Media for Time Course

1. YEP-lactate media (see Note 1): 1% yeast extract, 2% Bacto peptone, 0.004% adenine, 2.7% lactic acid (85%) (final pH 5.5). 2. YEPD media (see Note 2): 1% yeast extract, 2% Bacto peptone, 0.004% adenine, 2% dextrose (final pH 5.5). 3. YEPD plates (see Note 3): 1% yeast extract, 2% Bacto peptone, 0.004% adenine, 2% dextrose, 2.5% agar (final pH 5.5). 4. YEP-gal plates (see Note 4): 1% yeast extract, 2% Bacto peptone, 0.004% adenine, 2% galactose, 2.5% agar (final pH 5.5). 5. 20% galactose (w/v) (filter-sterilized) (see Note 5). 6. 20% dextrose (w/v) (filter-sterilized) (see Note 6). 7. 50 ml sterile disposable conical tubes. 8. Bucket of dry ice. 9. Autoclaved glass beads (3 or 4 mm) (Fisher).

2.3 Genomic DNA (gDNA) Preparation

1. Extraction buffer: 100 mM Tris pH 8.0, 50 mM EDTA, 2% SDS (filter sterilized). 2. Glass beads, acid-washed 425–600 μm (30–40 U.S. sieve) (Millipore Sigma). 3. Ultrapure phenol:chloroform:isoamyl alcohol 25:24:1 (ThermoFisher Scientific/Invitrogen). 4. Vortex-Genie (Scientific Industries). 5. Chloroform. 6. 3 M sodium acetate, pH 5.2. 7. Isopropanol. 8. 70% ethanol. 9. RNase A: RNase A 10 mg/ml (Millipore Sigma) in 10 mM Tris–Cl (pH 7.5), 15 mM NaCl. 10. TE: 10 mM Tris pH 7.6, 1 mM EDTA. 11. Speed-Vac vacuum concentrator.

2.4 Restriction Digest and Gel Electrophoresis for Southern Blot

1. Restriction enzyme of choice (New England Biolabs). 2. 3 M sodium acetate, pH 5.2. 3. Isopropanol. 4. 70% ethanol. 5. 6
loading buffer: 0.25% bromophenol blue, 0.25% xylene cyanol, 30% glycerol. 6. 1
TBE: 89 mM Tris–borate, 89 mM boric acid, 2 mM EDTA (see Note 7). 7. Speed-Vac vacuum concentrator.

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1. 0.25 M HCl. 2. 0.4 M NaOH. 3. NEN GeneScreen Plus (PerkinElmer). 4. Whatman 3MM chromatography paper. 5. Stacks of paper towels (multifold). 6. Glass tray. 7. Plexiglass support.

2.6 Radiolabeling Probe

1. Hexanucleotide mix (10
) (Millipore Sigma/Roche). 2. 5
random primer buffer: 5
hexanucleotide mix, 125 μM dGTP, dCTP, dTTP. 3. DNA polymerase I, large (Klenow) fragment; 5000 units/ml (New England Biolabs). 4. dATP, [α-32P]-3000 Ci/mmol 10 mCi/ml Easy Tide, 250 μCi (PerkinElmer). 5. Micro Bio-Spin 30 columns in Tris buffer (BioRad). 6. QIAquick PCR purification kit (Qiagen).

2.7 Hybridization and Washing Membrane

1. Hybridization buffer: 0.25 M sodium phosphate pH 7.2 (see Note 8), 7% sodium dodecyl sulfate, 1 mM EDTA. 2. Buffer 2: 1% sodium dodecyl sulfate, 20 mM sodium phosphate (pH 7.2) (see Note 8), 1 mM EDTA. 3. Buffer 1: 5% sodium dodecyl sulfate, 20 mM sodium phosphate (pH 7.2) (see Note 8), 1 mM EDTA. 4. Hybridization tube(s) and hybridization oven.

2.8 Stripping Membrane

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1. Stripping buffer: 2% SDS, 0.1
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Methods

3.1 Cultures for Time Course

Cultures are initially grown in YEPD media (see Note 10) and then transferred to lactose-containing media for GAL::HO induction. GAL::HO expression is under the control of the GAL1-10 promoter [9]. Upon galactose induction, samples are collected at the desired time points and stored frozen at -80  C. 1. Streak out strain of interest onto YEPD plate from frozen stock to obtain single colonies. Let grow for 2 days at 30  C. 2. Patch single colonies on YEPD plates. Let grow overnight at 30  C. 3. Replica plate onto various drop-out plates to confirm the strain phenotype (see Note 11). Also replica plate onto YEPEG plate

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to confirm that the strain is not petite (see Note 12). If working with mating-type switching strains, confirm mating type before inoculating (see Note 13). 4. Inoculate 5 ml YEPD with an isolate. Grow at 30  C on roller drum for approximately 8 h or overnight. 5. Spin down culture at low speed (40% (see Note 14). 22. From a height of around 2–3 feet from the bench, aliquot three drops (~30 μl per drop) of the final cell suspension onto a cleaned and polished glass slide using a glass pipette. Tilt the slide at about 30–45 from the horizontal. Do not let successive drops overlap one another (see Note 14). 23. Prepare as many slides as required and let them air-dry for ~10 min. The remaining cell suspension can be stored at 20  C and used for several days. The unstained slides can be stored at 4  C for at least 1 year. 24. Add 20 μl ProLong Gold antifade reagent containing 5 μg/ml DAPI per slide and mount with a cover glass. 25. Seal the slides with clear nail polish and image via epifluorescence microscopy (Fig. 5).

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Fig. 3 Layout of dishes for oocyte culture. Cartoons depicting the layout of media and solution droplets utilized, respectively, for oocyte collection, washing, culture, and removal of zona pellucida (ZP) 3.4 Metaphase-I Chromosome Spreads from Cultured Oocytes

1. Prepare four 35 mm dishes containing drops of M2 media, Tyrode’s solution, and PBS as shown in Fig. 3. Fully cover each drop with mineral oil to prevent evaporation. Prewarm these dishes to 37  C. These dishes are used, respectively, for the collection, washing and culture of oocytes, and the removal of zona pellucida (ZP; Fig. 3) (see Note 15). 2. Euthanize adult female mice by carbon dioxide inhalation followed by cervical dislocation. Dissect out the ovaries and place in the collection dish, into the prewarmed M2 medium drop supplemented with milrinone (see Note 16). 3. Under a stereo microscope, puncture mature antral follicles on the ovary surface using a 25-gauge needle until no antral follicles are visible. Oocytes are released into the medium. Use fine forceps to hold and manipulate the ovaries within the center of the droplet. 4. Discard the ovary tissue. 5. Let the oocytes settle for 2–3 min. 6. Using the mouth-pipette apparatus, select and transfer fully grown germinal-vesicle (GV) stage oocytes, surrounded by multiple layers of cumulus cells, to a peripheral part of the droplet. Repeat until all GV oocytes have been collected (see

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Fig. 4 Oocyte collection and culture. Representative bright-field images showing (a) cumulus cell-oocyte complexes (COCs), (b) germinal vesicle (GV) stage oocytes, (c) metaphase-I oocytes, and (d) ZP-free oocytes. Scale bars represent 100 μm

Note 17). A selected cumulus cell-oocyte complex (COC) is shown in Fig. 4a. 7. Mechanically remove the surrounding cumulus cells by sucking oocytes in and out of the pipette tip (see Note 18). 8. Wash off the milrinone and remaining cumulus cells by sequentially transferring oocytes through the four drops of M2 media in the washing dish (see Note 19). 9. Transfer oocytes into the M2 medium in a culture dish. 10. For in vitro maturation, the culture dish is placed in the dark on a heat block at 37  C. GV stage oocytes prepared for culture are shown in Fig. 4b. 11. After 3 h of culture, the oocyte GVs should have disappeared. Discard any oocytes remaining at the GV stage (see Note 20). 12. Metaphase-I oocytes are obtained after around 7 h of culture (see Fig. 4c; Note 20). Transfer oocytes into the first M2 medium droplet of the ZP removal dish. 13. Transfer up to 10 oocytes at a time into the first acid droplet to briefly wash and then rapidly transfer into the second acid droplet for ZP removal (see Note 21). 14. Continually monitor the oocytes for ZP removal and then immediately transfer ZP-free oocytes through the second M2 medium droplet and then into the third M2 droplet (see Note 22). 15. Repeat the previous step until all oocytes have undergone ZP removal. ZP-free oocytes are maintained in the third M2 medium droplet until chromosome spreads are performed (Fig. 4d). 16. Place 5-10 μl of 1% PFA, containing 0.15% Triton X-100 and 3 mM DTT, into each well of a 12-well slide (see Note 23). 17. Transfer a single ZP-free oocyte to the first well, carrying over as little M2 medium as possible.

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18. Wash away any residual PFA from the pipette tip in the PBS droplet (in the ZP removal dish), and repeat step 17 until all oocytes are transferred. 19. Allow the well slides to air dry at room temperature for overnight. 20. The chromosome spreads can either be processed directly for immunolabeling and DAPI staining, or stored at 4  C for at least 1 week (see Note 24). 3.5 Immunostaining and Imaging

1. Place slides in a humid chamber and pipette 1 ml TBST onto each slide. Leave for 5 min at room temperature (see Note 25). 2. Drain off the TBST and repeat step 1. 3. Drain off the TBST and wick off the excess by placing the edge of the slide on a paper towel. Add 1 ml blocking buffer. Incubate for 30 min at room temperature in humid chamber. 4. Drain off the blocking buffer and repeat the blocking buffer wash in step 3. 5. Dilute primary antibodies in ADB and add 100 μl per slide. Spread the antibody/ADB under a paraffin cover slip to cover the entire slide (or half slide for fetal oocyte spreads). 6. Incubate the slides overnight at room temperature in a humid chamber. 7. Remove the coverslip and add 1 ml TBST to each slide to rinse off the primary antibody. 8. Rinse slides three times with 1 ml TBST for 5 min each. Make sure that the slides never dry out during this process. 9. Dilute secondary antibodies in ADB. Add 100 μl of diluted secondary antibody per slide and spread with a paraffin cover slip. 10. Incubate for 60 min at 37  C in a dark humid chamber. 11. Remove the coverslip and add 1 ml TBST to each slide to rinse off the secondary antibody. 12. Rinse slides three times with 1 ml TBST for 5 min each in dark humid chamber. 13. ∗Dilute DAPI into TBST for a final concentration of 5 μg/ml and apply 200 μl per slide. Incubate for 10 min at room temperature in the dark (∗this step is only used for oocyte metaphase spreads). 14. Add 30 μl of ProLong antifade solution to each slide, and carefully mount a coverslip by placing one edge down first and slowly lowering the other edge. 15. Seal slides with clear nail polish and image via epifluorescence microscopy (Fig. 5).

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Fig. 5 Chromosome spreads. Representative images of pachytene and metaphase-I chromosome spreads from spermatocytes and oocytes. In pachytene nuclei, chromosome axes (SYCP3) are labeled in purple and Mlh1 is in green. In metaphase-I nuclei, positions of chiasmata are indicated with yellow arrows; yellow arrowheads highlight unconnected univalent chromosomes. In spermatocytes, white arrows indicate the sex chromosomes. In oocytes metaphase-I nuclei, centromeres (CREST) are labelled in red. Corner insets frame chromosomes that were from the same nucleus but located in different fields of view. Scale bars, 10 μm

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Notes 1. In vitro culture of mouse GV-stage oocytes requires limited laboratory equipment. An accurate temperature-controlled metal heat block is sufficient, and a CO2 incubator is not necessarily required. 2. Preferably use premade PBS from Invitrogen. However, if preparing your own PBS, carefully adjust the pH of 1 PBS to 7.4 prior to use and filter sterilize. 3. The average number of crossovers observed in juveniles (i.e., the first wave of spermatogenesis) and adults are reported to vary [23]; therefore, age-matched controls should be used for crossover quantification. 4. Do not disturb the loose pellet of tubule fragments at the base of the tube. Carefully transfer the cell suspension to a new tube and avoid transferring any tubule fragments by leaving the last ~500 μl behind. 5. Doubling the cell filter set (70 + 40 μm) significantly improves the removal of sperm from the cell suspension and improves spread quality and homogeneity. 6. The size of the pellet is crucial. If the pellet is too large, resuspend and remove some cell suspension. Conversely, if the pellet appears too small, resuspend and add some more of the cell suspension from see subheading 3.1, step 15. 7. Avoid spreading PFA all the way to the edges of the slide. This helps optimize the quality of chromosome spreads around the middle of the slide. 8. Drain off excess PFA by gradually tilting the slide and touching one corner to a wad of paper towels. This helps to accelerate drying of the slide during the next half an hour when the lid of the humid chamber is open. 9. The small orifice of a P20 pipette tip may mechanically damage the hypotonic-treated oocytes. 10. The volume of sucrose cell suspension and 1% PFA fixative solution can be varied depending on the area of the slide you want to cover and how many slides you need to prepare. However, keep the ratio between the volume of sucrose suspension, 1% PFA fixative solution, and the area on the slide constant; for example, 20 μl of sucrose suspension can be added to 100 μl of 1% PFA fixative solution to cover an entire slide. In this procedure, a pair of ovaries should yield ~1000 SYCP3-positive oocyte spreads, i.e., oocytes in prophase-I. 11. Longer incubations of cell suspensions in hypotonic buffer may result in overspreading of metaphase chromosomes.

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Fig. 6 Apparatus for metaphase spermatocyte chromosome spreads. Photo of setup used for dropping cell suspension onto glass slides to effect metaphase-I chromosome spreading

12. Cell pellets should be completely suspended to achieve homogenous and efficient fixation in see subheading 3.1, step 16. 13. Slow and homogenous fixation via dropwise addition of fixative solution with gentle vortexing is crucial to obtain well-spread metaphase chromosomes that enable quantification of chiasmata. 14. The distance between the glass pipette and the slide, and the humidity in the bench area are also critical for obtaining wellspread metaphase chromosome spreads. See apparatus in Fig. 6. 15. Each new batch of mineral oil should always be tested before general use because some batches can be toxic to oocytes. 16. Pregnant mare serum gonadotropin/equine chorionic gonadotropin (PMSG/eCG) can be used to stimulate follicle growth and, thereby, obtain larger numbers of GV stage oocytes. Intraperitoneal injection of 5–7.5 IU PMSG/eCG should be performed 44–48 h prior to oocyte collection. 17. New glass pipettes should be used for each collection day to avoid losing or contaminating oocytes. Before sucking oocytes into a pipette, always take up a small amount of fresh solution (2–5 μl) from the drop that you will be transferring into. This will minimize the risk of producing air bubbles that cause

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oocytes to be lost when you blow them out. Depending on your mouth pipetting skills, up to 20 cumulus cell-oocyte complexes can be picked up for each transfer. During oocyte collection, move the collection dish very gently and only as needed to ensure that collected oocytes do not get shuffled. 18. An optimally pulled glass pipette is critical for this procedure. The pipette opening should be similar to or slightly larger than the oocytes. A smaller pipette opening will damage the oocytes, and a larger opening is less effective for removing cumulus cells. 19. While transferring oocytes between droplets, always take up the oocytes with as little medium as possible. 20. In C57BL/6J mice, the majority of oocytes complete GV breakdown (GVDB) within 2 h and anaphase-I occurs around 8–10 h later. However, in some strains, such as CAST, anaphase-I can occur as early as 6 h post GVBD. 21. A brief wash in the first acid droplet is important since any residual M2 medium carried over to the acid droplet will adversely affect the rate of ZP removal. 22. Dependent on the precise pH of the acid droplet and the status of individual oocytes, ZP disappears in 10–30 s. The oocytes should be rapidly transferred into M2 medium as soon as the ZP dissolves. Overincubation in acid will damage the oocytes. However, the ZP has to be completely removed for optimal fluorescence staining. 23. A larger volume of PFA can be used if the chromosome spreads are not open enough; alternatively, reduce the volume if chromosomes are over spread. 24. Immunolabeling of fresh spreads is recommended. 25. Using a pipette tip to spread TBST over the entire 12-well slide during the first wash makes it easier for solutions to spread evenly over the slide at subsequent steps.

Acknowledgments M.I. was supported by a Japan Society for the Promotion of Science postdoctoral fellowship for research abroad. S.S. was supported by an A.P. Giannini Foundation postdoctoral fellowship. N.H. is an investigator of the Howard Hughes Medical Institute.

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first meiotic prophase in mammals. Endocr Rev 27(4):398–426 21. Chambon JP, Hached K, Wassmann K (2013) Chromosome spreads with centromere staining in mouse oocytes. Methods Mol Biol 957:203–212. https://doi.org/10.1007/ 978-1-62703-191-2_14 22. MacLennan M, Crichton JH, Playfoot CJ, Adams IR (2015) Oocyte development, meiosis and aneuploidy. Semin Cell Dev Biol

45:68–76. https://doi.org/10.1016/j. semcdb.2015.10.005 23. Zelazowski MJ, Sandoval M, Paniker L, Hamilton HM, Han J, Gribbell MA, Kang R, Cole F (2017) Age-dependent alterations in meiotic recombination cause chromosome segregation errors in spermatocytes. Cell 171(3):601–614. e613. https://doi.org/10.1016/j.cell.2017. 08.042

Chapter 20 Detection of DSBs in C. elegans Meiosis Tatiana Garcı´a-Muse Abstract Meiosis is a specialized reductional cell division responsible for the formation of gametes and the generation of genetic diversity. A fundamental feature of the meiotic process is the initiation of homologous recombination (HR) by the programmed induction of DNA double-strand breaks (DSBs). Caenorhabditis elegans is a powerful experimental organism, which is used to study meiotic processes mainly due to the germline that allows for visualization of sequential stages of meiosis. C. elegans meiosis-programed DSBs are resolved through HR; hence, the germline provides a suitable model to study DSB repair. Classically direct procedures to detect and study intermediate steps in DSB repair by HR in the nematode rely on germline immunofluorescence against the strand exchange protein RAD-51. Key words Double-strand breaks, C. elegans, RAD-51, Immunofluorescence, Homologous recombination

1

Introduction Meiosis consists of two cell divisions, called meiosis I and meiosis II, without replication in the middle that produces haploid gametes from diploid germline cells; each division goes through a prophase, metaphase, anaphase, telophase, and cytokinesis stages (named I or II respectively). During the two meiotic divisions, replicated chromosomes must segregate properly ensuring that each haploid cell has the correct number and type of chromosomes. Central to accurate chromosome segregation in meiosis is the establishment of the recombinationcrossovers resulting from meiotic DSB repair through the homolog by HR. In C. elegans, meiotic DSB repair by HR can be monitored by genetic approaches based on measuring crossover frequency using phenotypic markers or by direct cytological assays of the germline [1]. Immunofluorescence is a cytological assay used for light microscopy with a fluorescence microscope and can be utilized on tissue sections, cultured cells, or individual cells. This technique uses the specificity of antibodies to their antigen to analyze the presence and distribution of target proteins and is widely

Andre´s Aguilera and Aura Carreira (eds.), Homologous Recombination: Methods and Protocols, Methods in Molecular Biology, vol. 2153, https://doi.org/10.1007/978-1-0716-0644-5_20, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 C. elegans germline scheme. Germline nuclei in the distal tip of the gonad arm divide mitotically and thereafter undergo meiosis. As the nuclei move away from the distal end of the gonad they progress through prophase I. The regions are: 1- mitosis; 2- transition zone (leptotene and zygotene); 3-, 4-, and 5correspond to early, middle, and late pachytene, respectively; 6- diplotene and diakinesis

used in C. elegans [2, 3]. The nematode contains two germlines and each one is organized in a spatiotemporal manner starting from mitotic nuclei at the distal tip that will enter meiosis and undergo all the prophase I stages; hence it can be described as a meiosis time course [4]. As nuclei move proximally, they enter the transition zone that corresponds to the first stages of meiosis (leptotene and zygotene), followed by pachytene, diplotene, and reaching diakines (Fig. 1). During prophase I programmed DSBs are generated by the meiosis-specific endonuclease SPO-11 and subsequently repaired via HR [5, 6]. Another option is the generation of exogenous DSBs exposing the worms to ionizing radiation, which results in radiation-induced DSBs [7, 8]. One of the key proteins needed for homologous recombination repair is the DNA strand exchange Rad51, a homolog of the bacterial RecA protein [9]. During C. elegans meiosis RAD-51 becomes localized to discrete foci in nuclei of the first meiosis phases and it is most abundant in nuclei at early pachytene, and then those foci progressively disappear from DNA and no foci are observed in late pachytene [10]. Analysis of the assembly and disassembly dynamics of the strand exchange protein RAD-51 onto meiotic DSBs of germline nuclei in the gonad provides information regarding the process of DSB repair [7, 10]. This is a useful tool for the reason that unprogrammed DNA double-strand breaks (DSBs) are one of the most cytotoxic DNA lesions and must be repaired to preserve chromosomal integrity. Nevertheless, the identification of novel proteins involved in homologous recombination, such as COSA-1, a cyclin-related protein required to convert meiotic DSBs into cross-overs, suitable for the generation of transgenes containing fusion to fluorescence epitopes have allowed us to monitor HR in vivo [11].

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Materials

2.1 C. elegans Nematodes

1. 3 cm plates. 2. MYOB growth media (for 1 L: 4.6 g bactotryptone, 8 mg cholesterol, 2.0 g NaCl, 0.55 g Tris–HCl, 0.24 g TrisOH, 20 g agar). 3. HB101 or OP50 bacteria. 4. Worm pick made by a 1 cm of platinum wire 0.25 mm (Sigma 349402-250 mg) stick on a short Pasteur pipette tip by heat. 5. Stereoscope.

2.2 Poly-Lysine Slide Preparation

1. Microscope Slide. 2. Poly-L-lysine (Sigma P8920-100 ml). 3. Ethanol.

2.3 Tissue Extraction and Preparation

1. Scalpel or 23 gauge needle. 2. PBS tablets (Gibco). 3. Paraformaldehyde powder (Sigma P6148). 4. BSA (sigma A2153-10G). 5. Triton Tx-100 (Sigma T9284-100 ml). 6. TBS10 pH 8.0 (for 1 L: 80 g NaCl + 2 g KCl + 30 g Tris– HCl). 7. TBS-B (TBS1X, 0.05% BSA). Use under one-week old. 8. TBS-BTx (TBS-B, 0.001% Triton Tx-100). Use freshly made. 9. Pap-Pen 5 mm (Sigma Z377821). 10. Levamisole (Sigma 46944-250 mg).

2.4 Antibody Incubation

1. Anti-RAD-51 antibody (Novus Biologicals NB100-148). 2. Alexa Fluor-568 goat anti-rabbit antibody (Invitrogen). 3. 24 mm  50 mm coverslips. 4. DAPI solution (1 μg/ml in PBS). 5. Vectashield (H1000 Vector Laboratories). 6. Nail polish. 7. Fluorescence microscope.

3

Methods

3.1 Lab-Made Poly-Lysine Slide Preparation

1. Wash slides with a tissue soaked in 96 ethanol, and dry in an oven at 60  C for 5–10 min.

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2. Shake the cleaned slides in a poly-L-lysine solution for 5 min at RT. 3. Remove excess poly-L-lysine and dry in an oven at 60  C for 1 h. 4. Store at 4  C. 3.2 Tissue Extraction and Preparation

1. 30–50 L4-stage nematodes are transferred to MYOB plates seeded with bacteria (HB101 or OP50) and maintained at 20  C (see Note 1). 2. After 24 h, nematodes reach the right stage to be analyzed. To induce exogenous DSB, the worms can be irradiated (see Note 2). 3. Box an area on the slide using a Pap-Pen to contain solutions. One for each strain or condition. A maximum of eight squares per slide can be drawn (around 1  1 cm). 4. Place a drop of about 25 μl of PBS 1 just beside the Pap-Pen squares. One for each strain or condition. 5. Pick the worms into the drop of PBS (see Note 3). 6. Transfer the nematodes into a drop of about 25 μl PBS + levamisole (0.05 mM) inside the Pap-Pen squares. 7. To dissect out germline cut heads (below the pharynx) and/or tails (after the germline turn) off each animal using a scalpel or needle (see Fig. 2) (see Note 4). 8. Replace the PBS + levamizole with 4% paraformaldehyde (4% PFA) onto the cut animals (see Notes 5 and 6). 9. Incubate for 1 h at RT in a humidified chamber. 10. Replace the PFA with TBS-BTx solution (TBS + 0.5% BSA + 0.1% triton) for 5 min at RT (see Note 7). 11. Remove the TBS-BTx and wash two times with TBS-B solution (TBS + 0.5% BSA) for at least 10 min each time (see Note 8). 12. Incubate the samples in TBS-B for at least 30 min at RT.

Fig. 2 C. elegans scheme. The nematode presents a vermiform body shape with head and tail at opposite ends. Because C. elegans is transparent both gonads and the intestine are easily visualized. Dashed lines indicate the regions to cut the animal for germline extrusion

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1. Remove the TBS-B and add the anti-RAD-51 antibody (1:10000 in TBS-B). Incubate overnight at 4  C (preferred) or for 2 h at RT, in a humid chamber (see Note 9). 2. Rinse the samples with a quick wash with TBS-B solution (see Note 10). 3. Wash three to four times with TBS-B for at least 10 min for each wash. 4. Remove the TBS-B and incubate with Alexa Fluor goat antirabbit secondary antibody (1:5000 in TBS-B solution) for 2 h in the dark at RT. Store in the dark from now on. 5. Rinse the samples by a quick wash with TBS-B solution. 6. Wash four times in TBS-B solution for at least 15 min for each wash. 7. Remove the TBS-B and add PBS + DAPI (1 μg/μl) for at least 10 min (see Note 11). 8. Remove the PBS + DAPI and add 5–10 μl of mounting medium in each Pad-Pen square. 9. Cover the samples with a long coverslip. Seal the edges with nail polish. Place the slides flat and let dry for at least 1 h at RT and leave overnight at 4  C before storing at 20  C.

3.4 RAD-51 Quantification

1. Visualization of RAD-51 foci on germ cells (see Fig. 3) requires a fluorescence microscope (600 to 1000 magnification) that allows the acquisition in the Z axe. 2. To quantify DSB repair dynamics the number and distribution of RAD-51 foci are counted. 10 nuclei of each region (see Fig. 1) of at least 10 worms are analyzed from three independent experiments. 3. The number of nuclei with RAD-51 foci as well as the number of foci per nuclei are represented for each germline region (see a wild-type (N2) profile in Fig. 4). 4. Alternatively a time course of DSB repair can be performed from irradiated worm. RAD-51 foci form after a medium IR dose (75Gy) in less than 1 h and are repaired after 24 h.

Fig. 3 RAD-51 foci. Representative images of the meiotic region from N2(wt) fixed germlines immunostained with anti-RAD-51 antibody and counterstained with DAPI in normal conditions

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Fig. 4 RAD-51 distribution along the germline. Quantification of the recombination marker RAD-51 foci in the N2 (WT) in normal conditions. At least 15 gonads were analyzed, and 10 nuclei were scored in each zone (mitotic region (1), transition zone (2), early-mid-late pachytene regions (3-45), and diplotene-diakinesis regions (6)) for at least three independent experiments

4

Notes 1. A white spot in the center of the body characterizes L4 stage worms. 2. In typical survival experiments 30Gy represents low dose, 75Gy a medium dose, and 120Gy high dose. This correlates with the levels of DSB formation. 3. This step is to get rid of as much bacteria from the worms as possible before dissecting. If any bacteria get onto the polylysine slide, then the worms will not stick well. 4. The levamizole paralyzes the nematode movement and aids the dissection. However, too long in the presence of levimazole can kill the worms and then gonads do not extrude. Also some mutants are very sensitive; in that case use only PBS solution during dissection. 5. Is detrimental for the gonads to remain in the solution before fixation; therefore, beginners should start with a low number of worms. 6. Removal and addition of solutions needs to be gentle since the germlines can detach from the slide. 7. Formaldehyde is a sensitizing agent and a cancer hazard. Wear gloves and lab coat and always in a chemical fume hood. 8. PFA is prepared with H2O Milli-q. For 25 ml. preheat 22 ml H2O Milli-q at 60  C and add 1 g PFA while stirring in the chemical fume hood. If it does not dissolve add 2 μl NaOH (10 N). Repeat if necessary. Filter with a 0.45 μm filter and store at 20  C for long term. Once thawed, do not reuse.

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9. This step is for permeabilization, but more than 5 min with Triton-Tx is detrimental. 10. Store TBS-B at 4  C. 11. This step can be substituted using mounting medium with DAPI (1:500), which can be stored at 4  C in the dark.

Acknowledgments This work was supported by a Spanish Ministry of Science, Innovation and Universities grant (PGC2018-101099-B-I00). T.G-M. was the holder of postdoctoral grants from the CSIC JAE-Doc and the Junta de Andalucı´a Excellence Program (CVI-4567). References 1. Hillers KJ, Villeneuve AM (2009) Analysis of meiotic recombination in Caenorhabditis elegans. Methods Mol Biol 557:77–97 2. Hendricks M (2015) Observing and quantifying fluorescent reporters. Methods Mol Biol 1327:75–85 3. Phillips CM, McDonald KL, Dernburg AF (2009) Cytological analysis of meiosis in Caenorhabditis elegans. Methods Mol Biol 558:171–195 4. Lui DY, Colaia´covo MP (2013) Meiotic development in Caenorhabditis elegans. Adv Exp Med Biol 757:133–170 5. Dernburg AF, McDonald K, Moulder G, Barstead R, Dresser M, Villeneuve AM (1998) Meiotic recombination in C. elegans initiates by a conserved mechanism and is dispensable for homologous chromosome synapsis. Cell 94(3):387–398 6. Garcia-Muse T, Boulton SJ (2007) Meiotic recombination in Caenorhabditis elegans. Chromosom Res 15(5):607–621

7. Garcia-Muse T, Boulton SJ (2005) Distinct modes of ATR activation after replication stress and DNA double-strand breaks in Caenorhabditis elegans. EMBO J 24(24):4345–4355 8. Craig AL, Moser SC, Bailly AP, Gartner A (2012) Methods for studying the DNA damage response in the Caenorhabdatis elegans germ line. Methods Cell Biol 107:321–352 9. Ogawa T, Yu X, Shinohara A, Egelman EH (1993) Similarity of the yeast RAD51 filament to the bacterial RecA filament. Science 259:1896–1899 10. Alpi A, Pasierbek P, Gartner A, Loidl J (2003) Genetic and cytological characterization of the recombination protein RAD-51 in Caenorhabditis elegans. Chromosoma 112(1):6–16 11. Yokoo R, Zawadzki KA, Nabeshima K, Drake M, Arur S, Villeneuve AM (2012) COSA-1 reveals robust homeostasis and separable licensing and reinforcement steps governing meiotic crossovers. Cell 149(1):75–87

Chapter 21 Methods to Map Meiotic Recombination Proteins in Saccharomyces cerevisiae Aurore Sanchez and Vale´rie Borde Abstract Meiotic recombination is triggered by programmed DNA double-strand breaks (DSBs), catalyzed by the type II topoisomerase-like Spo11 protein. Meiotic DSBs are repaired by homologous recombination, which produces either crossovers or noncrossovers, this decision being linked to the binding of proteins specific of each pathway. Mapping the binding of these proteins along chromosomes in wild type or mutant yeast background is extremely useful to understand how and at which step the decision to repair a DSB with a crossover is taken. It is now possible to obtain highly synchronous yeast meiotic populations, which, combined with appropriate negative controls, enable to detect by chromatin immunoprecipitation followed by sequencing (ChIP-Seq) the transient binding of diverse recombination proteins with high sensitivity and resolution. Key words Chromatin immunoprecipitation, High-throughput sequencing, Genome-wide maps, DNA sonication, Meiotic proteins, Recombination intermediate mapping

1

Introduction Meiotic recombination in budding yeast is initiated via the programmed formation of DNA double-strand breaks in gene promoters, and generally, in nucleosome-depleted regions (NDR) [1]. Among several hundreds of DSBs formed per cell, only a subset of these will be repaired with a crossover, the other being repaired through noncrossover events [2]. The choice of repair pathway is controlled by many factors. At the single-cell level, it is governed by the phenomenom of interference, where the presence of a crossover-designated event lowers the probability of a nearby crossover occurring on the same chromosome [3]. In large cell populations, some studies have suggested that this decision could be also governed by specific chromosome features, such as the proximity to a centromere or a telomere [4–6]. One difficulty to map proteins binding to recombination events in meiosis is that this event is by nature transient, and occurs at a given site in only a small fraction of

Andre´s Aguilera and Aura Carreira (eds.), Homologous Recombination: Methods and Protocols, Methods in Molecular Biology, vol. 2153, https://doi.org/10.1007/978-1-0716-0644-5_21, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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the cells, at sites called hotspots. This makes it challenging to detect meiotic recombination proteins, especially those predicted to bind for a short time to recombination sites, in contrast with proteins such as transcription factors or chromosome structure proteins that have a longer residency on chromatin. In addition, nucleosomedepleted regions, where meiotic recombination occurs, due to their higher chromatin accessibility, are sometimes retrieved in different amounts than other regions upon purification, and therefore particular attention should be paid to using relevant controls, to reveal only the studied protein-binding sites, without any artifact due to the nature of the region being analyzed. Indeed, it is very important to use as a control a sample that is processed in an identical way as the real sample, ideally from cell lysates sonicated the same way and immunoprecipitated using the same antibody as the experimental sample. This is particularly important for proteins giving a very low signal over background. For instance, input DNA would not be suitable here as a control. To circumvent these difficulties, we describe here a method to map meiotic recombination proteins by ChIP-Seq, using highly synchronized cells, and using as a negative control a recombination-deficient strain, which allows to reveal only recombination-specific binding sites of a given protein. The combined use of highly synchronized cells and of a recombination deficient strain (such as spo11Δ) as the negative control allowed us to map recombination proteins that are difficult to detect by chromatin immunoprecipitation, such as the Pif1 helicase and the Mlh3 crossover protein. To obtain a highly synchronized meiosis, we used the approach developed by Chia and van Werven [7], where the expression of the master regulator of meiosis, IME1, is put under the control of an inducible promoter, which allows all cells of the population to “prepare” for meiosis and reach the same stage upon meiosis induction. The ChIP-Seq technique presented here was previously used in our lab to map “ZMM” meiotic recombination proteins [8].

2

Materials

2.1 Media for Yeast Culture

1. YPD: 1% Bacto yeast extract, 2% Bacto peptone, 2% dextrose. Autoclave (20 min at 120
C) (see Note 1). 2. “Semi-YPD”: 1% Bacto yeast extract, 2% Bacto peptone, 1% dextrose. Autoclave (20 min at 120
C) (see Note 1). 3. 1% potassium acetate (KAc). Autoclave. 4. 5% raffinose. Filter-sterilize. 5. 1% poly propylene glycol 2000 (PPG) (Sigma).

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6. Sporulation medium: add raffinose to 0.02% and PPG to 0.001% to autoclaved 1% KAc. 7. CuSO4: 500 mM. Filter-sterilize. 2.2 Cell Crosslinking and Lysis

1. FastPrep-24™ 5G instrument (MPbio). 2. Thin tips (200 μL) (Dutscher). 3. 2-mL FastPrep screw cap tubes (MPbio). 4. Zirconia/silicia beads BioSpec products (Fisher Scientific). 5. 16% Paraformaldehyde Sciences).

solution

(Electron

Microscopy

6. Lysis buffer: 50 mM Hepes-KOH pH 7.5, 140 mM NaCl, 5 mM EDTA, 1% Triton X-100, 0.1% sodium deoxycholate. Prepare 250 mL. Filter-sterilize. 7. 1 mg/mL aprotinin (Euromedex). 1 mg aprotinin is dissolved in 1 mL milliQ water. 8. cOmplete™, Mini EDTA-free protease inhibitor cocktail tablets (Roche). 9. 100 mM phenylmethylsulfonyl fluoride (PMSF) solution in isopropanol. 2.3 Immunoprecipitation

1. Bioruptor® Pico sonication device (Diagenode). 2. 15 mL Bioruptor® Pico tubes and sonication beads (Diagenode). 3. DynaMag™-2 Magnet rack (Thermo Fisher Scientific). 4. Rotating wheel for use at 4
C. 5. 1.5 mL Eppendorf DNA LoBind tubes. 6. 100 mg/mL BSA. 7. 1 mg/mL c-Myc mouse monoclonal antibody (9E10). 8. Magnetic Dynabeads Pan mouse IgG (Thermo Fisher Scientific) (see Note 2). 9. Wash and bind buffer (100 mL): 95.5 mM NaH2PO4, 4.5 mM NaH2PO4, 0.01% Tween 20 pH 8.2. Filter-sterilize. 10. Wash buffer 2: 50 mM Hepes-KOH pH 7.5, 500 mM NaCl, 5 mM EDTA, 1% Triton X-100, 0.1% sodium deoxycholate. Prepare 250 mL. Filter-sterilize. 11. Wash buffer 3: 10 mM Tris–HCl pH 8.0, 250 mM LiCl, 5 mM EDTA, 0.5% sodium deoxycholate, 0.5% NP-40. Prepare 100 mL. Filter-sterilize. 12. TE buffer: 10 mM Tris pH 7.5, 1 mM EDTA. Autoclave. 13. IPure kit v2 (Diagenode). 14. Isopropanol 100%.

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2.4 Sample Quality and Quantity Control

1. Qubit 4 fluorometer (ThermoFisher Scientific). 2. Qubit™ dsDNA HS Assay Kit (ThermoFisher Scientific). 3. Bioanalyzer instrument (Agilent). 4. Bioanalyzer High Sensitivity DNA Analysis (Agilent).

2.5 DNA Library Preparation and Sequencing

1. Illumina® TruSeq®Chip Library Preparation Kit. 2. Chloroform/isoamyl alcohol (v/v, 24: 1). 3. 5PRIME Phase Lock Gel-Heavy tubes, 2 mL (Quantabio). 4. 70% and 100% ethanol. 5. 35 mg/mL glycogen (Support—MP Biomedicals). 6. Illumina HiSeq 2500 instrument.

3

Methods

3.1 Highly Synchronous Meiotic Cultures

This section is based on [7] with minor modifications. In this system, cells expressing the IME1 gene under the pCUP1 inducible promoter are grown in rich medium containing twice less glucose than the regular YPD (“semi-YPD”), transferred in sporulation medium, to allow all cells to accumulate in the G1 phase (and perform the diauxic shift), and IME1 is induced by the addition of copper after 2 h in sporulation medium. In these settings, premeiotic DNA replication should be over at 4 h [7]. When compared to the “classical” sporulation protocol following a pregrowth in SPS medium [9], we observed that DSB and crossover recombination occur in a shorter period of time and DSBs peak at a higher level, due to better cell population synchronization (Fig. 1). In these conditions, DSBs appear at 4 h and crossovers start appearing at 5 h 30; therefore, to map an “early” recombination protein (such as Pif1) we use the 5 h time-point; for a “middle” recombination protein such as a ZMM protein, we use the 5 h 15 time-point; and for a “late” recombination protein such as Mlh3, we use the 5 h 30 time-point. 1. Day 4: Streak out the diploid on a YPD plate to generate single colonies and incubate at 30
C for two days. 2. Day 2: At 6 pm, inoculate a single colony in 5 mL YPD and incubate overnight (ON), at 30
C under agitation (250 rpm). 3. Day 1: At 11.15 am, dilute saturated YPD preculture 100-fold into 20 mL fresh YPD. Then incubate at 30
C under agitation (250 rpm). 4. At 4.45 pm, cells will be in exponential phase (OD < 2) with an OD600 ¼ 1.7. Inoculate at OD600 ¼ 0.05 into 60 mL “semiYPD.” Incubate ON at 30
C under agitation (250 rpm). Prewarm the 500 mL autoclaved 1% KAc bottle at 30
C ON.

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Fig. 1 Comparison of the timing of recombination events in “classical” SPS versus pCUP1-IME1-synchronized meiotic time-courses. (a) Southern blot monitoring the formation and repair of DNA DSBs at the HIS4LEU2 hotspot, in a strain containing pCUP1-IME1 and induced to undergo meiosis by the addition of Cu++ at 2 h after transfer to sporulation medium. P1 and P2: parental DNA fragments. CO1 and CO2: bands corresponding to the crossover repair of HIS4LEU2 DSB. DSB: DSB fragments from the DSB formed at the HIS4LEU2 hotspot [10]. (b) Quantification of DSB formation (%DSB) and repair as a crossover (%CO) from the experiment shown in (a). (c) Quantification of DSB formation (%DSB) and repair as a crossover (%CO) from a meiotic time-course using the “classical” SPS meiotic time-course. Data are from [11]

5. Day 0: At around 9–9.30 am, cells should be at OD600 ¼ 12–13. Centrifuge for 5 min at 5000  g and discard the supernatant (see Note 3). 6. Wash the pellet with one volume of prewarmed 1% KAc. Centrifuge 5 min at 5000  g and discard the supernatant. Make the sporulation medium. 7. Resuspend in sporulation medium to an OD600 ¼ 2.5. The 0 h time-point starts now. Incubate at 30
C under agitation (250 rpm). 8. After 2 h, add CuSO4 to a 50 μM final concentration to induce the expression of IME1 (20 μL from a 500 mM stock solution for 200 mL sporulation medium).

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9. After 8 h in sporulation medium, check meiotic divisions by DAPI staining. A typical percentage of 95–100% binucleate plus tetranucleate cells is expected. 3.2

Cross-Linking

The remaining protocol is written for one sample of one strain, where the protein to be immunoprecipitated is tagged with a myc epitope. It is strongly recommended to process the test and control strains in parallels. We usually perform test and control experiments each in duplicates. 1. Cool PBS at 4
C. 2. At the desired time point (for instance 5 h 30 for a late recombination protein), take 1.109 cells (50 mL of the sporulation culture at DO600 ¼ 2.5) and divide them into two 50 mL conical tubes (see Notes 4 and 5). 3. Add 1.56 mL of 16% paraformaldehyde solution (final 1%) to each 25 mL culture and incubate at room temperature (RT) for 15 min on a rotating wheel (40 rpm). 4. Add 1.25 mL 2.5 M glycine (final 0.125 M) to quench the cross-linking reaction. 5. Incubate at RT for 5 min on a rotating wheel (40 rpm). 6. Centrifuge for 5 min at 5000  g and discard the supernatant under a fume hood in a suitable container for waste containing formaldehyde. 7. Resuspend the pellet in 25 mL of cold PBS 1. Centrifuge 5 min at 5000  g and discard the supernatant. 8. Repeat step 7. 9. Resuspend each pellet in 2.5 mL PBS 1. Pool the pellets (5 mL total) and distribute into five FastPrep tubes. 10. Centrifuge 1 min max speed at 4
C. 11. Carefully discard supernatants; no liquid should remain in the tubes. 12. Freeze pellets in liquid nitrogen and store at 80
C (or process immediately) (see Note 6).

3.3 Preincubation of the Beads with Antibody

1. Resuspend Pan mouse IgG magnetic beads by vortexing >30 s. 2. Transfer 250 μL of magnetic beads to a 2 mL Eppendorf tube. 3. Place the tube on the magnet to separate the beads from the solution and remove the supernatant (use of the magnetic rack, see Note 7). 4. Wash two times with 1 mL wash and bind buffer. Use the magnet between each wash to remove the supernatant.

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5. Resuspend the beads in 1 mL wash and bind buffer. Add 50 μL of 100 mg/mL BSA and incubate on a rotating wheel at 4
C for 4 h (see Note 8). 6. Remove the supernatant with the magnet. 7. Wash two times with 1 mL wash and bind buffer. Use the magnet between each wash to remove the supernatant. 8. Resuspend the beads in 1 mL of wash and bind buffer. 9. Add 40 μl 200 μg/mL c-Myc monoclonal antibody (20 μg total) and incubate on a wheel at 4
C for 4 h. 3.4

Cell Lysis

1. Prepare 10 mL lysis buffer supplemented with protease inhibitors by adding 1 mM PMSF, 50 μg/mL aprotinin and 1 cOmplete™, Mini, EDTA-free protease inhibitor cocktail and keep on ice (see Note 9). 2. Remove the five FastPrep tubes from the freezer from Subheading 3.2, step 12 and thaw cells on ice. 3. Resuspend each pellet in 500 μL of cold lysis buffer supplemented (Prepared in 1). 4. Add an approximately equal amount of Zirconia/silicia beads (a full 500 μL Eppendorf tube). 5. Process cell lysis in a FastPrep-24™ 5G instrument with the following program to do three times: Intensity 6, 30 s ON and 3 min pause on ice between each program repeat. 6. Collect the five lysates by pipetting with thin tips to avoid catching beads and pool them to a 15 mL conical tube (see Note 10). 7. Adjust to a final volume of 2.5 mL with lysis buffer supplemented with protease inhibitors.

3.5 Sonication (See Note 11)

1. Add 900–1000 mg Bioruptor™ beads into a 15 mL Bioruptor™ Tubes graduation (equivalent to the fourth line of the 15 ml Bioruptor™ Tubes graduation scale). Wash beads by vortexing in three volumes of PBS 1. Remove all PBS using thin tips. 2. Add 2.5 mL lysate (from Subheading 3.4, step 7). 3. Sonicate to an average fragment size of approximatively 0.1–0.5 kb with the precooled Bioruptor™ Pico instrument at 4
C with the following program: 45 cycles of 30 s pulse, 30 sec pause (45 min total). 4. Collect the 2.5 mL sonicated lysate by pipetting with thin tips to avoid catching beads and split it into five 1.5 mL Eppendorf tubes.

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5. Clear the sonicated lysate by centrifugation for 5 min at 19,000  g and at 4
C. Transfer the supernatant into new tubes. 6. Keep a 10 μL aliquot of this supernatant as input control, freeze in liquid nitrogen, and store at 20
C (see Note 12). 3.6 Immunoprecipitation

1. Split 1 mL of magnetic beads plus antibody prepared in Subheading 3.3, step 9 into five 1.5 mL Eppendorf tubes (200 μl each). 2. Wash two times with 1 mL lysis buffer. Use the magnet between each wash to remove the supernatant. 3. Load the sonicated lysate from Subheading 3.5, step 5 on the beads and incubate ON at 4
C on a rotating wheel. 4. The next day, put tubes on the magnet and remove the supernatant. 5. Perform the following washes each for 1 min on a rotating wheel at room temperature. Use the magnet between each wash to remove the supernatant (see Note 13). (a) Wash two times with 1 mL lysis buffer. (b) Wash two times with 1 mL wash Solution 2. (c) Wash two times with 1 mL wash Solution 3. (d) Wash one time with 1 mL TE. 6. Completely remove the supernatant (see Note 13).

3.7 Elution and DNA Purification

The IPure Diagenode kit is used with the following modifications to elute DNA. (For more details see the manufacturer’s protocol): 1. Prewarm buffer A at 25
C during 30 min before use. 2. Prepare the elution buffer by mixing 692.4 μL Buffer A and 27.6 μL B. 3. Thaw the 10 μL input sample (Subheading 3.5, step 6) on ice and add 120 μL of elution buffer. 4. To each of the five tubes of IP (Subheading 3.6, step 6), add 120 μL of elution buffer and pool them in a new 1.5 mL Eppendorf DNA LoBind tube. From this stage use only Eppendorf DNA LoBind tubes. 5. Incubate samples and input for 4 h (or overnight) at 65
C on a thermomixer, with continuous shaking. 6. Put IP samples and input tubes on the magnet and after 1 min, transfer the supernatants to a new Eppendorf DNA LoBind tube. Keep the samples on ice. 7. Add 2 or 10 μL of carrier to the input or IP sample, respectively. Vortex briefly and do a short spin.

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8. Add 100 or 500 μl of 100% isopropanol to the input or IP sample, respectively. Vortex briefly and do a short spin. 9. Resuspend the magnetic beads from the kit and add 10 μL to each input and IP sample. 10. Incubate samples for 50 min at room temperature on a rotating wheel (40 rpm). 11. Prepare the wash buffer A by mixing 330 μL wash buffer 1 w/o isopropanol and 330 μL isopropanol 100%. 12. Perform a short spin. 13. Put IP samples and input tubes on the magnet and after 1 minute, discard the supernatants and add 100 μL wash buffer A to each tube. From this moment, beads stick very strongly to the tube’s walls. Be careful to leave the liquid and the beads at the bottom of the tube. Mix only by tapping the tubes six times. 14. Incubate 5 min at RT, and mix by tapping every 1 min. 15. Prepare the wash buffer B by mixing 330 μL wash buffer 2 w/o isopropanol and 330 μL isopropanol 100%. 16. Perform a short spin. 17. Put IP samples and input tubes on the magnet and after 1 min, discard the supernatants and add 100 μL wash buffer B to each tube. Mix only by tapping the tubes six times. 18. Incubate 5 min at RT, and mix by tapping every 1 min. 19. Perform a short spin. 20. Put IP samples and input tubes on the magnet and after 1 min, discard the supernatants and add 40 μL Buffer C to each tube. Mix only by tapping the tubes six times. 21. Incubate 30 min at RT, and mix by tapping every 5 min. 22. Perform a short spin. 23. Put IP samples and input tubes on the magnet and after 1 min, collect the supernatants and transfer them to a new Eppendorf DNA LoBind tube. 3.8 Sample Quality and Quantity Control

1. Quantify the sample concentration. For precise quantification of DNA quantity, we strongly recommend the use of the Qubit® instrument. Follow the manufacturer’s protocol. A typical yield of 0.8–2.5 ng/μL for input and IP DNA is expected. 2. Test the efficiency of DNA fragmentation by analyzing only the input DNA size using Bioanalyzer instrument. Follow the manufacturer’s protocol. A typical yield of 1–3 ng/μL DNA is expected. The size of DNA fragments should be between 150 and 300 bp. 3. Freeze samples with liquid nitrogen and store at 80
C.

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3.9 DNA Library Preparation and Sequencing

The purpose of this part is to add adapter sequences onto the ends of DNA fragments to generate libraries for paired-end 50 bp sequencing reads. Many new sequencing machines can be used for the sequencing step (Illumina, Ion Torrent...); here we used Illumina HiSeq 2500 instrument. Ten nanogram of both immunoprecipitated (ChIP-DNA) and nonimmunoprecipitated (input DNA) are used to prepare the DNA library for next generation sequencing. The Illumina® TruSeq®Chip Library Preparation Kit was used in the following steps (for more details see the manufacturer’s protocol): 1. Repair of 30 and 50 ends. 2. Library size selection. Library size is selected using a 1.8 ratio of AMPure XP beads/sample. 3. Addition of adenylate 30 ends. 4. Ligation of indexed paired-end adapters. 5. Library size selection. Library size is selected using two rounds of purification with 1 ratios of AMPure XP beads/sample. 6. Amplification via polymerase chain reaction (PCR) of adaptorligated DNA. 7. Library size selection. Library size is selected using one round of purification with 1 ratios of AMPure XP beads/sample. 8. Library validation. 9. Normalization and pooling libraries.

3.10 Read Normalization and Bioinformatic Analyses

Our procedure for ChIP-Seq data analysis has been published previously and we summarize here its main steps. Sequencing data are analyzed using custom Bash and R scripts [8]. 1. Reads are aligned to the sacCer2 version (SGD June 2008) of the S. cerevisiae S288C genome, using Bowtie, allowing for two mismatches. Reads that matched more than once in the genome or matched mitochondrial or ribosomal DNA were eliminated from further analysis. 2. Aligned paired-end reads are extended and then normalized as follows: for the control, the top and bottom outliers of the distribution are removed and the average read number per position (coverage) of the remaining reads is then set to 1. Similarly, top and bottom outliers of the distribution are removed from the average read number in the experimental samples, and average coverage for the remaining reads is calculated, and its ratio compared to the control is used to normalize all reads of the experimental sample. 3. Both scaled experimental and control samples are then converted to bigwig format, and the control bigwig file (usually from a spo11 mutant) is subtracted from the experimental file to

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generate a normalized, subtracted file that can be used to visualize the protein enrichment along chromosomes.

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Notes 1. Volvic™ water quality is constant and it is weakly mineralized so yeast cultures prepared in media with Volvic™ water are much more reproducible. We recommend to use Volvic™ water but distilled water could also be used. 2. PanMouse IgG beads are only used when the primary antibody is a mouse antibody. For antibodies generated from other species, we use protein G magnetic beads. 3. Example: for one culture at OD600 ¼ 12, take 41.7 mL culture. After washing in 1% KAc, resuspend the pellet in 200 mL sporulation medium to obtain OD600 ¼ 2.5. 4. The time point for sample collection must be determined for each new protein. To determine the precise moment when the protein enrichment is maximal, it is recommended to make a classical ChIP experiment followed by qPCR analysis as described in [8]. For this, take 10 mL of a pCUP1-IME1 synchronized culture every hour until 8 h after transfer to sporulation medium and follow the protocol described in [8]. 5. It is strongly recommended to take 50 mL twice. One sample is processed, and the second serves as a backup sample. 6. This is a freeze point. Flash-freezing of pellets must be done with liquid nitrogen and stored at 80
C until needed. Experiment can be stopped at this point. 7. Place the tube on the magnetic rack and wait 30 s until beads are separated from the solution. Invert the magnetic rack three times from top to bottom to recover all the beads contained in the tube cap. Wait again for 30 s and remove all supernatant by aspirating with a needle connected to a vacuum pump. 8. Magnetic bead incubation with BSA before IP permit to decrease nonspecific background. The other option is to do a preclearing of your sample. 9. PMSF must be stored at 20
C, and it is recommended to make it fresh. The addition of PMSF to the lysis buffer is done just at the time of the experiment. 10. The lysate can also be collected by making a hole in the bottom of the FastPrep tube with a sterile needle. After removing its screw-cap, quickly place the tube into a 15 mL Falcon tube and spin for 10 s at 1000  g to collect the lysate. 11. We recommend using the Bioruptor™ Pico, and the sonication conditions described are adapted to this instrument. If using

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another instrument, it is important to properly adapt the sonication conditions to obtain DNA fragmentation around 100–300 bp. Set the cooling device of the sonicator Bioruptor™ Pico at 4
C at least 30 min before use. 12. To decrease even more the nonspecific background in the IP experiment, you can do a second preclearing of your lysate. To do this, incubate your sample with magnetic beads only (without antibody) on a wheel for four hours at 4
C. After four hours, collect the lysate cleared twice and proceed to the next step. 13. We recommend to use a needle and aspiration to completely remove all traces of supernatant remaining in the lid and the bottom of the tube. References 1. Pan J, Sasaki M, Kniewel R, Murakami H, Blitzblau HG, Tischfield SE, Zhu X, Neale MJ, Jasin M, Socci ND, Hochwagen A, Keeney S (2011) A hierarchical combination of factors shapes the genome-wide topography of yeast meiotic recombination initiation. Cell 144 (5):719–731. https://doi.org/10.1016/j.cell. 2011.02.009 2. Hunter N (2015) Meiotic recombination: the essence of heredity. Cold Spring Harb Perspect Biol. https://doi.org/10.1101/cshperspect. a016618 3. Muller HJ (1916) The mechanism of crossing over. Am Nat 50:193–434 4. Chen SY, Tsubouchi T, Rockmill B, Sandler JS, Richards DR, Vader G, Hochwagen A, Roeder GS, Fung JC (2008) Global analysis of the meiotic crossover landscape. Dev Cell 15 (3):401–415. https://doi.org/10.1016/j. devcel.2008.07.006 5. Serrentino ME, Borde V (2012) The spatial regulation of meiotic recombination hotspots: are all DSB hotspots crossover hotspots? Exp Cell Res 318(12):1347–1352. https://doi. org/10.1016/j.yexcr.2012.03.025 6. Serrentino ME, Chaplais E, Sommermeyer V, Borde V (2013) Differential association of the conserved SUMO ligase Zip3 with meiotic double-strand break sites reveals regional variations in the outcome of meiotic recombination. PLoS Genet 9(4):e1003416. https://doi.org/ 10.1371/journal.pgen.1003416

7. Chia M, van Werven FJ (2016) Temporal expression of a master regulator drives synchronous sporulation in budding yeast. G3 (Bethesda). https://doi.org/10.1534/g3. 116.034983 8. De Muyt A, Pyatnitskaya A, Andreani J, Ranjha L, Ramus C, Laureau R, FernandezVega A, Holoch D, Girard E, Govin J, Margueron R, Coute Y, Cejka P, Guerois R, Borde V (2018) A meiotic XPF-ERCC1-like complex recognizes joint molecule recombination intermediates to promote crossover formation. Genes Dev 32(3-4):283–296. https://doi.org/10.1101/gad.308510.117 9. Murakami H, Borde V, Nicolas A, Keeney S (2009) Gel electrophoresis assays for analyzing DNA double-strand breaks in Saccharomyces cerevisiae at various spatial resolutions. Methods Mol Biol 557:117–142 10. Hunter N, Kleckner N (2001) The single-end invasion: an asymmetric intermediate at the double-strand break to double-holliday junction transition of meiotic recombination. Cell 106(1):59–70 11. Brachet E, Beneut C, Serrentino ME, Borde V (2015) The CAF-1 and Hir histone chaperones associate with sites of meiotic double-strand breaks in budding yeast. PLoS One 10: e0125965. https://doi.org/10.1371/journal. pone.0125965

Chapter 22 Investigation of Break-Induced Replication in Yeast Beth Osia, Rajula Elango, Juraj Kramara, Steven A. Roberts, and Anna Malkova Abstract Repair of double-strand DNA breaks (DSBs) is important for preserving genomic integrity and stability. Break-induced replication (BIR) is a mechanism aimed to repair one-ended double-strand DNA breaks, similar to those formed by replication fork collapse or by telomere erosion. Unlike S-phase replication, BIR is carried out by a migrating DNA bubble and is associated with conservative inheritance of newly synthesized DNA. This unusual DNA synthesis leads to high level of mutagenesis and chromosomal rearrangements during BIR. Here, we focus on several genetic and molecular methods to investigate BIR using our system in yeast Saccharomyces cerevisiae where BIR is initiated by a site-specific DNA break, and the repair involves two copies of chromosome III. Key words Break-induced replication, Double-strand break, Single-stranded DNA, Homologous recombination, Contour-clamped homogenous electric field electrophoresis, Gross chromosomal rearrangements, APOBEC

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Introduction Double-strand breaks (DSBs) are the most lethal type of DNA damage, typically resulting from problems during DNA replication or chromosome segregation or due to a cell’s exposure to various chemicals or radiation (reviewed in [1]). DSBs can be repaired by nonhomologous end joining (NHEJ) or by homologous recombination (HR). HR uses a homologous template to repair broken DNA and is believed to be more accurate than NHEJ, which proceeds via direct ligation of broken DNA ends. Several HR pathways have been described, including gene conversion (GC) that proceeds by gap repair or by synthesis-dependent strand annealing (SDSA), single-strand annealing (SSA), and break-induced replication (BIR). Here we focus on BIR, which is specific for situations where only one DSB end can find homology in the genome for invasion and repair. BIR starts with extensive 50 to 30 resection of a DSB end followed by strand invasion of a 30 end into a homologous

Andre´s Aguilera and Aura Carreira (eds.), Homologous Recombination: Methods and Protocols, Methods in Molecular Biology, vol. 2153, https://doi.org/10.1007/978-1-0716-0644-5_22, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Yeast experimental system to investigate BIR. (a) Schematic of the AM1003 experimental disome system that contains two copies of chromosome III. One copy contains a Gal::HO endonuclease recognition site (black arrow) at MATa and is truncated by insertion of LEU2 and telomere sequence next to MATa. The other copy of chromosome III is full-length and cannot be cut by HO due to the presence of a MATα-inc (uncuttable) allele. (b) Mechanism of BIR initiation and progression. BIR is initiated by 50 to 30 resection of the one-ended DSB, which leads to the formation of exposed 30 ssDNA. The ssDNA end invades into the homologous template initiating DNA synthesis. Leading strand synthesis progresses by a migrating bubble. ssDNA accumulates behind the migrating bubble due to uncoupled leading and lagging strand synthesis. Completed repair results in conservative inheritance of newly synthesized DNA. (c) Repair outcomes following DSB initiation in AM1003. The following repair outcomes and their phenotypes are depicted: gene conversion (Ade+ Leu+), BIR (Ade+ Leu), chromosome loss (Adered Leu), and half crossover (Adewhite Leu)

DNA donor (Fig. 1a, b). This leads to initiation of extensive DNA synthesis, which is carried out by a migrating DNA bubble, proceeding with asynchrony between leading and lagging strand

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synthesis and leading to conservative inheritance of newly synthesized DNA (Fig. 1b) [2–4]. This abnormal DNA synthesis is responsible for the increased level of mutagenesis and chromosome rearrangements associated with BIR [2, 5–8]. One important source of BIR-associated mutagenesis is long single-stranded DNA (ssDNA) resulting from extensive resection as well as from the asynchrony between leading and lagging strand synthesis. Such long ssDNA accumulates behind the BIR migrating bubble and can accumulate DNA damage, which leads to hyper-mutagenesis associated with BIR. In addition, ssDNA can promote formation of lethal recombination intermediates that can also form during BIR [9]. For several decades, yeast Saccharomyces cerevisiae has been used as a model to investigate BIR. Several different experimental systems were developed that mainly differed in the way the doublestrand break was introduced [10–13]. Here we focus on our experimental system where BIR is initiated by a site-specific HO endonuclease and proceeds between two copies of chromosome III (Fig. 1a) [14]. Using this system has allowed us to perform a detailed study of the mechanism of BIR by employing various genetic and molecular approaches, identify proteins that carry out and regulate BIR, and investigate mutagenesis and gross chromosomal rearrangements associated with BIR [2, 5–7, 14]. Here we focus on determining the efficiency of BIR using genetic methods. In addition, we describe how to determine the amount of ssDNA formed during BIR by using APOBEC3A (A3A)-induced mutagenesis. In particular, APOBEC cytidine deaminase (reviewed in [15]) expressed in yeast converts cytidines in ssDNA accumulated during BIR into deoxyuridines (dUs) [16]. In ung1Δ mutants lacking uracil glycosylase, these dUs are not repaired and all result in C to T mutations, which reveals the full mutagenic potential of A3A during BIR and can be used to characterize the amount of ssDNA formed during BIR. Finally, we describe the investigation of BIR kinetics using Contour-clamped homogenous electric field (CHEF) gel electrophoresis and Southern blot hybridization.

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Materials To investigate BIR, we use our yeast strain AM1003 disomic for chromosome III (Fig. 1a) with the genotype hmlΔ::ADE1/hmlΔ:: ADE3 MATa-LEU2-tel/MATα-inc hmrΔ::HPH FS2Δ::NAT/FS2 leu2/leu2–3112 thr4 ura3–52 ade3::GAL::HO ade1 met13 [14] or its derivatives. In this strain, BIR is initiated by HO endonuclease regulated by a galactose-inducible promoter. A DSB is initiated at the MATa locus of the truncated copy of chromosome III, and the break is repaired by BIR using another intact, full-length copy of chromosome III containing MATα-inc.

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Yeast Media

All media, materials, and solutions used for investigation of BIR by genetic methods need to be autoclaved (unless stated otherwise). 1. Sc-Leu dropout liquid: 2% (w/v) D-glucose, 0.67% (w/v) yeast nitrogen base without amino acids (US Biological, #Y2025), 0.087% (w/v) Leu dropout amino acid mix, pH 5.5. 2. YEP-lactate liquid: 3.15% (v/v) lactic acid, 1% (w/v) yeast extract, 2% (w/v) peptone pH 5.5. 3. 20% (w/v) galactose: 20 g galactose dissolved in 100 mL of MilliQ water and filter sterilized. 4. YEP-galactose plates: 2% (v/v) galactose, 2.5% (w/v) agar, 1% (w/v) yeast extract, 2% (w/v) peptone pH 5.5. (Autoclave everything except for galactose. Following autoclaving, add 20% filter-sterilized galactose (1:10 dilution to final concentration of 2%).) 5. Sc-Ade dropout plates: 2% (w/v) D-glucose, 2.5% agar, 0.67% (w/v) yeast nitrogen base without amino acids, 0.087% (w/v) Ade dropout amino acid mix, pH 5.5. 6. Sc-His dropout plates: 2% (w/v) D-glucose, 2.5% (w/v) agar, 0.67% (w/v) yeast nitrogen base without amino acids, 0.087% (w/v) His dropout amino acid mix, pH 5.5. 7. Sc-Leu dropout plates: 2% (w/v) D-glucose, 2.5% (w/v) agar, 0.67% (w/v) yeast nitrogen base without amino acids, 0.087% (w/v) Leu dropout amino acid mix, pH 5.5. 8. Sc-Ura dropout plates: 2% (w/v) D-glucose, 2.5% (w/v) agar, 0.67% (w/v) yeast nitrogen base without amino acids, 0.087% (w/v) Sc-Ura dropout amino acid mix. 9. Sc-Ura, Ade dropout plates: 2% (w/v) D-glucose, 2.5% (w/v) agar, 0.67% (w/v) yeast nitrogen base without amino acids, 0.087% (w/v) Sc-Ura, Ade dropout amino acid mix. 10. YEPD plates: 2% (w/v) D-glucose, 2.5% (w/v) agar, 1% (w/v) yeast extract, 2% (w/v) peptone, pH 5.5. 11. YEPD liquid: 2% (w/v) D-glucose, 1% (w/v) yeast extract, 2% (w/v) peptone, pH 5.5. 12. 50 mg/mL Hygromycin B stock solution (EMD Millipore #400052). 13. YEPD + Hyg plates: 2% (w/v) D-glucose, 2.5% (w/v) agar, 1% (w/v) yeast extract, 2% (w/v) peptone, pH 5.5, 0.05% (v/v) hygromycin B. (Add 10 mL of 50 mg/mL hygromycin B stock solution per liter.) 14. Sc-Leu dropout + Hyg liquid: 2% (w/v) D-glucose, 0.67% (w/v) yeast nitrogen base without amino acids (US Biological, #Y2025), 0.087% (w/v) Leu dropout amino acid mix,

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pH 5.5, 0.05% hygromycin B (v/v). (Add 10 mL of 50 mg/ mL hygromycin B stock solution per liter.) 15. YEP-lactate + Hyg liquid: 3.15% (v/v) lactic acid, 1% (w/v) yeast extract, 2% (w/v) peptone pH 5.5, 0.05% (v/v) hygromycin B. (Add 10 mL of 50 mg/mL hygromycin B stock solution per liter.) 16. Petri dishes (100  15 mm). 2.2 Yeast Strains and Plasmids

1. AM1003 [14] or its derivatives. 2. AM3647 that is isogenic to AM1003, but ung1Δ::KanMX, hmrΔ::ura3–29-HPH (containing insertion of ura3–29 reporter [17] at 90 kb position centromere distal from MATα-inc). 3. pSR435 and pSR419: centromeric plasmids [18]. pSR435 contains human APOBEC3A gene under the control of the TetO7 promoter. pSR419—empty vector control (similar to pSR435, but without A3A).

2.3 Detecting ssDNA Accumulated on the Course of BIR

All solutions used for the molecular analysis of BIR need to be prepared using MilliQ water. 1. 1 M sorbitol (RPI, 10043–35-3): 18.217 g dissolved in 100 mL MilliQ water. 2. 2 U/μL Zymolyase solution: 0.1 g of 20 T zymolyase (MP, #320921) in 1 mL of 50% (v/v) glycerol. 3. TE buffer: 10 mM Tris, 1 mM EDTA, pH 7.6. 4. 20% (w/v) SDS (Fisher, #BP166-5): 20 g of sodium dodecyl sulfate dissolved in 100 mL MilliQ water. 5. 5 M potassium acetate (KAc): 49.1 g of potassium acetate dissolved in 100 mL MilliQ water. 6. RNase A stock solution: 10 mg of RNase A dissolved in 1 mL MilliQ water. 7. Phenol/chloroform/isoamyl alcohol (25:24:1).

2.4 Assessing BIR Kinetics and BIR Outcomes Using CHEF Gel Electrophoresis

1. 50 mL conical tubes. 2. 10% (w/v) sodium azide solution: 10 g of sodium azide (Sigma Aldrich) dissolved in 100 mL MilliQ water. 3. 50 mM EDTA, pH 7.5: Diluted 1:10 from 0.5 M EDTA in MilliQ water. 4. Plug molds (Bio-Rad #1703713). 5. SCE solution: 0.1 M sorbitol, 1 mM sodium citrate, 50 mM EDTA pH 7.5. 6. 1% (w/v) LMP agarose (Fisher, 9012-36-6): in 0.75 mL water and 0.25 mL 0.5 M EDTA pH 7.5.

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7. Zymolyase buffer: 0.005 g 20 T zymolyase, 1 mL SCE solution, 50 μL 2-mercaptoethanol. 8. Incubation buffer 1: 50 mM EDTA pH 7.5, 0.01 M Tris–HCl pH 7.5. 9. Incubation buffer 2: 0.1 M Tris–HCl pH 7.5, 0.25 M EDTA pH 7.5, 1% (v/v) sarkosyl. 10. Wash buffer: 10 mM Tris pH 7.5, 50 mM EDTA pH 7.5. 11. Storage buffer: 40 mM EDTA pH 7.5, 55% (v/v) glycerol. 12. Proteinase K: RPI, #39450-01-6. 13. Ethidium bromide (Sigma Aldrich). 14. 5 TBE: 64 g Tris base (RPI, #77-86-1), 31 g boric acid, 3.35 g EDTA disodium salt dihydrate (Fisher, BP120-1) dissolved in 1L of MilliQ water. 15. 1 TBE: dilute 5 TBE by adding 200 mL 5 TBE to 800 mL of MilliQ water. 16. 1% Pulse-field certified agarose gel (Bio-Rad, #L004315B): 2 g pulse-field certified agarose in 200 mL 0.5 TBE. 17. 20 SSC pH 7: 175.3 g NaCl, 88.2 g sodium citrate, and 900 mL MilliQ water. 18. Alkaline solution: 20 g NaOH, 87.7 g NaCl in 1 L of MilliQ water. 19. Neutralization solution: 87.7 g NaCl, 121.4 g Tris base in 1 L of MilliQ water, pH 7.0. 20. QIAGEN gel extraction kit: QIAGEN, # 28704. 21.

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22. RmT Random primer kit (Agilent, # 300392). 23. Hybridization buffer: 50 mL of 0.5 M sodium phosphate buffer pH 7.2, 25 mL of 20% SDS, 200 μL of 1 M EDTA pH 8. Make up to 100 mL using MilliQ water. 24. Low Stringency Wash Buffer: 2 SSC, 0.1% (w/v) SDS.

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Methods

3.1 Determining the Efficiency of BIR

1. Inoculate a single colony of AM1003 or its derivative in liquid Sc-Leu dropout media (see Note 1) and grow for 24 h at 30  C to approximately 1  108 cells/mL (to saturation). 2. Transfer 5 mL of the saturated culture (inoculum) from Subheading 3.1, step 1 to 45 mL of YEP-lactate liquid media in 250 mL flask and grow at 30  C with constant aeration for 16 h (see Note 2).

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3. Using a hemocytometer count the cells and calculate the cell density. The expected cell density is approximately 2  107 cells/mL. 4. Perform serial dilutions of the cell culture to reach a density of approximately 5  102 cells/mL by first adding 5  105 cells to 1 mL of sterile water. Perform three serial 1:10 dilutions in sterile water. 5. Plate 0.1 mL of the yeast cell suspension (5  102 cells/mL) on at least three YEPD plates and five to ten YEP-galactose plates. 6. Incubate all plates (YEPD and YEP-galactose) for 3–5 days at 30  C. Usually, the colonies will form in approximately 3 days on YEPD, while it may take approximately 5 days for the colonies to form on YEP-galactose. 7. Cell viability following DSB induction is determined by comparing the number of colonies grown on YEP-galactose to those grown on YEPD. Cell viability should be calculated using the formula: Cell viability ð%Þ ¼

Mean number of colony‐forming units ðc:f :uÞ=per plate of YEP‐Gal  100 : Mean c:f :u:per YEPD plate (see Note 3). 8. Replica plate the colonies grown on YEP-galactose onto Sc-Ade, Sc-Leu, and Sc-His dropout media plates. 9. Score repair outcomes based on auxotrophic marker phenotypes (Figs. 1c and 2a). (a) BIR: Ade+ Leu. (b) Gene conversion: Ade+ Leu+. (c) Half crossover: Ade-white Leu. (d) Chromosome loss: Ade-red Leu. 10. Calculate the frequency of BIR outcomes (FBIR) using the following formula:

F BIR ð%Þ ¼

2  ð# of full BIR coloniesÞ þ # of colonies with a BIR sector  100 : 2  ðtotal coloniesÞ (see Note 4 and Fig. 2b).

3.2 Determining the Amount of ssDNA Formed During BIR

1. Inoculate a single colony of AM3647 (Fig. 3a) transformed with pSR435 (containing A3A gene) into 1 mL of Sc-Leu dropout + Hyg liquid media, and one colony of AM3647 transformed with pSR419 (empty vector control) into 1 mL of Sc-Leu dropout + Hyg liquid media and incubate for 24 h at 30  C with constant aeration until it reaches saturation (to the density approximately 1  108 cells/mL).

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Fig. 2 (a) BIR repair plating assay. Cells were pre-grown in YEP-lactate liquid media followed by plating on YEP-galactose plates. The colonies were replica plated on the Sc-Ade (left) and Sc-Leu (right) dropout plates. The rad9Δ derivative of AM1003 is shown that produces various repair outcomes. Arrows point to different repair outcomes, green arrows indicate a BIR outcome, blue arrows indicate a half crossover, black arrows indicate gene conversion, and red arrows indicate a chromosome loss outcome. In addition, the depicted mutant shows formation of many sectored colonies (containing sectors of several repair outcomes in the same colony), which can result from two sister chromatids repairing differently, or from delayed repair that occurs following one or several cellular divisions. The latter is frequent in DNA damage checkpoint mutants including rad9Δ depicted here. (b) CHEF gel analysis of Ade+Leu repair outcomes, including BIR repair outcomes and repair outcomes containing chromosomal rearrangements. Top: ethidium bromide-stained CHEF gels; middle: Southern blot hybridization with ADE1-specific probe and with ADE3-specific probe (bottom). Red arrow head denotes BIR band. Blue arrow head denotes donor band. Asterisks denote rearranged recipient (ADE1containing) and donor (ADE3-containing) chromosomes

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Fig. 3 Mutations accumulating in ssDNA formed during BIR (From data in Elango et al. [16] published under Oxford University Press’s open access and a Creative Commons Attribution 4.0 International License). (a) The increase of Ura+ reversions resulting from BIR in the presence of expression of APOBEC3A. Top: experimental disomic system (similar to that shown in Fig. 1a, but containing ura3–29 mutation reporter inserted at position

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2. For the culture transformed with the A3A gene (pSR435), transfer 0.5 mL of inoculum into 4.5 mL YEP-lactate + Hyg media (in a 125 mL flask). For the culture transformed with the empty vector control (pSR419), transfer 1 mL of inoculum into 9 mL YEP-lactate + Hyg media (in a 125 mL flask). Both should have a starting cell density of ~1  106 cells/ mL. Incubate all cultures at 30  C for ~16–20 h with constant aeration, to a final concentration of approximately 1–2  107 cells/mL. 3. From the A3A transformed cultures, collect 0.5 mL of culture from Subheading 3.2, step 2 into a conical tube, and likewise from the empty vector transformed cultures collect 3 mL of the culture from Subheading 3.2, step 2 into a conical tube. Use these as the “0 h” time point to assess the level of mutagenesis associated with S-phase replication (as a control). 4. Take an additional 0.5 mL from all cultures from Subheading 3.2, step 2 and serially dilute them by transferring 0.1 mL of culture into 0.9 mL sterile water. Typically, four tenfold dilutions are made to obtain a concentration of approximately 1  103 cells/mL. 5. Plate each suspension from Subheading 3.2, step 4 with a concentration of approximately 1  103 cells/mL by plating 0.1 mL per Sc-Ade dropout media (three Sc-Ade dropout plates total with approximately 100 cells per plate). From these plates, the concentration of cells in the culture will be determined. 6. To estimate the number of mutations accumulated in the culture at 0 h (before induction of the DSB by HO), centrifuge the samples collected in Subheading 3.2, step 3 at 5000  g for 2 min at 25  C. Decant the supernatant and keep the pellet. Resuspend the pellet in 0.5 mL of sterile water and spread onto five Sc-Ura, Ade dropout plates (0.1 mL per plate). Be cautious not to plate >1  108 cells per plate to avoid formation of a lawn due to residual growth. 7. Add 440 μL of 20% galactose (filter sterilized) to the remaining 4 mL of the A3A culture from Subheading 3.2, step 2 and 715 μL of 20% galactose (filter sterilized) to the remaining 6.5 mL of the ä Fig. 3 (continued) 90 kb centromere distal to MATα-inc). Bottom: Ura+ rates for ura3–29 inserted at 90 kb Chr. III position following BIR in the presence of APOBEC3A containing plasmid (A3A) or empty vector (EV). Asterisk and pound symbol indicate statistically significant differences from No-DSB (0 h) and from no-A3A (EV), respectively. (b) A3A-induced mutations (black vertical lines) in one BIR outcome in the progeny of ung1Δ derivative of AM1003 formed in the presence of A3A. Enlarged: mutation cluster on the track of BIR. (c) Clustered mutations in BIR isolates formed in the presence of A3A. Positions of mutations (black lines) are depicted along the chromosome III reference. The total number of mutations in each cluster is indicated by a number (on the right)

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empty vector culture from Subheading 3.2, step 2, to obtain a final concentration of 2% galactose in all cultures. The addition of galactose initiates formation of the DSB. Incubate with constant aeration at 30  C for 7 h. 8. For A3A cultures, collect 25 μL of culture from Subheading 3.2, step 7, and from empty vector cultures, collect 2 mL of culture. Subheading 3.2, step 7 (see Note 5). Centrifuge at 5000  g for 2 min at 25  C. Decant the supernatant (or remove with a pipette for small volumes) and resuspend each pellet in 0.5 mL of sterile water and plate onto five Sc-Ura dropout plates (0.1 mL of suspension per plate). Be cautious to not plate >1  108 cells per plate to avoid formation of a lawn. 9. Take an additional 0.5 mL from all cultures from Subheading 3.2, step 7 and serially dilute them by transferring 0.1 mL of culture into 0.9 mL sterile water. Typically, four 10-fold dilutions are made to obtain a concentration of approximately 1  103 cells/mL. 10. Plate each suspension from Subheading 3.2, step 9 with a concentration of approximately 1  103 cells/mL by plating 0.1 mL per YEPD plate (three YEPD plates total with approximately 100 cells per plate). From these plates, the concentration of cells in the culture will be determined. 11. Incubate all plates for 5 days at 30  C and count the colonies on each plate. 12. Calculate the rate of Ura+ reversions (see Note 6) associated with BIR using the following formulas: Uraþ BIRrate ¼ Uraþ f requency ð7 hÞ  Uraþ f requency ð0 hÞ: Total number of colonies on all Ura dropoutplates ð0 hÞ : Uraþ f requency ð0 hÞ ¼ Mean number of colonies on YEPD plates ð0 hÞ  105 Total number of colonies on all Ura dropout plates ð7 hÞ Uraþ f requency ð7 hÞ ¼ : Mean number of colonies on YEPD plates ð7 hÞ  105 (see Notes 6 and 7). It is expected that the Ura+ BIR rate (at 7 h) will significantly (more than ten times) exceed the Ura+ rate at 0 h for both cells transformed with A3A plasmid and with empty vector. In addition, Ura+ BIR rate in cells with A3A vector is expected to be more than 100 times higher than in cells transformed with empty vector [16] (Fig. 3a) (see Note 8). 13. Pick Ura+ colonies from Sc-Ura dropout plates (7 h, formed after BIR) from the culture containing the A3A expression plasmid, and streak on SC-Ura dropout plates to isolate single colonies. 14. Take the singles from the last step and patch them on YEPD, grow at 30  C.

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15. Replica plate patches on Sc-Leu, Sc-Ade, Sc-Ura, and also on YEPD + Hyg plates to confirm that the colonies have completed BIR repair, that they are Ura+, and that they lost the A3A-containing plasmid (They should be HygsAde+ Ura+Leu). 16. Use the patch from Subheading 3.2, step 14 to start a culture in 50 mL YEPD, grow to saturation (approximately 18–24 h). 17. Pellet the culture from Subheading 3.2, step 16 by centrifugation at 5000  g for 5 min. 18. Resuspend yeast pellet in 5 mL of 1 M sorbitol. 19. Add 500 μL of 2 U/μL zymolyase solution. Mix by vortexing. 20. Incubate for at least 1 h at 37  C and periodically agitate gently. 21. Centrifuge at  4000  g for 7 min. Discard as much as possible of the supernatant. 22. Resuspend the cells in 10 mL of TE buffer. 23. Add 1.1 mL of 20% SDS. Invert the tube quickly 20 times to mix. 24. Incubate at 65  C for 30 min (see Note 9). 25. Add 11 mL 5 M KAc. Invert the tube gently ten times to mix. Set the tube on ice for 30–60 min. 26. Centrifuge for 15 min at 14,000  g. Transfer 10–15 mL of supernatant into a fresh centrifuge tube (see Note 10). 27. Add equal volume of room temperature isopropanol to the supernatant. Invert the tube ten times to mix. 28. Allow the tube to sit at room temperature for 15 min (see Note 11). 29. Centrifuge for 10 min at 14,000  g. Pour off the supernatant. 30. Partially air-dry the pellet for 15 min. Place the tube upside down on a napkin. 31. Add 20 mL of 70% ethanol (EtOH). Invert the tube ten times to mix. 32. Allow the tube to sit overnight at 4  C. 33. Centrifuge for 2 min at 14,000  g. Pour the supernatant away immediately once it is removed from centrifuge (see Note 12). Allow the tube to sit with its lid open to dry further. 34. Resuspend yeast genomic DNA in 2 mL TE. Transfer to a 15 mL polypropylene centrifuge tube. 35. Add 30 μL of RNase A stock solution. Incubate at 37  C for 30 min.

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36. Add 2 mL of phenol/chloroform/isoamyl alcohol mix. Vortex and spin tubes at 4000  g for 10 min. 37. Take the top phase (try not to get white protein at phase interface; about 1.8 mL). 38. Add 100 μL of 5 M KAc and mix gently by inverting the tube. 39. Split into five aliquots in 1.5 mL microcentrifuge tubes (400–500 μL per tube). Precipitate with 2 volume (800–1000 μL) of 100% ethanol. Spin all aliquots in a microcentrifuge at 14,000  g for 10 min (see Note 13). 40. Wash twice with 70% ethanol, then leave the tubes open to allow the DNA to dry. 41. Dissolve dry DNA in a total of 0.5 mL TE (100 uL per DNA pellet). 42. Submit 2 μg of purified yeast genomic DNA for library preparation/Illumina sequencing. Library preparation can be performed using KAPA DNA Hyper kit (Kapa Biosystems). Whole genome sequencing can be performed with the HiSeq4000 PE 2  150 (Illumina) sequencing platform. 43. Once sequencing reads are received, perform read alignment to a reference and mutation calling using the CLC Genomic Workbench version 7.5 software or later, specifically: (a) Trim (if poor sequencing quality necessitates) and align the sequence reads to a reference genome of AM1003 that was created for the [5] study, but was tailored to the AM3617 strain using the “Trim Reads” and “Map Reads to Reference” functions of CLC software. (b) Use the aligned read track for mutation calling using the “Fixed Ploidy Variant Detection tool” with ploidy parameter set to 2, requiring SNVs to have greater than 23% allele frequency, and having a read coverage greater than 10% of the mean coverage across the samples. (c) Filter out preexisting variants and SNVs due to mapping artifacts. Parental variants can be removed with the “Remove Variants Present in Control Reads” function. Alternatively, remove mutations that occur in more than one sample (including a list of SNVs generated for an untreated parental control strain). (d) Use the filtering tool to remove all types of mutations from the resulting list except for single nucleotide variants (SNVs). Unless indicated otherwise, all the parameters are kept at default values (see Note 14).

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44. Identify mutation clusters using the Cluster Finder software developed by Dmitry Gordenin’s lab [19, 20] (see Note 15). The length of mutation clusters identified in ung1Δ serves as an estimate of the ssDNA length formed during BIR [16] (see Fig. 3b, CF for examples). 3.3 Investigation of BIR Kinetics and Outcomes Using CHEF Gel Electrophoresis

1. Inoculate a single colony of AM1003 or its derivative in 50 mL of Sc-Leu dropout media, and incubate for 18–24 h at 30  C with constant aeration until the culture reaches a density of ~108 cells/mL. 2. Add the inoculum from Subheading 3.3, step 1 (50 mL) to 500 mL of YEP-lactate in a 2 L-flask. Incubate the culture at 30  C with constant aeration to reach a cell density of ~2  107 cells/mL (should take approximately 16 h). 3. Collect 50 mL of culture for a 0 h sample (sample taken before galactose addition) in a 50 mL conical tube, and then add 0.5 mL of 10% sodium azide to this aliquot. Mix thoroughly. 4. Induce the DSB by adding 50 mL of 20% galactose to the YEP-lactate culture (see Note 16). 5. Use a tabletop centrifuge to pellet the sample collected at time point “0 h” (5000  g for 5 min). 6. Discard the supernatant into hazardous waste container, and resuspend the pellet in 50 mL of 50 mM EDTA pH 7.5 solution. 7. Pellet the sample using a tabletop centrifuge (spin at 5000  g for 5 min). Discard supernatant. Pellets can be stored in 80  C freezer until needed for plug preparation. 8. Collect 50 mL of culture every hour (or every 2 h) and repeat Subheading 3.3, step 5–7 (see Notes 17 and 18). 9. When ready to prepare DNA plugs, thaw all pellets from the time-course assay on ice. 10. Prepare LMP agarose by adding 0.01 g of LMP agarose to 0.75 mL of MilliQ water, and then adding 0.25 mL 0.5 M EDTA pH 7.5. Heat LMP agarose on heating block to 100  C to dissolve the LMP agarose. 11. Prepare zymolyase buffer by adding 50 μL of 2-mercaptoethanol and 0.005 g 20T zymolyase to 1 mL of SCE. Keep on ice. 12. Equilibrate LMP agarose to 40  C by placing in 40  C water bath (see Note 19). 13. Use a vortex to loosen the pellet. Equilibrate the pellet in a 40  C water bath. 14. Quickly mix 0.085 mL zymolyase buffer with 0.415 mL of agarose, and then add 0.25 mL cells to this mixture.

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15. Transfer quickly the cell agarose mixture into plug molds by pipetting (see Note 20). 16. Chill plug molds containing cell agarose mixture in refrigerator at 4  C for approximately 5 min. 17. Prepare incubation buffer 1 mix by adding 750 μL mercaptoethanol and 10 μL 10 mg/mL RNase into 10 mL of incubation buffer 1. Dispense 2.5 mL of incubation buffer 1 mixture into 50 mL conical tubes. 18. Remove solidified plugs from the plug molds into conical tubes containing incubation buffer 1 by a toothpick. Incubate at 37  C in a water bath for 1 h (see Note 21). 19. Discard the incubation buffer from conical tubes; use a scoopula to keep the plugs in place. 20. Rinse plugs with MilliQ water, and dump the water. Use a scoopula to keep plugs in place. 21. Prepare incubation buffer 2 mix by adding 0.005 g proteinase K into 10 mL of incubation buffer 2. 22. Add 2.5 mL of incubation buffer 2 mixture into each conical tube with plugs. Incubate at 50  C for 16 h. 23. Rinse plugs with MilliQ water. Use a scoopula to keep the plugs in place. 24. Rinse plugs with 2.5 mL of wash buffer. 25. Plugs can be stored in wash buffer for two days at 4  C before they are used for CHEF gel electrophoresis. For long-term storage, decant wash buffer, and use scoopula to keep the plugs. Add 50 mL of storage buffer; keep at 20  C for up to 6 months (see Note 22). (a) Make 3 L of 0.5 TBE by adding 300 mL of 5 TBE and 2.7 L of MilliQ water in a 4 L beaker. (b) Assemble gel casting tray (a part of CHEF Bio-Rad equipment) with black base plate on the bottom. Make sure to tighten screws on each side. (c) Remove plugs using a scoopula and rinse in wash buffer. Place each plug against the longer side of the gel casting tray. (d) Prepare 1% pulse-field agarose gel by melting 2 g agarose in 200 mL of the previously made 0.5 TBE in a microwave oven. Allow the gel to cool to about 60  C before pouring it into gel casting tray (see Note 23). 26. Transfer the remaining 0.5 TBE buffer into CHEF electrophoresis cell. Circulate and cool the buffer to 14  C. 27. Once the gel solidifies, disassemble the gel casting tray while leaving the gel adhered to the black plate (see Note 24).

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28. Place the gel and the black base plate into frame in the middle of the CHEF electrophoresis cell. Close the lid of the CHEF electrophoresis cell. 29. Enter the parameters for the initial and final switch times, the duration of the run, and the voltages on the control module. These parameters will vary depending on the sizes of yeast chromosomes that you aim to separate. If you are aiming to separate the donor chromosome III (355 kb) from the repaired by BIR chromosome III (345 kb) of AM1003, and using CHEF DRII apparatus, the parameters should be as follows: initial switch time of 10 s, final switch time of 35 s, running time of 40 h, and the voltage is 6 V/cm (see Note 25). 30. When the CHEF run is finished, add 300 mL 0.5 TBE into a glass baking dish. 31. Remove the CHEF gel from the CHEF apparatus and place into the glass baking dish that contains 0.5 TBE. 32. Add ethidium bromide to a final concentration of 0.5 μg/mL into the glass dish containing the CHEF gel and 0.5 TBE. Incubate for 30 min with constant gentle shaking. 33. Carefully remove the gel from black base plate. Take a picture of the CHEF gel with separated chromosomes using a UV gel imager. 34. Carefully remove buffer with ethidium bromide, and rinse the gel with water. Dispose the EtBr containing solution appropriately. Place the gel into UV crosslinker and set the exposure to 600 μJ/cm2 at 254 nm wavelength. Expose the gel to this UV dose. 35. Submerge the gel into alkaline solution, and incubate for 30 min with constant gentle shaking. Decant alkaline solution. 36. Submerge gel in neutralization solution and incubate for 30 min with constant gentle shaking. 37. To transfer gel, construct gel transfer sandwich in the following order from bottom to top: glass tray containing 10 SSC, plexiglass surface, filter paper wick cut to the width of the gel that contacts the SSC below and is wetted with 10 SSC, CHEF gel (DNA side up), Nylon transfer membrane wetted with 2 SSC, filter paper cut to the size of the gel wetted in 10 SSC, two stacks of paper towels (~20 cm high), second plexiglass surface, and 500 g weight. Capillary transfer methods further described in [21]. Let the transfer occur for 20 h. 38. Place the membrane (DNA side up) in the UV crosslinker. Expose the membrane to UV (254 nm wavelength) using the auto-crosslink function (120 mJ/cm2). 39. Prepare the DNA probes for Southern blot hybridization.

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Fig. 4 BIR kinetics assay using CHEF gel electrophoresis (From data in Elango et al. [9] published under open access and a Creative Commons Attribution 4.0 International License). (a) The BIR repair kinetics in wild-type (SRS2) strain isogenic to AM 1003 (left) and BIR-defective srs2Δ derivative. The aliquots were taken from the liquid cultures undergoing BIR repair before (0 h) and following the addition of galactose to the media at indicated time points. The genomic DNA was separated by CHEF. The upper panels represent ethidium bromide-stained gels of the separated chromosomes. The middle panels represent the results of Southern blot analysis using an ADE3-specific probe that highlights the donor chromosome III. The bottom panels represent the results of Southern blot analysis using an ADE1-specific probe highlighting the recipient chromosome III. Chr.: chromosome; Trunc. Chr III: truncated chromosome III. (b) Quantification of BIR product (top) and of donor chromosome entering the gel (bottom) in wild-type (SRS2) and srs2Δ strains from the results of analysis shown in (a)

(a) ADE1-specific DNA probe targets not only the recipient copy of chromosome III (before and after the DSB, and also following DSB repair) but also chromosome I containing a native ADE1 gene (see Fig. 4a). This probe is prepared by PCR amplification of the ADE1 fragment from the yeast genome (using genomic DNA prepared from AM1003) and using the following PCR primers: FP: 50 -GGTTTGAAACAACCTCAAGGACTT-30 . RP: 50 -AAGTCCTTGAGGTTGTTTCAAACC-30 . (b) ADE3-specific DNA probe targets the donor copy of chromosome III (before and after the DSB, and also following DSB repair) but also chromosome VII containing the native ADE3 gene. This probe is prepared by PCR amplification of the ADE3 fragment from the yeast genome (using genomic DNA prepared from AM1003) and using the following PCR primers: FP: 50 -GCAGGGTTCGATTTCACTATGGGT-30 . RP: 50 -ACCGTCATCATCGACTTCCACGGC-30 .

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40. Run the ADE1-specific or ADE3-specific probe fragment(s) on a 1% agarose 1 TBE gel and excise the fragment(s). 41. Using the QIAGEN gel extraction kit extract the probe PCR fragment. 42. Resuspend and dissolve the contents of one tube of the RmT Random primer kit in 37 μL of MilliQ water, and transfer to a 1.5 mL microcentrifuge tube. 43. Add 5 μL (~50 ng) of the gel-extracted PCR probe fragment. 44. Incubate RmT Random primer kit and probe fragment mixture at 100  C for 5 min. Be sure to add a microcentrifuge tube locker to keep the lid closed. 45. After incubation period, place reaction mixture on ice for 1 min and briefly spin down (to collect all the liquid on the bottom of the tube). 46. Add 5 μL of 32P-dCTP to the cooled reaction mixture, and add 3 μL of Magenta polymerase from the RmT Random primer kit, and incubate at 37  C dry bath for 10 min. 47. Add 2 μL of stop mix from the RmT Random primer kit to stop the reaction. 48. Add 200 μL of MilliQ water to the reaction mixture and denature at 100  C dry bath for 10 min and snap cool on ice for 1 min. 49. Pre-hybridize the crosslinked membrane from step 14 in 25 mL of hybridization buffer at 65  C for at least 10 min. 50. Hybridize the membrane at 65  C with 50 μL of the labeled probe diluted in 25 mL of hybridization buffer for 24 h [22]. 51. Use low stringency wash buffer preheated to 65  C to wash the membrane in an open tray. Gently rock the wash buffer and membrane for 10 min. Discard the waste in a special radioactive waste container. 52. Repeat the same wash two more times or until background is significantly diminished, as being checked using a Geiger counter. 53. Using 20 cm  25 cm GE Phosphor screen (or similar screen), expose membrane for 1 day and image on a Typhoon Phosphorimager FLA7000 (or similar brand) (see Fig. 4a for examples of images). 54. To calculate the amount of BIR product present in the culture at each time point, the intensity of BIR product (345kb band) hybridizing to ADE1-specific probe should be measured using ImageQuant (GE Healthcare Life Sciences). In addition, the intensity of an intact, truncated copy of chromosome III (243kb long) that also hybridizes to ADE1-specific probe and that is present at the time “0” should be also calculated. To

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account for variation in DNA loads, intensities of the bands corresponding to the intact chromosome III, as well as to the repaired chromosome III, should be normalized to intensities of the bands corresponding to chromosome I, which also hybridizes to the ADE1-specific probe. The efficiency of BIR repair, presented as the percentage of truncated chromosome III that was converted to BIR product, should be calculated by dividing the normalized intensity of a repair band by the normalized intensity of uncut, truncated chromosome III. Results of three time-course experiments can be used to calculate the average  SD BIR efficiency at each time point for each strain. The intensities of the template (donor) Chr. III molecule (355kb band) at each time point (visualized by hybridization with radioactively labeled ADE3 probe) should be normalized to intensities of the bands corresponding to chromosome VII, which also hybridizes to the ADE3-specific probe. In wild-type cells, the amount of the donor DNA should remain constant throughout the course of BIR. However, in some mutants, for example, in srs2Δ, the amount of the donor is drastically decreased following the initiation of BIR (for example, at 8 h time point following the initiation of BIR) (Fig. 4a, b) [9]. This decrease of donor molecules in the agarose gel in srs2Δ is indicative of the accumulation recombination intermediates as branched DNA structures. The percent decrease of the donor DNA (Fig. 4b) can be used to estimate the amount of branched lethal intermediates formed in the course of BIR.

4

Notes 1. Sc-Leu dropout media is used for the inoculum to eliminate cells that underwent spontaneous BIR and therefore became Leu prior to the beginning of the experiment. 2. The lack of glucose in the media enables the efficient galactose uptake and the maximum HO induction, and therefore DSB initiation. It is expected that more than 90% of cells will undergo HO-induced DSB in less than 20 min following galactose addition. 3. Loss of viability after the DSB induction is rare due to the design of the disomic system that contains two chromosomes III (Fig. 2a). These cells remain viable even when repair of the one (broken) chromosome is unsuccessful and it is lost. However, the loss of viability can occur due to more complex events affecting both recombining chromosomes [9]. 4. It is important to mention that not all Ade+ Leu events have been proven to be BIR. Some Ade+ Leu outcomes contain gross chromosome rearrangements (GCRs). BIR can be

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distinguished from GCR outcomes using CHEF gel electrophoresis. In particular, BIR events without chromosomal rearrangements contain two copies of chromosome III: a 355 kb chromosome that hybridizes to ADE3-specific probe and a 345 kb chromosome that hybridizes to an ADE1-specific probe. BIR outcomes with rearrangements are defined as having two copies of Chr. III where at least one of the copies deviates from its expected size (see Fig. 2b for examples). The fraction of GCRs among Ade+Leu should be taken into account when calculating BIR efficiency (see in [6] for details). 5. Typically, for wild-type strains, 2-5 mL samples are collected at 7 h. However, for certain mutants, more cells need to be collected due to a lower mutation rate. 6. The ura3–29 mutation located at 90 kb position centromere distal from MATα-inc can revert to a Ura+ phenotype via C to T, C to G, or C to A base substitutions [17]. 7. The best way to determine the mutation rate during S-phase is to use a no-DSB control strain. A no-DSB strain has a genotype similar to AM1003 except it contains Matα-inc-LEU2-tel, and in these strains, DSBs cannot be induced by HO endonuclease. 8. Due to the accumulation of mutations over several generations, the spontaneous mutagenesis rate (the rate of mutations during S-phase replication) is calculated using Drake equation: μ ¼ 0.4343 f/log (Nμ), where μ is the rate of spontaneous mutagenesis, f is mutation frequency, and N is the number of cells in yeast culture [23]. However, during BIR, the cells are arrested at G2/M by DSB induction until they are repaired, and then they recover from the arrest. This means that there are no cell divisions between 0 and 7 h time points following galactose addition. This is why the mutation rate during BIR can be calculated using a modified Drake equation (Mutation rate during BIR ¼ Mutation frequency (7 h)-Mutation frequency (0 h)). 9. It is expected that in ung1Δ cells transformed with pSR419 (empty vector), the S-phase Ura+ rate will be approximately 1  107, while the BIR Ura+ rate will be approximately 1  106, while in the cells containing pSR435 (A3A vector), the S-phase and BIR Ura+ rates are expected to be approximately 1  105 and 5  104, respectively (Fig. 3a). 10. The tubes can also be kept overnight at ambient temperature. In addition, ice bath can be prepared at this point. 11. Do not disturb the pellet. It may help to filter through sterile cheesecloth. 12. During this time, you can prepare napkins on lab bench.

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13. Be careful not to lose the pellet. Use pipette to remove rest of the liquid. Place the tip of the pipette on the bottom of the tube next to the pellet. 14. You may need to split the sample before spinning—for example, split into four 1.5 mL Eppendorf tubes ~ 0.5 mL in each. 15. The procedure can be streamlined by assembling all the steps into a single workflow. It is also possible to perform all the steps with some tools available in open access, including Bowtie (for the alignments) and GATK (for mutation calling). 16. The Cluster Finder program uses the filtered mutation calls list from the previous step as an input. The mutation table needs to be reformatted first according to the authors’ instructions. Enter the path to the modified table into the control script and then execute the control script to launch the cluster search program. 17. DSB formation should occur within the first hour following the addition of galactose. 18. BIR repair products appear usually at 4–6 h following galactose addition (DSB induction). 19. To prevent degradation of zymolyase, the LMP agarose mix should not be hot at the time of mixing with zymolyase. 20. While transferring cell mixture, with LMP agarose and zymolyase, into plug molds, avoid air bubbles in the mold so as to have a solid plug. 21. A pressurized stream of air can also be used to remove plugs from the plug mold as an alternative to a toothpick. 22. For long-term storage of the plugs, the volume of storage solution should be at least half of the storage tube to prevent freezing of the plugs as well as to protect against breaking of the plugs. 23. While you are pouring the gel into the casting tray, make sure that the holes in each corner of the black plate are filled without air bubbles. 24. When removing the gel from the casting tray, make sure to remove as much excess gel from the edges and bottom of the black plate. 25. For different CHEF machines, the optimal running parameters to separate the chromosomes of interest vary.

Acknowledgments A.M. is supported by R35GM127006 grant from NIGMS, R03ES029306, and R21 ES030307 from NIEHS. S.A.R. is supported by R01 award CA218112 from the National Cancer Institute.

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References 1. Mehta A, Haber JE (2014) Sources of DNA double-strand breaks and models of recombinational DNA repair. Cold Spring Harb Perspect Biol 6(9):a016428 2. Saini N et al (2013) Migrating bubble during break-induced replication drives conservative DNA synthesis. Nature 502(7471):389–392 3. Wilson MA et al (2013) Pif1 helicase and Poldelta promote recombination-coupled DNA synthesis via bubble migration. Nature 502 (7471):393–396 4. Donnianni RA, Symington LS (2013) Breakinduced replication occurs by conservative DNA synthesis. Proc Natl Acad Sci U S A 110 (33):13475–13480 5. Sakofsky CJ et al (2014) Break-induced replication is a source of mutation clusters underlying kataegis. Cell Rep 7(5):1640–1648 6. Vasan S et al (2014) Cascades of genetic instability resulting from compromised breakinduced replication. PLoS Genet 10(2): e1004119 7. Sakofsky CJ et al (2015) Translesion polymerases drive microhomology-mediated break-induced replication leading to complex chromosomal rearrangements. Mol Cell 60 (6):860–872 8. Smith CE, Lam AF, Symington LS (2009) Aberrant double-strand break repair resulting in half crossovers in mutants defective for Rad51 or the DNA polymerase delta complex. Mol Cell Biol 29(6):1432–1441 9. Elango R et al (2017) Break-induced replication promotes formation of lethal joint molecules dissolved by Srs2. Nat Commun 8 (1):1790 10. Malkova A, Ivanov EL, Haber JE (1996) Double-strand break repair in the absence of RAD51 in yeast: a possible role for breakinduced DNA replication. Proc Natl Acad Sci U S A 93(14):7131–7136 11. Malkova A et al (2005) RAD51-dependent break-induced replication differs in kinetics and checkpoint responses from RAD51mediated gene conversion. Mol Cell Biol 25 (3):933–944 12. Morrow DM, Connelly C, Hieter P (1997) “Break copy” duplication: a model for

chromosome fragment formation in Saccharomyces cerevisiae. Genetics 147(2):371–382 13. Davis AP, Symington LS (2004) RAD51dependent break-induced replication in yeast. Mol Cell Biol 24(6):2344–2351 14. Deem A et al (2008) Defective break-induced replication leads to half-crossovers in Saccharomyces cerevisiae. Genetics 179(4):1845–1860 15. Roberts SA, Gordenin DA (2014) Clustered and genome-wide transient mutagenesis in human cancers: hypermutation without permanent mutators or loss of fitness. BioEssays 36 (4):382–393 16. Elango R et al (2019) Repair of base damage within break-induced replication intermediates promotes kataegis associated with chromosome rearrangements. Nucleic Acids Res 47 (18):9666–9684 17. Shcherbakova PV, Pavlov YI (1996) 30 -->50 exonucleases of DNA polymerases epsilon and delta correct base analog induced DNA replication errors on opposite DNA strands in Saccharomyces cerevisiae. Genetics 142 (3):717–726 18. Hoopes JI et al (2017) Avoidance of APOBEC3B-induced mutation by error-free lesion bypass. Nucleic Acids Res 45 (9):5243–5254 19. Chan K et al (2015) An APOBEC3A hypermutation signature is distinguishable from the signature of background mutagenesis by APOBEC3B in human cancers. Nat Genet 47 (9):1067–1072 20. Saini N et al (2016) The Impact of environmental and endogenous damage on somatic mutation load in human skin fibroblasts. PLoS Genet 12(10):e1006385 21. Sambrook J, Russell DW (2006) Southern blotting: capillary transfer of DNA to membranes. CSH Protoc 2006(1):pii: pdb. prot4040 22. Church GM, Gilbert W (1984) Genomic sequencing. Proc Natl Acad Sci U S A 81 (7):1991–1995 23. Deem A et al (2011) Break-induced replication is highly inaccurate. PLoS Biol 9(2):e1000594

Chapter 23 Measurement of Homologous Recombination at Stalled Mammalian Replication Forks Nicholas A. Willis and Ralph Scully Abstract Site-specific replication fork barriers (RFBs) have proven valuable tools for studying mechanisms of repair at sites of replication fork stalling in prokaryotes and yeasts. We adapted the Escherichia coli Tus-Ter RFB for use in mammalian cells and used it to trigger site-specific replication fork stalling and homologous recombination (HR) at a defined chromosomal locus in mammalian cells. By comparing HR responses induced at the Tus-Ter RFB with those induced by a site-specific double-strand break (DSB), we have begun to uncover how the mechanisms of mammalian stalled fork repair differ from those underlying the repair of a replication-independent DSB. Here, we outline how to transiently express the Tus protein in mES cells, how to use flow cytometry to score conservative and aberrant repair outcomes, and how to quantify distinct repair outcomes in response to replication fork stalling at the inducible Tus-Ter chromosomal RFB. Key words Homologous recombination, Sister-chromatid recombination, Replication fork barrier (RFB), Short-tract gene conversion, Long-tract gene conversion, Tandem duplication, Tus-Ter, GFP, RFP, Mouse embryonic stem cell, Flow cytometry

1

Introduction Replication fork stalling (sometimes termed “replication stress”) is an important trigger of genomic instability in human diseases, including cancer and developmental disorders [1, 2]. The use of site-specific chromosomal replication fork barriers (RFBs) in bacteria and yeast has greatly facilitated the study of stalled fork repair in these model organisms [3–12]. However, progress in understanding stalled fork repair in mammalian cells has been hindered by a dearth of tractable tools for establishing a site-specific chromosomal RFB. To understand how stalled fork repair is regulated in mammalian cells and how these repair processes become corrupted in certain human disease states, we adapted the Escherichia coli Tus-Ter RFB [13–16] for use in mammalian cells and used it to trigger site-specific replication fork stalling at a defined

Andre´s Aguilera and Aura Carreira (eds.), Homologous Recombination: Methods and Protocols, Methods in Molecular Biology, vol. 2153, https://doi.org/10.1007/978-1-0716-0644-5_23, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Tus-Ter-induced stalled fork repair outcomes in mammalian cells. 6Ter reporter and Tus-Ter-induced homologous recombination and aberrant repair products. Fork stalling (represented as a bidirectional fork block) induces sisterchromatid recombination measured by gene conversion of a mutant green fluorescent protein (GFP)-encoding allele to wild-type GFP. Grey-blue boxes, mutant GFP. Green box, wild-type GFP. Ovals A and B denote 50 and 30 artificial RFP exons. Tr-GFP, promoterless 50 -truncated GFP heteroallele. Orange triangle represents 6Ter element array adjacent to the I-SceI endonuclease cut site (blue vertical line). STGC short-tract gene conversion; LTGC long-tract gene conversion, TD tandem duplication. LTGC and TD repair outcomes produce wildtype RFP expression through RNA splicing

chromosomal locus in living mammalian cells [17]. This system enables a direct comparison between repair induced by the Tus-Ter RFB and repair triggered by a replication-independent chromosomal double-strand break (DSB) produced by the I-SceI homing endonuclease [18]. The general structure of the Ter-HR reporter and detectable repair outcomes, as measured by flow cytometry, are summarized in Fig. 1. The reporter contains two nonfunctional heteroalleles of the gene encoding the enhanced green fluorescent protein (GFP), each lacking essential sequence required for function: the 50 -truncated GFP allele (“50 Tr-GFP”) lacks a promoter and contains an N-terminal truncation; within the 6Ter-GFP allele, five nucleotides in the central region of the GFP open reading frame are replaced with an array of six 23 bp Ter elements and a single I-SceI endonuclease target site. These two GFP heteroalleles flank an additional “RFP cassette,” consisting of two artificial exons derived from the RFP(3.1) cDNA [19, 20], splice donor and acceptor elements, and independent promoter and polyadenylation

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sequences. The two exons are in reverse order, so that a single copy of the RFP cassette does not result in the expression of wild-type RFP [20]. The full HR reporter is targeted as a single copy to the Rosa26 locus of mouse chromosome 6. Transient expression of Tus or I-SceI induces site-specific replication fork stalling or site-specific DSB induction, respectively, within the 6Ter-GFP allele of the HR reporter. The cellular responses to these triggers include the induction of homologous recombination (HR), which can be quantified by conversion of the HR reporter cells to GFP+ and/or RFP+. The 6Ter-HR reporter allows for the rapid detection and quantitation of three distinct repair outcomes, each marked by GFP or RFP protein expression and therefore detectable by flow cytometry. An overview of these repair events is described below and in Fig. 1. 1. Short-tract gene conversion (STGC): STGCs are detected as GFP+RFP products, reflecting HR between the two GFP heteroalleles. Gene conversion converts the 6Ter-GFP allele to wild type. In normal cells, STGC is the major HR product in response to either Tus-Ter-induced fork stalling or an I-SceIinduced DSB [17, 20]. Structural analysis of Tus-Ter-induced STGCs revealed that they are noncrossover products of two-ended HR [17, 21]. This, in turn, suggests that Tus-Terinduced STGCs are products of bidirectional replication fork stalling at the Tus-Ter RFB, and that they engage the synthesisdependent strand annealing (SDSA) pathway of somatic HR [17, 22, 23]. Tus-Ter-induced STGC therefore likely represents a physiological, conservative, error-free HR outcome. Consistent with this, Tus-Ter-induced STGC is mediated by the canonical BRCA/Fanconi anemia/Rad51 pathway [17, 21]. In contrast to STGC induced by a replicationindependent DSB, Tus-Ter-induced STGC is not subject to competition by classical nonhomologous end joining (cNHEJ) [24]. Further, the accumulation of Rad51 at a Tus-Ter RFB is more intense and more tightly focused than at an ISceI-induced DSB. These observations suggest that Tus-Terinduced STGC entails DNA structural intermediates distinct from those formed during the repair of a replicationindependent DSB [23]. 2. Long-tract gene conversion (LTGC): LTGCs are detected as GFP+RFP+ products. LTGC is an error-prone, replicative HR outcome, normally observed as a minor proportion of all detectable HR products for both Tus-Ter-induced and I-SceIinduced repair [17, 20, 25–27]. LTGCs arise by gene conversion between the two GFP heteroalleles, but extended copying from the donor sister chromatid during gene conversion results in the expansion of the reporter to generate three tandem

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copies of GFP (the second of which is wild type; Fig. 1). In the context of GFP triplication, the RFP cassette is duplicated. This places two RFP exons in a productive arrangement, and splicing enables the expression of wild-type RFP [20]. Unlike ISceI-induced LTGC, which is Rad51-dependent, Tus-Terinduced LTGC is Rad51-independent and is quantitatively increased in cells lacking BRCA1 [17]. We speculate that some Tus-Ter-induced LTGCs may be products of aberrant stalled fork restart. 3. Tandem duplication (TD): TDs are detected as GFPRFP+ products. They are aberrant repair outcomes that arise through a nonhomologous duplication of the RFP cassette [21]. TDs are detectable in response to a Tus-Ter RFB, but not in response to an I-SceI-induced DSB. Thus, TD formation appears to be specific to the stalled fork response. Mechanistically, Tus-Terinduced TDs arise by an aberrant fork restart mechanism, terminated by end joining. Because the break point of the TD is nonhomologous, the span of individual TDs, as well as RFP+ signal intensity, varies significantly between independent TD repair clones. TDs occur frequently in cancer genomes, with typical spans of >100 kb [28, 29]. Notably, breast and ovarian cancer genomes lacking the hereditary breast and ovarian cancer predisposition gene BRCA1 contain numerous short-span (~10 kb) TDs [21, 28–30]. BRCA1 mutant mES HR reporter cells produce Tus/Ter-induced TDs of similar size, recapitulating the TD phenotype of BRCA1 mutant human cancers [21]. While a low frequency of GFPRFP+ events is observed in response to an I-SceI-induced DSB, our unpublished structural analysis of these I-SceI-induced GFPRFP+ repair products shows that they are not nonhomologous TDs of the type detected at the Tus-Ter RFB. Here we present how to propagate, prepare, and transfect the 6Ter-HR reporter mES cells to reliably and accurately measure repair at the Tus-Ter RFB. We describe in detail how to plot, gate, and score repair by flow cytometry and how to properly quantify Tus-Ter-induced or I-SceI-induced STGC, LTGC, and TD repair frequencies.

2

Materials

2.1 Tissue Culture Media and Solutions

Prepare all solutions using deionized water and sterilize by filter sterilization or autoclaving unless otherwise specified. 1. Mouse embryonic stem cell medium: DMEM with L-glutamine, 4.5 g/L glucose and sodium pyruvate further supplemented to 15% serum (see Note 1), 20 mM HEPES pH 7.6,

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0.1 mM beta-mercaptoethanol, 500 U/mL rLIF, 1 MEM nonessential amino acids (NEAA), 2 mM L-glutamine, 100 U/ mL penicillin, 100 μg/mL streptomycin, and 0.001 M sodium pyruvate. To DMEM, add HEPES, beta-mercaptoethanol, rLIF, NEAA, and sodium pyruvate using aseptic technique in a sterile tissue culture hood and store at 4  C. Before first use, add additional supplements bringing serum to 15%, L-glutamine to 2 mM, penicillin to 100 U/mL, streptomycin to 100 μg/mL, and MycoZap prophylactic to 1. Store at 4  C. DMEM, Fisher Scientific. 0.1 M sodium pyruvate, Fisher Scientific 10,000 U/mL penicillin, 10,000 μg/mL streptomycin, Fisher Scientific 0.2 M L-glutamine, Cellgro 1 M HEPES pH 7.6, Fisher Scientific 100 NEAA (nonessential amino acids), Fisher Scientific ESGRO rLIF ten million units/mL, EMD Millipore. 10 MycoZap Prophylactic, Lonza. 2. MEF medium: DMEM with L-glutamine, 4.5 g/L glucose and sodium pyruvate further supplemented to 15% serum, 0.8 mM beta-mercaptoethanol, 2 mM L-glutamine, 100 U/mL penicillin, and 100 μg/mL streptomycin. To DMEM, add betamercaptoethanol, serum to 10%, L-glutamine to 2 mM, penicillin to 100 U/mL, and streptomycin to 100 μg/mL. Store at 4  C. 3. 10 PBS: 136 mM NaCl, 2.6 mM KCl, 10 mM Na2HPO4∙7H2O, and 1.7 mM KH2PO4. Weigh 80 g NaCl, 2 g KCl, 14.4 g Na2HPO4, and 2.4 g KH2PO4. To 900 mL deionized water and pH to 6.7 with 4 M HCl (when diluted to 1 PBS will be pH 7.4). Autoclave 30 min on liquid cycle. Store at RT. 4. 1 PBS: 13.6 mM NaCl, 0.26 mM KCl, 1 mM Na2HPO4∙7H2O, and 0.17 mM KH2PO4. Measure 50 mL 10 PBS and dilute with 450 mL deionized water. Autoclave 30 min on liquid cycle. Store at RT. 5. 1 PBS 0.2% gelatin: Measure 50 mL 10 PBS solution, add 1 g porcine gelatin, and dilute with 450 mL deionized water. Autoclave 30 min on liquid cycle. While media is still hot but not superheated or boiling, swirl gently to fully dissolve gelatin. Store at RT. 6. Trypsin/EDTA: 1 PBS, 0.25% trypsin, and 1 mM EDTA. Measure 100 mL 10 PBS solution and dilute with 800 mL deionized water. Add 2 mL sterile 0.5 M EDTA pH 8.0 and autoclave 30 min on liquid cycle. Allow the solution to cool to

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room temperature. Add 100 mL 2.5% trypsin (Fisher Scientific, cat #MT25054CI) using aseptic technique under sterile conditions. Aliquot to 50 mL conical tubes and store at 20  C. Thaw at 37  C prior to first use and store at 4  C until fully consumed. 7. DMEM/EDTA (2 mM EDTA): Add 0.5 M EDTA pH 8.0 to 2 m final concentration in DMEM. 2.2 Plastic Consumables

1. 24-well flat-bottom plate. 2. 6-well flat-bottom plate. 3. 96-well flat-bottom plate. 4. 10 cm tissue culture dish. 5. 5 mL sterile polystyrene round-bottom tube (12  75 mm).

2.3 Mouse Embryonic Stem Cell Transfection Reagents

1. Lipofectamine 2000, Life Technologies. 2. Gibco Opti-MEM (serum-free media), Life Technologies. 3. Endo-free Plasmid Maxiprep Kit, Qiagen. 4. Horizon Scientific SMARTpool.

(Dharmacon)

siRNA

On-Target

5. Hemocytometer. 6. 0.4% Trypan blue solution. 7. Beckman GS65KR, outfitted with a GM-3.8 swinging bucket rotor (204 mm max distance). 2.4 Flow Cytometry and Repair Quantitation

1. Beckman Coulter Cytoflex LX fitted with yellow (561 nm) and blue (488 nm) lasers (see Note 2). 2. Flowjo 10. 3. Microsoft Excel.

3

Methods Perform all steps using aseptic technique in a tissue culture hood unless otherwise specified. Carry out all tissue culture manipulations using reagents warmed to room temperature with the exception of the Lipofectamine 2000 (stored at 4  C with limited time at room temperature) and plasmid or siRNA aliquots (stored at 20  C and maintained on ice until used after thawing at room temperature).

3.1 Endotoxin-Free Plasmid Amplification and Preparation

1. Transform high-copy mammalian empty vector (EV), Tus, and I-SceI expression plasmids into competent dh5alpha bacteria, plate on LB media supplemented with appropriate selection agent, and incubate overnight (16–20 h) at 37  C.

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2. Pick 1-day-old colonies directly into 300 mL LB broth containing the appropriate selection agent and incubate the cultures overnight (16–20 h) at 37  C, 200 rpm orbital shaking (see Note 3). 3. Prepare endotoxin-free plasmids using Qiagen Endo-free Maxiprep or Megaprep kits as per manufacturer’s instructions (see Note 4). 4. Dilute purified EV/Tus/I-SceI expression plasmids to 0.5 μg/ μL in commercial, nuclease-free water, and aliquot to 100 μL in sterile 1.7 mL Eppendorf tubes to minimize freeze-thaw cycles. For GFP expression plasmid, dilute to 0.25 μg/μL. Store all plasmid aliquots at 20  C. 3.2 Thaw and Recover Mouse Embryonic stem (mES) Cell HR Reporter

1. Thaw mES cells onto six-well plates previously coated with “feeder” mouse embryonic fibroblasts (MEF) in fresh mES cell media (see Note 5). 2. Incubate the mES cells under standard tissue culture conditions (37  C, 5% CO2) to form small, light-colored, smooth, undifferentiated, refractile colonies after approximately 5 days growth time (see Note 6). 3. Aspirate mES cell media and wash each well with PBS. 4. Add 0.25 mL trypsin/EDTA and incubate for 1 minute at room temperature (see Note 7) followed by quenching the trypsin enzymatic activity with the addition of 0.75 mL (three volumes) mES cell media. 5. Disaggregate mES cell colonies by mechanical action: carefully pipette the contents of each well five times using a p1000 pipette (see Note 8). 6. Transfer the 1 mL mES cell suspension to a 15 mL conical tube and pellet cells, 1000 rpm/200  g, 4  C, 3–5 min. 7. Resuspend the cell pellet in mES cell media and return the cell suspension to the six-well plate in a final volume of 2 mL (see Notes 9 and 10).

3.3 Adapt and Maintain mES Cell Reporters in Culture

1. After sufficient proliferation of the reseeded culture, disaggregate the mES culture using a combination of trypsin digest and mechanical action (see Subheading 3.1, steps 3–6) and plate an appropriate dilution of each culture on fresh gelatinized plates (see Notes 11 and 12). 2. Return the culture(s) to grow under standard conditions allowing ~3 days for mES cell recovery and proliferation. 3. Maintain mES cells in culture by passaging cells every three days alternating between reseeding cells onto the currently used, conditioned plate or passaging the cells onto fresh, gelatinized media. For guidelines and best practices, see Notes 13–15.

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3.4 Prepare mES HR Reporter Cells for Transfection

1. Expand mES cell cultures appropriately for transfection (see Notes 16 and 17). 2. One day prior to transfection, reseed mES cell culture(s) to stimulate proliferation (see Notes 18 and 19). 3. On the day of transfection, harvest cells as normal and count viable cells using trypan blue and a hemocytometer (see Note 20). While performing the cell count, pellet the cells to remove the trypsin. 4. Resuspend the mES cells in mES cell media to a density of 0.8 M cells/mL (see Note 21). 5. Label an appropriate number of 24-well plates for transfections as shown in Fig. 2a (see Notes 22–24). Subsequent steps will follow the use of the single 24-well plate shown in Fig. 2a. 6. Cover the wells of all the 24-well plates with PBS/gelatin and incubate the plates for at least 5 min at room temperature to gelatinize (see Note 25). 7. Aspirate the PBS/gelatin and immediately plate 200 μL 0.8 M cells/mL mES cell suspension (160,000 cells) to each well using a p1000 pipette (see Note 26).

3.5 Prepare Transfection Mixes and Lipofection Reactions for mES Cell Transfection

1. Prepare four plasmid mixes and a single Lipofectamine 2000 mix as outlined in Fig. 2b in sterile 5 mL polystyrene tubes. Plasmid and Lipofectamine mixes to combine to produce a single lipofection reaction for transient transfection of a single well is listed below and should be scaled up appropriately (see Note 27). SeeNotes 28 and 29 for additional details regarding more complex mixes. 1 Plasmid mix (EV, Tus, I-SceI, GFP)

1 Lipofectamine mix

0.5 μg plasmid

1.2 μL Lipofectamine 2000

33 μL Opti-MEM

33 μL Opti-MEM

2. First prepare each plasmid mix (EV, Tus, I-SceI, and 10% GFP expression plasmid) in parallel and agitate each tube by gently flicking the tubes to mix (see Note 30 and 31). 3. Prepare the Lipofectamine mix after the plasmid mixes are ready and mix by capping and inverting the tube several times (see Note 32). 4. Incubate the fresh plasmid and Lipofectamine mixes at room temperature for 5 min. 5. Invert the Lipofectamine mix several times again before proceeding to setting up the lipofection reactions.

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Fig. 2 Twenty-four-well plate transfection format and transfection reaction summary. (a) To measure cellular repair response to replication fork stalling or to a site-specific DSB, four clones are transfected with empty vector (“EV”), Tus-expression plasmid (“Tus”), or I-SceI expression plasmid (“I-SceI”). In parallel, to accurately measure transfection efficiency, cells are co-transfected with plasmid mixes consisting of 90% EV and 10% GFP expression plasmid (“10 % GFP”) by mass. Limiting GFP expression plasmid provides more accurate measurement of transfection efficiency. Transfection efficiency is quantified by GFP+ frequency (see Notes 16 and 17). (b) Transfection mix setup for transfections shown in panel (a). Due to residual volumes during pipetting, both plasmid and Lipofectamine mixes and the final lipofection reactions are prepared in slight excess to what is required. Following this scheme consumes more reagent but ensures enough material is prepared for proper lipofection of all wells on the 24-well plate shown

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6. Transfer one equal volume of Lipofectamine mix to each plasmid mix to set up the lipofection reactions. Agitate each 5 mL reaction tube by gentle flicking or pipetting using a p1000 to mix (see Note 33). 7. Incubate the lipofection reactions at room temperature for at least 5 min (see Note 34). 3.6 Transiently Transfect mES Reporter Cells

1. Set up all 24-well plates so that the wells to receive EV or the GFP reactions are open—place the plate lid to cover the wells designated to receive Tus or I-SceI lipofection reactions to prevent GFP plasmid contamination of these wells. 2. Flick each lipofection mix gently before addition and transfect target wells by transferring 70 μL each lipofection reaction to each designated well. Begin by adding the EV control reaction to all appropriate wells, then add all GFP transfection efficiency reactions to appropriate wells, then add Tus, and finish with the addition of the I-SceI lipofection reaction mixes (see Note 35). 3. After the addition of the lipofection reactions, gently agitate the 24-well plate(s) being careful not to swirl the contents of the wells (see Note 36). 4. Place the 24-well transfection plate(s) in the tissue culture incubator for 6 h (see Note 37). 5. Gently add 1 mL mES cell media to each transfected well using a 25 mL pipette without the pipette tip meeting any individual wells contents (see Note 38). Return the plate(s) to the tissue culture incubator to proliferate overnight (16–24 h). 6. The following day, between 16 and 28 h after transfection, replace the media in each well with 1 mL fresh mES cell media and return the plates to the incubator for another two days (see Notes 39 and 40).

3.7 Reporter Cell Preparation and Analysis by Flow Cytometry

1. Remove mES cell media from each well for the entire plate by forcefully decanting the contents of the wells (see Note 41). 2. Wash each well with PBS: using a 25 mL pipette, add PBS to wash each well of residual media and serum and repeat the decanting process (see Note 41). 3. Using a 5 mL pipette, add two drops (~100 μL) 0.25% trypsin/ EDTA to each well and incubate at 37  C for 5 min. Firmly tap to agitate each plate to loosen and dislodge adherent cells from the plastic and to help break up any cell clumps (see Note 42). 4. Using a 5 mL pipette, add two drops (~100 μL) DMEM/ EDTA to each well and swirl the wells to mix. 5. Using nonfilter p1000 tips, pipette each well five times and transfer the cell suspension (~240 μL) to a single well of a 96-well plate and keep these plates at room temperature (see

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Note 43). Arrange all EV transfection samples in row A, Tus in rows B and C, I-SceI in rows D and E, and lastly GFP in row F thereby segregating each clone or condition to a single column on the 96-well plate (see Note 44). 6. Set up the Beckman Coulter Cytoflex LX cytometer configuration and gating to exclude small subcellular debris and any events containing more than a single cell. Exclude cellular debris after initially plotting FSC-A versus SSC-A. Further refine by excluding events produced by cell doublets after plotting FSC-width versus FSC-A. These exclusions will improve the accuracy of measurements of both the absolute raw values of repair events scored in real-time and calculated repair frequency for STGC, LTGC, and TD repair events (see Note 45 and gating provided in Fig. 3a). 7. For each independent 96-well plate, record event data for all EV samples first, followed by Tus, and finally I-SceI. Backflush 5 between 96-well plates and repeat this maneuver until all plates are finished (see Notes 46–49). 8. Record event data for all the GFP transfection wells for all plates to conclude the cytometer run. 9. Back flush the system several times and run 6–12 wells containing 0.2 mL PBS to clear the system of residual cells and debris. 10. Export the data as FCS files for analysis using Flowjo 10 or appropriate software. If 600,000 events were captured per sample well, each 96-well plate will produce 1–2 GB of data. 3.8 Flowjo Analysis and SCR Repair Quantitation

1. Import FCS files for analysis into a new Workspace using Flowjo 10. 2. Define two groups within the Flowjo Workbook: one group to quantify SCR and aberrant repair, and a second group to quantify transfection efficiency. Be sure to enable “synchronization” to ensure that any changes produced within any single scatter plot (axis range, gate position, etc.) is applied universally to all samples within the group. 3. Within Flowjo, restrict events for repair quantitation through successive rounds of hierarchical gating excluding events too small or too large to be solitary mES cells. Gate to exclude events such as cellular debris which are too small to be solitary, intact mES cells after plotting FSC-H versus SSC-H. Gate to exclude additional events too large to be mES cells after plotting FSC-A versus SSC-A. Lastly, gate to exclude cell doublets after plotting FSC-width versus FSC-H (see Fig. 4a for gating ancestry and Notes 50 and 51). Flowjo will provide raw repair frequencies (see Figs. 4b and 5a) which must be further manipulated to estimate the true repair frequencies (seestep 4 below).

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Fig. 3 Cytoflex LX flow cytometer gating setup for data acquisition. (a) Sample data shown (adapted from CytExpert software shell) is for a single representative technical replicate of BRCA1Δ/exon11 6Ter-HR reporter cells co-transfected with wild-type Tus and short interfering (si) RNA against siBRCA1. Cytoflex LX forward scatter (FSC) and side scatter (SSC) voltages are adjusted, and hierarchical gating is applied to analyze events representative of single cells. Left panel: P1 gating (red) applied to the FSC-A versus SSC-A plot excludes small events such as autofluorescent cellular debris or dead cells. Middle panel: P2 gating (blue) applied to the FSC-width versus FSC-H plot excludes events too large or complex to be produced by a single cell including cell doublets. This rough gating scheme will exclude events detected outside the profile generated by a typical single, solitary mES cell which may erroneously score as a repair event (i.e., for a droplet containing two cells, one GFP+RFP and the other GFPRFP+, this excluded event would score as a GFP+RFP+ LTGC repair outcome). Right panel: FITC-A versus mCHERRY-A plot depicting quadrant gating used to roughly identify STGC, LTGC, and TD events highlighted in green, orange, and red, respectively. (b) Events and raw frequency of repair at the conclusion of representative sample run

4. To properly quantify repair frequencies first subtract background event frequency displayed in EV samples for STGC, LTGC, and RFP+ repair events from the raw values found for Tus or I-SceI transfected sample. Then normalize these values against transfection efficiency to compare across conditions within a single experiment (see Fig. 5b) and also between biological replicates produced from independent experiments.

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Fig. 4 Flowjo gating and raw repair frequency determination. (a) The combined data collected from four independent experiment duplicate samples for BRCA1 mutant reporter cells treated with siBRCA1 and transiently co-transfected with Tus-expression vector is shown. Left panels: biexponential mCherry-A versus FITC-A pseudo-color scatter plot (top) and contour plot with outliers (bottom). Scatter plot and contour plots are used to refine STGC, LTGC, and TD gate placement to properly exclude uncolored events. Right panels: gating ancestry excluding cellular debris/dead cells (FSC-H vs. SSC-H plot), larger events (FSC-A vs. SSC-A plot), and events generated by doublets or small cell clumps containing more than a solitary mES cell (FSC-Width vs. FSC-H plot). Plots generated using Flowjo 10.6. (b) Raw numbers and frequencies calculated for Tus-induced repair events from gating applied to the scatter plot shown in panel (a)

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Fig. 5 Normalized STGC, LTGC, and TD repair frequency calculation. (a) Left: scatter plots corresponding to combined data from BRCA1 mutant reporter cells treated with siBRCA1 and co-transfected with either empty vector (“EV”) or Tus-expression vector (“Tus”). Scatter plot displaying Tus-induced repair identical to data is shown in Fig. 4. Gated STGC, LTGC, and TD repair events are highlighted in green, orange, and red, respectively. Inset values indicate raw STGC, LTGC, and TD repair frequencies measured. Right: the pseudo-color plot indicates transfection efficiency as measured after transient transfection of GFP expression plasmid (at 1:10 dilution with EV). Plots generated using Flowjo 10.6. (b) To correct for background frequencies of repair products, the observed raw induced frequencies are adjusted by subtracting raw background frequencies measured for empty vector (“EV % ”) from Tus-induced repair frequencies (“Tus %”). To correct for transfection efficiency, the background-adjusted raw repair frequencies are normalized against the measured transfection efficiency (e.g., in this figure, the background-adjusted Tus-induced TD frequency of 0.009485% is divided by 0.85 to calculate the normalized TD frequency of 0.01116% per transfected cell)

Often between five and twelve independent experiments must be performed and compiled to generate datasets suitable for publication.

4

Notes 1. Six to twelve lots of serum from multiple vendors should be tested to identify batches with optimal mES cell proliferation and plating efficiency with minimal differentiation and cell death. Growth assays are typically conducted over a four-week

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period allowing several passages for initial mES cell adaptation to the new sera. We find no correlation between quoted cost per bottle and performance. 2. Any flow cytometer allowing independent detection of GFP and RFP fluorescence outfitted with both 461 nm (yellow) and 488 nm (blue) laser coupled to BP 610/20 or 525/40 nm, respectively, may be used. We use either a Beckman Coulter Cytoflex LX running CytExpert 2.3.0.84 or a Becton Dickinson 5 Laser LSR II running FACSdiva to detect RFP(3.1) and GFP fluorescence. 3. Plasmid yield may be improved by inoculating a single colony directly into a large volume of LB broth without the use of a starter culture. Briefly freeze and then fully thaw the bacteria pellet prior to plasmid preparation to improve plasmid yield. 4. Plasmid purity is an important determinant of transfected mES cell survival and transfection efficiency—both are important factors for reliable and reproducible quantitation of repair frequencies for which both the cell number analyzed and the absolute number of repair outcomes scored during the flow cytometry run are critical. We observe increased cytotoxicity, decreased proliferation, and decreased transfection efficiency when comparing transfections performed using nonendotoxin-free plasmid preparations to transfections performed using endotoxin-free preparations. 5. mES cells grow extremely poorly if thawed directly onto plastic, displaying low plating efficiency and significant cell death. Initially, we thaw mES cells into a single well of a six-well plate pre-coated with mouse embryonic fibroblast (MEF) “feeder” cells that have been mitotically inactivated by lethal irradiation. MEFs provide both matrix support for mES cell attachment and secretion of factors that enhance cell survival and pluripotency. Feeders or nonirradiated MEFs may be purchased from commercial suppliers. mES cells must be adapted to enable them to proliferate under MEF-free culture conditions on gelatin prior to transfection for HR analysis. The reason for this is that MEFs, if present in the HR assay, will capture a significant proportion of transfected nucleic acid (plasmid and/or siRNA), thereby interfering with the accurate measurement of repair responses in the mES reporter cells. 6. Normally, mES cells do not grow as a monolayer, but form discrete three-dimensional colonies when grown under standard culture conditions. As mES cell colonies begin to overgrow, cells at the colony interior begin to differentiate—under brightfield visualization, large colony interiors take on a darkened, mottled appearance. It is important to disaggregate colonies by trypsin digestion combined with and mechanical

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shearing (pipetting using a p1000 regular-bore tip) and passaging before they show signs of differentiation (seesteps 3–5 for details). 7. Do not overly trypsin digest mES cell cultures. If cells slough off the plate surface after tilting the plate without mechanical agitation, cells have been overly digested. The trypsin must be removed under most circumstances through neutralization with serum, followed by washing and centrifugation (seestep 6). 8. Pipette the medium sufficiently to dislodge the mES cell colonies from the plate surface and then pipette several more times to efficiently disaggregate all clumps to single cells. MEF feeders will also be dislodged and suspended with the mES cell suspension. 9. Passaging the mixed culture will lead to a loss of feeders through a failure of MEFs to adhere after replating and through simple dilution. First, reseed mES reporter cells after sufficient growth (see Note 6). Initially, reseed cultures back to the source feeder well used for thawing: for wells containing several hundred to thousands of colonies, reseed the culture 1:5, replating only 20% of the culture; for wells containing less than 100 colonies, replate the entire disaggregated culture. 10. Excess cells may be frozen in 90% serum, 10% DMSO at 80  C in the short term and transferred to liquid nitrogen cooled cyto-storage for the long term. 11. After the first reseed, cultures must be passaged to fresh gelatinized media and may not be reseeded a second time onto the source well. 12. Adaptation to gelatin is a shock to mES cells, which proliferate best on feeders. Often a moderate number of cells die during the initial, early passages to and on gelatinized plates lacking feeder MEFs. For healthy, highly proliferative cultures with plate coverage approaching confluency, passage cells 1:5, plating ~20% of the culture. For cultures displaying 60%, 50%, and ~30% coverage, passage 1:4, 1:3, and 1:2, respectively. For cultures that are significantly undergrown, displaying less than 30% coverage after three days growth, composed of isolated colonies, passage all or the majority of the population to a new well. 13. After adaption to proliferation on gelatinized plates, mES cell populations often look best (more smooth, light-colored, refractile, uniformly shaped colonies) after reseeding to conditioned plastic (i.e., conditioned by previous use for culturing ES cells). mES cells double approximately every 14 h, requiring frequent passaging multiple times each week to prevent overgrowth (see Note 14). Typically, mES cells in a six-well plate format are passaged 1:10 to achieve a similar

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confluency (cell surface density) after 48 h proliferation and may thereby be passaged Monday to Wednesday to Friday to Monday. 14. Care should be taken to prevent mES cell cultures from reaching or exceeding 100% confluency. Overgrowth may result in significant cell death and potential loss of the culture. Frequently allowing mES reporter cells to overgrow will cause reporter repair characteristics to change over time. Passaging cells 1:5 will achieve a similar confluency within 24 h. Passaging cells 1:10 will achieve a similar cell density within 48 h, and passaging 1:15 will allow cells to proliferate without attention over a 72-h period. mES cell plating efficiency is highly sensitive to cell number with cultures displaying approximately 50% plating efficiency when plating a limited number (hundreds) of cells. For routine mES culture maintenance, do not split cells more aggressively than 1:15. 15. Passaging mES cells, whether reseeding or splitting cells to fresh gelatinized plates, most often requires pelleting of cells to remove the trypsin. However, routine passaging of mES cells in the six-well plate format may be achieved without trypsin removal if wells are more than 75% confluent: effectively passage cells 1:15 by transferring two drops (60%) well from the six-well plate to the larger dish is effectively a 1:5 dilution. A single fairly (80–90%) confluent well of a six-well plate could also be split between two 10 cm dishes for amplification over this same time period. Under normal growth conditions, such a maneuver may be performed over a weekend. For individual clone repair assays, 1–2 M cells are needed, and a single well of a six-well plate will be sufficient. To survey a large number of test conditions such as a panel of siRNAs, 4–40 M cells are required, and individual clone mES cell cultures must be expanded to 1 or more 10 cm dishes. 17. For cells grown in the 10 cm dish format, wash with PBS and add 1 mL trypsin/EDTA to digest. Rock the plate to ensure uniform trypsin coverage and incubate the plate 30–60 s at

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room temperature. Add 2 mL (2 V) mES cell media to quench the trypsin activity. Using a p1000, forcibly dislodge the cells off the surface of the plate and pipette five to ten times to disaggregate any cell clumps. Pellet cells to remove the trypsin and resuspend the culture as normal. 18. Generally, the higher the culture confluency and plate coverage the day of the reseed, the better conditioned the plastic will be and the better the reseeded mES cells will appear the following day. If cells are counted, plating 3–4 M cells will result in a healthy culture of 50–65% confluency consisting of highly refractile, smooth colonies. If counting is not performed and reseeding is based on culture confluency, for a confluent dish, reseed 1:5 in the morning or 1:6 if reseeded in the afternoon (assuming transfection will be performed the following morning). For ideal cultures displaying 80–90% confluency, reseed approximately 1/4 of cells. For cultures displaying 40–60% confluency, reseed approximately 1/3–½ of cells. For plates displaying a cell density of 30% or less, reseed at least ¾ of the culture. 19. Reseed the cells so as to reach 80–90% confluency the following day (16–24 h). This strategy permits the harvest of the maximum number of proliferating cells without incurring unnecessary cell stress due to culture overgrowth or undergrowth due to overly sparse cell growth and minimal plate conditioning. 20. To minimize cellular exposure to trypsin, pellet the cells while performing the cell count. To count the cells, dilute 10 μL of cell suspension into 90 μL trypan blue. The cell concentration is calculated by multiplying the number of cells counted by 10,000 and the dilution factor of 10 (i.e., a count of 34 indicates a concentration of 3.4 M cells/mL). 21. Typically for transfection, 200 μL cells at 0.8 M cells/mL are transferred to each well. However, transfection of mES cells in the 24-well plate format is able to tolerate minor changes to mass of nucleic acid(s) used, cell density and absolute number, and transfection volume. The transfection conditions described may accommodate cell densities up to 2 M cells/ mL with no reduction in the transfection efficiency or significant changes to the repair frequencies measured. Transfection efficiency may be increased by reducing the cell suspension volume slightly while maintaining the total cell number deposited into each well (i.e., 175 μL cells with a cell density of 0.9 M cells/mL). Transfections performed with the intention of harvesting the cells within 24–48 h often require a greater number of transfected cells up-front, and increasing the cell density will limit the number of wells or the number of plates to be processed.

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22. For functional assays, mES cells are transfected in suspension in a 24-well plate format. Tissue culture plate surface area influences transfection efficiency. Larger plate formats with greater surface area such as six-well plates or 10 cm dishes require more cells, transfection reagent, and nucleic acid while displaying reduced transfection efficiencies compared with transfections performed in the 24-well plate format. 23. To measure transfection efficiency, co-transfect cells in parallel with a limiting amount of GFP expression plasmid combined with empty vector control plasmid in the plasmid mix. The frequency of GFP+ cells measured 72 h after transfection is used to quantify transfection efficiency and used to normalize functional repair data between independent experiments. Transfection efficiencies greater than 65% are optimal while experiments displaying a transfection efficiency below 40% are thrown out—often these low transfection efficiency experiments display divergent, atypical functional results when compared with functional data generated from high transfection efficiency experiments. 24. Assessing any condition in triplicate is best. However, we find very little variation in repair frequencies observed for cells transfected with empty vector (EV) or significant variation in transfection efficiencies measured by transfection with limiting GFP expression plasmid. In an effort to minimize costs related to reagent consumption, time and labor, and flow cytometer use, transfections are restricted to include a single well for measuring transfection efficiency, and five wells for measuring repair—a single well for the EV control, two wells for Tus, and two wells for I-SceI-induced repair. The 24-well plate format shown in Fig. 2a allows the assessment of four independent clones or the clonal response to four different treatment groups such as siRNA treatment or supplemental plasmid co-transfection. This strategy is especially important when surveying a large number of clones or treatments and allows for a greater number of biological repeats to be performed with similar consumable use. 25. PBS/gelatin may be added in advance and gelatinizing plates placed in the tissue culture hood for up to several hours. 26. Minimize the production of bubbles when transferring cells to the 24-well plate by performing the following: (a) after taking up 200 μL cell suspension, rest the side of the p1000 tip against the lip of the well and touch the tip of the p1000 to the bottom edge of the well; (b) gently expel the cell suspension pressing the pipette to “first stop” and do not continue to “second stop”; (c) without relaxing this grip on the pipette, return the tip to the cell suspension to take up another 200 μL cell

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suspension by releasing the pipette from “first stop.” Repeat this process until all aspirated wells contain cells. Do not allow gelatinized wells to dry. Typically, cell transfer to two 24-well plates may be completed before gelatinized wells dry out. 27. Because pipetting is associated with a residual dead volume, prepare extra volumes for each plasmid mix (i.e., for the transfection of four clones as shown in Fig. 2, four wells of EV and eight wells of Tus are required, therefore prepare 5 and 10 mixes, respectively (see Fig. 3)). 28. The basic expression plasmid transfection mix is shown. Co-transfection of EV/Tus/I-SceI/GFP plasmids with siRNA and or additional expression plasmids is routinely performed. For transient transfection of expression plasmids, the total mass of plasmid used per lipofection reaction should not deviate significantly from 0.5 μg per reaction. Perform titration experiments to determine the optimal balance of EV/Tus/ISceI expression plasmid to supplemental expression plasmid for the gene of interest maintaining the total mass of plasmid for all test reactions. Between 50 and 150 ng expression plasmid is sufficient in most maneuvers. EV/Tus/I-SceI expression plasmid should not be reduced below 0.25 μg per transfection reaction. GFP expression plasmid should be maintained at 50 ng per reaction regardless of other components in the lipofection reaction. Lipofectamine concentration remains unchanged. Examples of these strategies are published [17, 21, 24]. 29. For siRNA, no more than 20 pmol On-Target SMARTpool is added per reaction along with 0.35 μg EV/Tus/I-SceI expression plasmid. Ten pmol siRNA produces similar levels of mRNA or protein depletion to transfections performed using 20 pmol per reaction. When depleting multiple targets by siRNA treatment, keep any single siRNA dose constant by supplementing any single siRNA with control siRNA (i.e., for assessment of codepletion of BRCA1 and FANCM, compare reactions containing 10 pmol siBRCA1 and 10 pmol siFANCM to reactions containing 10 pmol siBRCA1 or 10 pmol siFANCM supplemented with 10 pmol siLUC, and with reactions containing 20 pmol siLUC control). 30. For the measurement of transfection efficiency, combine 0.45 μg EV and 50 ng GFP expression plasmid per reaction (90% EV, 10% GFP plasmid). Transfection efficiency varies very little between mES cell populations transfected with EV, Tus, or I-SceI expression plasmids. There is no need to measure transfection efficiency for each expression plasmid. 31. When performing transfections incorporating supplemental siRNA or expression plasmids, prepare a single EV/Tus/I-

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SceI expression plasmid master mix in excess of what is needed for any single mix containing independent siRNAs. Calculate the master mix volume needed by considering making each single mix in slight excess. 32. Prepare a single Lipofectamine master mix to ensure that all transfected cells receive a nearly identical amount of the lipofection agent. To determine the minimum Lipofectamine mix to prepare, total the number of mixes assembled for EV, Tus, I-SceI, and GFP plasmids, and prepare extra to account for dead volume associated with pipetting. 33. Change tips between each transfer of the Lipofectamine mix to each plasmid mix to prevent contamination of any lipofection reaction with the GFP plasmid mix. Do not vortex lipofection reactions to mix. 34. Lipofection reactions should be added immediately to target cells after 5 min incubation at room temperature. However, this transfection protocol tolerates a delay of up to 25 min after combining the plasmid and Lipofectamine preparations before addition to cells. This flexibility in timing accommodates up to two dozen 24-well plates to undergo lipofection during any single round of transient transfection. 35. Use sterile nonfilter p200 tips for the transfection maneuver. After taking up 70 μL lipofection reaction, and while resting the side of the p200 tip on the lip of the well without dipping the tip into the cell suspension, expel the transfection mix into each well by firmly pressing the pipette fully to “second stop.” Release the pipette to return to “first stop” and immediately take up another aliquot of the current lipofection reaction. Repeat this process for all EV wells and then for all GFP wells changing tips between different lipofection reactions. Be sure to change tips before proceeding to the Tus reactions and again for the I-SceI reactions. 36. A circular mixing motion will swirl the contents of the wells and will concentrate cells toward the periphery of each well. This consequence may limit transfection efficiency. Rock the contents of the wells, scraping the 24-well plates strictly forward and backward followed by side-to-side across the surface of the tissue culture incubator. 37. Six hours incubation for transfection appears optimal. However, cells may be fed earlier if required with 4.5 h incubation time being sufficient. In unpublished work, we observe no significant differences in STGC or LTGC frequency between transfected cells held for 2, 3, 4, 5, or 6 h prior addition of 1 mL mES cell media (seestep 5). mES cells take about 4 h to adhere strongly to gelatinized plastic. Feeding wells earlier than 4 h risks dislodging cells and reducing transfected cell survival.

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38. Long-term exposure to cationic lipid transfection agent is toxic to mES cells. Dilution of the lipofection reaction in the 24-well plate format (200 μL cells, 70 μL lipofection containing 1.2 μL Lipofectamine 2000) with 1 mL mES cell media effectively limits this toxicity. 39. Use a single Pasteur pipette to aspirate up to three 24-well plates at any one time. While to risk of cross contamination is never eliminated, there appears no need to change tips between any set of conditions including the GFP plasmid transfected wells. 40. For assessment of siRNA-mediated mRNA depletion or protein accumulation after transient transfection, maximal changes to mRNA or protein levels are observed approximately 48 h after transfection. A minimum of two wells for RNA or four wells for protein isolation should be pooled and processed accordingly. 41. Plates may be processed under nonsterile conditions. Forcefully decant the contents of the wells into a sink. There is no need to aspirate the media or the PBS wash by vacuum since cells are strictly adherent, lipofection reactions harboring plasmids have long been removed, and flow cytometry is normally not performed under sterile conditions. 42. Five minutes of trypsinization is more than adequate to loosen transfected cells. To agitate the contents of the wells, firmly tap one side of the 24-well plate. Cells should slough off the well surface immediately upon agitation and clumps easily be broken up by gentle agitation. 43. Cells should remain viable and not irreversibly clump for up to 6 h at room temperature suspended in the trypsin/EDTA, DMEM/EDTA mix. Cells in suspension will settle over time requiring resuspension. For this, we recommend using a multichannel pipette capable of taking up 200 μL volume. Placing plates on ice is not recommended and actually accelerates cell clumping. Fixing cells will decrease or destroy GFP and RFP (3.1) activity. Storing prepared 96-well plates at 4  C is not ideal and doing so will reduce the number of events captured. 44. This plate configuration will allow three 24-well plates to be well organized in a 96-well plate format saving rows G and H for PBS, water, extra samples for cytometer setup, cleaning agents, etc. Additionally, consistency in 96-well plate setup will allow for easier data analysis. 45. When setting up the Cytoflex LX, adjust FSC and SSC voltages to keep mES cells in range of detection. mES cells are quite small, just 7–17 μm in diameter [31], and as a consequence, the FSC and SSC voltages required may be far lower than traditionally expected for mammalian cell flow cytometry. Rough

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gating to exclude either debris or cell doublets is adequate at this stage with more refined gating performed during Flowjo analysis (see Subheading 3.8). Plot mCherry-A versus FITC-A measuring RFP(3.1) and GFP fluorescence, respectively, and use a quadrant gate to get a rough real-time estimate of raw STGC, LTGC, and GFPRFP+ numbers and frequency. Adjust 561 and 488 nm laser voltages to separate STGC events (GFP+RFP), LTGC events (GFP+RFP+), and TD events (GFPRFP+) from the main uncolored population by four to five decades (logs). A biexponential plot is shown in Fig. 4, which improves the presentation of RFP and GFP plots during the cytometer run and for analysis. 46. Set up acquisition settings on the Cytoflex LX with upper limits both on the number of events to acquire and maximum recording time for each sample—600,000 events or 1 min 20 s of recording time for any EV, Tus, and I-SceI well. For GFP transfection wells, record up to 20,000 events or for up to 20 s. Set a 5-s back flush and 3-s mix for each EV, Tus, and I-SceI well. These settings ensure that each 96-well plate run populated with samples from three 24-well plates will take approximately 2 h to complete with minimal carryover between wells. 47. Perform a deep clean cycle and several prime cycles followed by running several wells of PBS or sterile water to ensure the cytometer fluidics are working properly and lines sufficiently free of debris, air bubbles, or cells from previous cytometer runs. It is important to thoroughly clean the lines since repair events, especially TDs, are exceptionally rare even under the most ideal conditions (0.05%). 48. Acquire events for samples harboring the lowest frequency of repair events first (i.e., EV followed by Tus and then I-SceI). For samples expected to display abnormally high frequencies of HR (such as I-SceI-induced HR frequencies observed in Xrcc4 mutants [24] or aberrant repair events such as tandem duplications in BRCA1 mutant cells challenged with stalled fork repair when treated with siFANCM [21]), it is best to record data from these wells last or to manually back flush the system after recording to prevent carryover to immediate subsequent samples awaiting acquisition. Similarly, do not measure GFP transfection efficiency wells until all EV/Tus/I-SceI wells on all 96-well plates are recorded. 49. mES cells will settle over an hour’s time. Remove the plate from the cytometer after the EV and Tus sample wells are recorded and, using a multichannel pipette, resuspend I-SceI and GFP transfection wells (see Note 43). 50. Flowjo will display both raw numbers of STGC, LTGC, and RFP+ events, and also raw frequency of each category of repair

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within the sample population. We find that 300,000 events per sample are the minimum event number required to produce consistent data with event totals of 600,000 or more to be optimal. 51. Sample STGC event count totals in the 100 s and LTGC event count totals of several dozen appear the minimum required to generate consistent data. RFP+ events, defined as TDs when induced by Tus, often occur at a frequency of less than 1 in 100,000 unless induced by the functional loss of BRCA1, which increases the observed TD frequency ten- to 20-fold to ~15 events per 100,000. The majority of RFP+ events induced by I-SceI are homologous tandem duplications for which the interior of the reporter has been duplicated and the concatenated cassette has three copies of GFP, all nonfunctional with the central GFP allele maintaining the 6Ter-I-SceI sequence. References 1. Ciccia A, Elledge SJ (2010) The DNA damage response: making it safe to play with knives. Mol Cell 40(2):179–204. https://doi.org/ 10.1016/j.molcel.2010.09.019. S1097-2765 (10)00747-1 [pii]. PubMed PMID: 20965415; PubMed Central PMCID: PMC2988877 2. Zeman MK, Cimprich KA (2014) Causes and consequences of replication stress. Nat Cell Biol 16(1):2–9. https://doi.org/10.1038/ ncb2897 3. Bidnenko V, Ehrlich SD, Michel B (2002) Replication fork collapse at replication terminator sequences. EMBO J 21(14):3898–3907 4. Michel B (2000) Replication fork arrest and DNA recombination. Trends Biochem Sci 25 (4):173–178 5. Lambert S, Mizuno K, Blaisonneau J, Martineau S, Chanet R, Freon K et al (2010) Homologous recombination restarts blocked replication forks at the expense of genome rearrangements by template exchange. Mol Cell 39 (3):346–359. https://doi.org/10.1016/j. molcel.2010.07.015 6. Lambert S, Watson A, Sheedy DM, Martin B, Carr AM (2005) Gross chromosomal rearrangements and elevated recombination at an inducible site-specific replication fork barrier. Cell 121(5):689–702 7. Ahn JS, Osman F, Whitby MC (2005) Replication fork blockage by RTS1 at an ectopic site promotes recombination in fission yeast. EMBO J 24(11):2011–2023 8. Jalan M, Oehler J, Morrow CA, Osman F, Whitby MC (2019) Factors affecting template switch recombination associated with restarted

DNA replication. eLife 8. https://doi.org/10. 7554/eLife.41697. PubMed PMID: 30667359; PubMed Central PMCID: PMCPMC6358216 9. Nguyen MO, Jalan M, Morrow CA, Osman F, Whitby MC (2015) Recombination occurs within minutes of replication blockage by RTS1 producing restarted forks that are prone to collapse. eLife 4:e04539. https://doi.org/ 10.7554/eLife.04539. PubMed PMID: 25806683; PubMed Central PMCID: PMCPMC4407270 10. Larsen NB, Hickson ID, Mankouri HW (2014) Tus-Ter as a tool to study site-specific DNA replication perturbation in eukaryotes. Cell Cycle 13(19):2994–2998. https://doi. org/10.4161/15384101.2014.958912. PubMed PMID: 25486560; PubMed Central PMCID: PMCPMC4614373 11. Larsen NB, Liberti SE, Vogel I, Jorgensen SW, Hickson ID, Mankouri HW (2017) Stalled replication forks generate a distinct mutational signature in yeast. Proc Natl Acad Sci U S A 114(36):9665–9670. https://doi.org/10. 1073/pnas.1706640114. PubMed PMID: 28827358; PubMed Central PMCID: PMCPMC5594675 12. Larsen NB, Sass E, Suski C, Mankouri HW, Hickson ID (2014) The Escherichia coli Tus-Ter replication fork barrier causes sitespecific DNA replication perturbation in yeast. Nat Commun 5:3574. https://doi.org/10. 1038/ncomms4574 13. Berghuis BA, Dulin D, Xu ZQ, van Laar T, Cross B, Janissen R et al (2015) Strand separation establishes a sustained lock at the Tus-Ter replication fork barrier. Nat Chem Biol 11

HR at Stalled Mammalian Replication Forks (8):579–585. https://doi.org/10.1038/ nchembio.1857 14. Elshenawy MM, Jergic S, Xu ZQ, Sobhy MA, Takahashi M, Oakley AJ et al (2015) Replisome speed determines the efficiency of the Tus-Ter replication termination barrier. Nature 525 (7569):394–398. https://doi.org/10.1038/ nature14866 15. Mulcair MD, Schaeffer PM, Oakley AJ, Cross HF, Neylon C, Hill TM et al (2006) A molecular mousetrap determines polarity of termination of DNA replication in E. coli. Cell 125 (7):1309–1319 16. Pandey M, Elshenawy MM, Jergic S, Takahashi M, Dixon NE, Hamdan SM et al (2015) Two mechanisms coordinate replication termination by the Escherichia coli Tus-Ter complex. Nucleic Acids Res 43 (12):5924–5935. https://doi.org/10.1093/ nar/gkv527. PubMed PMID: 26007657; PubMed Central PMCID: PMCPMC4499146 17. Willis NA, Chandramouly G, Huang B, Kwok A, Follonier C, Deng C et al (2014) BRCA1 controls homologous recombination at Tus/Ter-stalled mammalian replication forks. Nature 510(7506):556–559. https:// doi.org/10.1038/nature13295. PubMed PMID: 24776801; PubMed Central PMCID: PMC4118467 18. Jasin M (1996) Genetic manipulation of genomes with rare-cutting endonucleases. Trends Genet 12(6):224–228 19. Shaner NC, Campbell RE, Steinbach PA, Giepmans BN, Palmer AE, Tsien RY (2004) Improved monomeric red, orange and yellow fluorescent proteins derived from Discosoma sp. red fluorescent protein. Nat Biotechnol 22 (12):1567–1572 20. Chandramouly G, Kwok A, Huang B, Willis NA, Xie A, Scully R (2013) BRCA1 and CtIP suppress long-tract gene conversion between sister chromatids. Nat Commun 4:2404. https://doi.org/10.1038/ncomms3404. ncomms3404 [pii]. PubMed PMID: 23994874; PubMed Central PMCID: PMC3838905 21. Willis NA, Frock RL, Menghi F, Duffey EE, Panday A, Camacho V et al (2017) Mechanism of tandem duplication formation in BRCA1mutant cells. Nature 551(7682):590–595. https://doi.org/10.1038/nature24477. PubMed PMID: 29168504; PubMed Central PMCID: PMCPMC5728692 22. Paques F, Haber JE (1999) Multiple pathways of recombination induced by double-strand breaks in Saccharomyces cerevisiae. Microbiol Mol Biol Rev 63(2):349–404

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23. Scully R, Panday A, Elango R, Willis NA (2019) DNA double-strand break repairpathway choice in somatic mammalian cells. Nat Rev Mol Cell Biol. https://doi.org/10. 1038/s41580-019-0152-0 24. Willis NA, Panday A, Duffey EE, Scully R (2018) Rad51 recruitment and exclusion of non-homologous end joining during homologous recombination at a Tus/Ter mammalian replication fork barrier. PLoS Genet 14(7): e1007486. https://doi.org/10.1371/journal. pgen.1007486. PubMed PMID: 30024881; PubMed Central PMCID: PMCPMC6067765 25. Johnson RD, Jasin M (2000) Sister chromatid gene conversion is a prominent double-strand break repair pathway in mammalian cells. EMBO J 19(13):3398–3407 26. Nagaraju G, Odate S, Xie A, Scully R (2006) Differential regulation of short- and long-tract gene conversion between sister chromatids by Rad51C. Mol Cell Biol 26(21):8075–8086. PubMed PMID: 16954385; PubMed Central PMCID: PMC1636746 27. Puget N, Knowlton M, Scully R (2005) Molecular analysis of sister chromatid recombination in mammalian cells. DNA Repair (Amst) 4 (2):149–161. PubMed PMID: 15590323; PubMed Central PMCID: PMC2967438 28. Menghi F, Barthel FP, Yadav V, Tang M, Ji B, Tang Z et al (2018) The tandem duplicator phenotype is a prevalent genome-wide cancer configuration driven by distinct gene mutations. Cancer Cell 34(2):197–210 e5. https://doi.org/10.1016/j.ccell.2018.06.008 29. Menghi F, Inaki K, Woo X, Kumar PA, Grzeda KR, Malhotra A et al (2016) The tandem duplicator phenotype as a distinct genomic configuration in cancer. Proc Natl Acad Sci U S A 113(17):E2373–E2382. https://doi.org/ 10.1073/pnas.1520010113. PubMed PMID: 27071093; PubMed Central PMCID: PMCPMC4855596 30. Nik-Zainal S, Davies H, Staaf J, Ramakrishna M, Glodzik D, Zou X et al (2016) Landscape of somatic mutations in 560 breast cancer whole-genome sequences. Nature 534(7605):47–54. https://doi.org/ 10.1038/nature17676. PubMed PMID: 27135926; PubMed Central PMCID: PMCPMC4910866 31. Pillarisetti A, Ladjal H, Ferreira A, Keefer C, Desai JP (2009) Mechanical characterization of mouse embryonic stem cells. Conf Proc IEEE Eng Med Biol Soc 2009:1176–1179. https:// doi.org/10.1109/IEMBS.2009.5333954

Chapter 24 Super-Resolution Imaging of Homologous Recombination Repair at Collapsed Replication Forks Donna R. Whelan and Eli Rothenberg Abstract Single-molecule super-resolution microscopy (SRM) combines single-molecule detection with spatial resolutions tenfold improved over conventional confocal microscopy. These two key advantages make it possible to visualize individual DNA replication and damage events within the cellular context of fixed cells. This in turn engenders the ability to decipher variations between individual replicative and damage species within a single nucleus, elucidating different subpopulations of stress and repair events. Here, we describe the protocol for combining SRM with novel labeling and damage assays to characterize DNA double-strand break (DSB) induction at stressed replication forks (RFs) and subsequent repair by homologous recombination (HR). These assays enable spatiotemporal mapping of DNA damage response and repair proteins to establish their in vivo function and interactions, as well as detailed characterization of specific dysfunctions in HR caused by drugs or mutations of interest. Key words Homologous recombination, Super resolution, DNA damage response, DNA doublestrand break, DNA replication

1

Introduction Single-molecule super-resolution microscopy (SRM) relies on the temporal separation of single-molecule fluorescence emissions such that at any one time during data acquisition only one emitter is fluorescing within a diffraction limited area. This temporal separation of individual emitters is achieved by photochemical or photophysical manipulation of the fluorophore to achieve a metastable dark state. The approach described here uses the stochastic optical reconstruction microscopy (STORM) SRM variant which employs high laser powers, a thiol-containing buffer and oxygen removal in order to photophysically convert the vast majority of organic fluorophore labels present within a sample into a reduced nonfluorescent state [1]. Thousands to millions of stochastically switched individual fluorophore emissions are imaged across thousands of frames to produce the raw SRM data. Subsequently, each individual

Andre´s Aguilera and Aura Carreira (eds.), Homologous Recombination: Methods and Protocols, Methods in Molecular Biology, vol. 2153, https://doi.org/10.1007/978-1-0716-0644-5_24, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 An overview of multicolor single-molecule SRM. (a) A diffraction limited micrograph of a section of cell nucleus fluorescently labeled for nascent DNA (naDNA) using EdU incorporation and detection via click chemistry and Alexa Fluor 647, and immunolabeled for DNA damage (targeting the histone modification γH2A.X with Alexa Fluor 568) and replication (targeting the replication processivity factor PCNA with Alexa Fluor 488). (b) Sequential single-molecule imaging in each of the three channels, using high laser flux and a reducing buffer to achieve spatially isolated individual fluorophore emission patterns [i.e., point spread functions (PSFs)]. (c) Mathematical fitting of each detected PSF generates a coordinate list of fluorescent molecule locations which are then registered using a predetermined elastic transformation to correct for chromatic aberrations. (d) The super-resolution image can then be generated by mapping each molecular coordinate in space, taking into account localization precision and label offsets. The image shows the same area and labels as in (a)

emission pattern is approximately fit such that the underlying location of the molecule is determined to within a few nanometers. The resulting coordinate list of molecules can then be reconstructed to form a pseudo-image (Fig. 1). Importantly, this approach does not rely on intensity in the same way as most conventional fluorescence and other SRM methods, particularly insofar as the dominance of brighter regions in these images. Instead, in single-molecule SRM, fluorophores are detected without clustering or cumulative brightness-based effects. This engenders that individual proteins, replication forks (RFs), and double-strand breaks (DSBs) can be detected and examined within the cellular context. Moreover, SRM allows for spatial resolutions of approximately 20 nm when performing multicolor nuclear imaging, allowing for visual separation of spatially distinct RFs and DSBs, as well as unprecedented levels of insight into the internal organization of repair foci [2]. The SRM assay described herein is specifically designed to investigate homologous recombination (HR) repair of singleended DSBs (seDSBs) generated by RF stress. This damage induction pathway is a prevalent and important endogenous genomic

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instability source and a major contributor to human disease [3, 4]. SeDSB are a unique class of chromosomal breaks that occur at replication forks because of replication stress. SeDSBs cannot be repaired via the canonical nonhomologous end-joining (NHEJ) pathway because they lack a second end to ligate and, therefore, are necessarily repaired via HR [5]. In order to generate single-ended DSBs that mimic this endogenous stress, low doses of the small molecule drugs camptothecin (CPT) [6] or hydroxyurea (HU) [7] are applied to synchronized S phase cells in the assay described. Nascent DNA (naDNA) is simultaneously pulse labeled by addition of ethynyl deoxyuridine (EdU) so that RFs coincident with DSBs can be detected by “click” reaction after cell pre-extraction and fixation [8]. An array of replicative and repair proteins can then be detected using immunolabeling, including, as described previously and among many others, MRE11, CtIP, BRCA1, BRCA2, RAD51, PCNA, and RPA [2, 9–12]. SRM imaging is undertaken using either a commercially available singlemolecule SRM setup or, as in our case, a homebuilt bespoke microscope [13]. Finally, the single-molecule data are analyzed and rendered using openly available software or code, to produce a spatiotemporal map of repair/misrepair events [14, 15]. Herein, we detail these steps, including the required reagents, optimizations, and common troubleshooting.

2

Materials All solutions should be prepared using ultrapure water unless stated otherwise.

2.1 Cell Culture, Fixation, and Labeling

1. U-2 OS human bone osteosarcoma cells (ATCC HTB-96). 2. Culture medium: McCoy’s 5A medium supplemented with 10% fetal bovine serum and 100 U/ml penicillin-streptomycin. 3. Starvation medium: McCoy’s 5A medium supplemented with 100 U/ml penicillin-streptomycin. 4. 6-well plates. 5. No. 1.5 borosilicate coverslips 22
22 mm. 6. Dulbecco’s phosphate-buffered saline (D-PBS). 7. Dimethyl sulfoxide (DMSO). 8. 100 μM camptothecin in DMSO (see Note 1). 9. 10 mM ethynyl deoxyuridine (EdU) (see Note 2). 10. 10 mM bromodeoxyuridine (BrdU) (see Note 3). 11. 10–1000 μl autopipettes and sterile tips (see Note 1).

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12. Cytoskeleton pre-extraction (CSK) buffer: 10 mM HEPES, 300 mM Sucrose, 100 mM NaCl, 3 mM MgCl2, and 0.5% Triton X-100, pH ¼ 7.4, store without Triton X-100 for up to 2 weeks at 4  C. Solution should be used immediately after Triton X-100 added. 13. Fixation solution: Isotonic PBS containing 0.3% glutaraldehyde, 3.7% paraformaldehyde, pH ¼ 7.4. Make immediately prior to cell fixation using electron microscopy grade 32% paraformaldehyde and 70% glutaraldehyde. 14. Phosphate-buffered saline (PBS). 15. Blocking buffer: 2% glycine, 2% BSA, 0.2% gelatin, and 50 mM NH4Cl in isotonic PBS. 16. Click iT Plus EdU labeling kit with Alexa Fluor 488, 532, 568, or 647 conjugated fluorophore (ThermoFisher). 17. Primary antibodies against protein targets of interest, e.g., MRE11, RAD51, BRCA1, etc. (see Note 4). 18. Alexa Fluor 488, 532, 568, and 647 conjugated secondary antibodies compatible for multicolor immunolabeling as in confocal microscopy. 19. Anti-BrdU antibody (for labeling of ssDNA, see Note 3). 2.2

SRM Imaging

1. Glass slides. 2. Power drill capable of 20,000 rpm with 0.75 mm diamond drill bits. 3. Ultrapure water. 4. Double-sided tape. 5. Clear nail polish. 6. Photoswitching buffer: Isotonic PBS containing 1 mg/mL glucose oxidase, 0.02 mg/mL catalase, 10% glucose, 100 mM mercaptoethylamine (MEA), pH ¼ 8. Store frozen aliquots of 100
enzyme mixture (100 mg/ml glucose oxidase and 2 mg/ml catalase) and 1 M MEA for up to 6 months. Store 20% glucose in ultrapure water at 4  C for 2–4 weeks. Immediately prior to imaging, make up photoswitching buffer using 10
PBS such that the final solution is isotonic (see Note 5). 7. Tetraspecks mounted on cover glass to give clear individual bead signals.

3

Methods

3.1 Cell Culture, Fixation, and Labeling

1. Seed U-2 OS cells at 20% confluence in pre-warmed 37  C culture medium onto coverslips placed in the wells of a 6-well plate to a total volume of 1.5 ml per well. Tap coverslips down

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gently with a pipette tip to release any oxygen captured underneath. 2. Culture the cells for 12–24 h at 37  C in 5% CO2. 3. Wash the cells in pre-warmed 37  C DPBS and then replace the culture medium with pre-warmed 37  C starvation medium. Incubate for 48–72 h at 37  C in 5% CO2. 4. Replace starvation medium with 1.5 ml pre-warmed 37  C culture medium per well and incubate for 16 h at 37  C in 5% CO2 (see Note 6). 5. To achieve a low level of synchronized seDSB induction with naDNA labeling, remove half the culture medium (750 μl per well) and then add 750 μl pre-warmed 37  C culture medium containing 200 nM CPT and 20 μM EdU (see Note 7). To detect ssDNA using BrdU, add 20 μM BrdU. This will achieve final concentrations of 100 nM CPT, 10 μM EdU/BrdU. 6. Incubate for 1 h. 7. Remove the culture medium and immediately add 1.5 ml of 4  C CSK buffer to each well for 2–3 min (see Note 8). 8. Remove most of the CSK Buffer (~80–90%) and add 1.5 ml of fixation solution to each well. Allow 10 min for fixation to occur. 9. Wash cells 3
with PBS. 10. Incubate cells in blocking buffer overnight at 4  C or for at least 1 h at room temperature. With the introduction of fluorophores, all subsequent steps should be performed with minimal exposure to light. To achieve this, reaction and antibody incubation should be performed in the dark, and samples should be stored wrapped in foil. 11. After blocking cells, remove all liquids from each well and perform click labeling of the naDNA as detailed in the ClickiT Plus protocol, using 100 μl of reaction cocktail per coverslip and labeling for 1 hour. The reaction cocktail contains the Alexa Fluor picolyl azide along with other proprietary reagents for an optimum reaction (see Note 9). 12. Wash cells 3
in PBS. 13. Dilute primary antibodies (including anti-BrdU) in blocking buffer and apply 100 μl to each coverslip. Allow staining for 2 h (see Note 10). 14. Wash cells 3
in PBS. 15. Apply 100 μl of diluted secondary antibodies in blocking buffer and stain for 1 h (see Note 10). 16. Wash cells 3
in PBS and store at 4  C for up to 2 weeks if needed.

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3.2 SRM Imaging, Rendering, and Analysis

1. Using the 0.75 mm drill bit at 20,000 rpm, drill two holes into a standard slide, vertically separated by 1 cm. Make sure both the top and bottom surfaces of the slide are wet to minimize friction, and that the glass is placed over soft plastic during drilling. 2. Prior to imaging, remove the cover glass from the 6-well plate and place it over the drilled glass slide such that the cells are sandwiched in between and both holes are as near the center of the cover glass as possible. Use double-sided tape as a spacer along the edge of the glass slide. 3. Seal the cover glass by applying nail polish around its edge and allowing it to dry. 4. Using an autopipette, push 100 μl of PBS into the chamber through one of the two holes. Liquid should escape the second hole but not anywhere else (see Note 11). 5. Immediately prior to imaging, replace the PBS in the chamber by pushing through >200 μl of photoswitching buffer. 6. Mount the slide on an appropriate SRM setup and bring the sample into focus by visualizing the nascent DNA channel at low illumination power. If possible, use highly inclined and laminated optical sheet (HILO) illumination (see Note 12). 7. Focusing through the middle of nuclei of interest, increase the excitation laser power to achieve single molecule blinking and acquire 2–20,000 frames at 20–100 Hz (see Note 13). 8. Following acquisition of the naDNA channel, switch dichroic and emission filters and sequentially image the remaining fluorophore channels (see Note 14). 9. Repeat steps 7 and 8 for other fields of view until sufficient data are collected. 10. Raw multi-frame image stacks of single-molecule data can be processed using any one of the commercially or openly available software or code packages (for a summary of those available see [16]). These programs produce a list of molecular coordinates which can be rendered into a pseudo-image of the naDNA and immunolabeled proteins. 11. To correct for channel-to-channel chromatic aberration, a polynomial elastic transformation describing these aberrations must be produced. To achieve this, images of dispersed Tetraspeck beads in each imaged channel should be acquired immediately prior or subsequent to an imaging session. 12. These images of Tetraspecks can then be used to calculate the necessary transformations, which can then be applied to correct coordinate lists and rendered SRM images.

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13. Coordinate lists and rendered pseudo-images can be analyzed using several approaches, including common confocal methods, to quantify total signal, colocalization, spatial arrangement, and protein–protein interrelationships at DSBs. For a review of these methods see [17].

4

Notes 1. CPT is not readily soluble in water and must be stored in DMSO. CPT should be prepared as 1000
stock in DMSO and diluted into >10 ml of culture medium prior to dosing. This limits the potential interference of DMSO in the experiment while maintaining a reliable dosage value. Furthermore, well-calibrated autopipettes are required for drug dilutions. Control cells must be treated with equivalent doses of DMSO. If different drug doses are required, stock concentrations and control experiments should take this into account. 2. EdU is provided as part of the Click-iT Plus EdU Imaging kit. 3. BrdU is used to detect single-stranded DNA and should only be used if this is a desired target of the experiment. 4. Ideally, choose primary antibodies that have previously been used for SRM. If these are not available, antibodies validated for confocal immunofluorescence are likely to work well for SRM but will often require different dilutions. Unproven antibodies must be validated for use in SRM. 5. The composition of the photoswitching buffer should be optimized for different SRM setups and different samples, including changes in damage induction and antibodies. This is because the photoswitching buffer is used to achieve single molecule blinking by controlling the rate at which fluorophores are “switched off” via the MEA concentration, and the rate at which they are switched back “on” via the molecular oxygen concentration. Depending on the intensity of the excitation light at the sample, and the density of fluorophores present, these rates will change. 6. 16 h release after starvation causes the U-2 OS cells to be in mid-S phase. This time will need to be optimized using cell cycle analysis for different cell lines. 7. Do not remove and replace all of the culture medium as this will likely affect the rate of replication and, therefore, the nascent DNA signal. 8. The CSK buffer removes much of the cytosolic and soluble nuclear components which ultimately lowers nonspecific staining and biases the proteins detected to those bound to chromatin which are, therefore, more likely to be playing a role in

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repair. However, this pre-extraction can cause cells to detach from the coverslips and so the timing must be carefully optimized. If possible, cells should be observed using a bright-field microscope during this process. High confluence increases the likelihood of cell detachment. 9. For all labeling reactions on the coverslips, only 100 μl of reagent is needed. To avoid drying out of the slides during labeling, make sure that all liquid within the well is aspirated and that the coverslip is centered with no corners touching the sides, before careful application of the 100 μl to the cover glass. If required, cells can be labeled within a humidity chamber. 10. Antibody concentrations should be determined for each new target by titrating concentrations and analyzing signal and structure. Recommended concentrations for immunofluorescence are a useful starting point; however, for SRM, samples typically need higher concentrations to satisfy the Nyquist sampling theorem. Incubation times should also be tested to optimize staining. 11. At this point, the mounted cover glass can be stored for 1–2 weeks at 4  C by sealing the two drilled holes with tape. 12. SRM setups, whether commercial or homebuilt, comprise a high-NA 60
or 100
objective, high power continuous wave lasers, and highly sensitive EMCCD or sCMOS cameras. 13. It is imperative that single molecule blinking is achieved with minimal overlapping emission patterns present during data acquisition. Overlapping emissions result in image artifacts and severely impact analyses. To achieve good single molecule blinking, various parameters should be empirically optimized including photoswitching buffer composition, laser intensity, beam alignment, and imaging frame rate. For a complete discussion see [18, 19]. 14. The order in which channels are acquired should also be optimized because different lasers can bleach and/or photoswitch other channels via crosstalk mechanisms. Typically, channels should be imaged in order of decreasing wavelength, but this may differ for specific antibody and fluorophore combinations.

Acknowledgments D.R.W. would like to acknowledge support from a Bruce Stone Fellowship from the La Trobe Institute for Molecular Science and funding via the Bendigo Tertiary Education Anniversary Foundation. Research in the Rothenberg lab is supported by funds from the NIH R01 GM108119, American Cancer Society (ACS: 130304-RSG-16-241-01-DMC), the V Foundation for Cancer Research (D2018-020), and Fondation Leducq (17CVD02).

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References 1. Bates M, Huang B, Dempsey GT, Zhuang X (2007) Multicolor super-resolution imaging with photo-switchable fluorescent probes. Science 317(5845):1749–1753 2. Whelan DR et al (2018) Spatiotemporal dynamics of homologous recombination repair at single collapsed replication forks. Nat Commun 9(1):3882 3. Saleh-Gohari N et al (2005) Spontaneous homologous recombination is induced by collapsed replication forks that are caused by endogenous DNA single-strand breaks. Mol Cell Biol 25(16):7158–7169 4. Zeman MK, Cimprich KA (2014) Causes and consequences of replication stress. Nat Cell Biol 16(1):2–9 5. Conlin MP et al (2017) DNA ligase IV guides end-processing choice during nonhomologous end joining. Cell Rep 20(12):2810–2819 6. Liu LF et al (2000) Mechanism of action of camptothecin. Ann N Y Acad Sci 922(1):1–10 7. Petermann E, Luis Orta M, Issaeva N, Schultz N, Helleday T (2010) Hydroxyureastalled replication forks become progressively inactivated and require two different RAD51mediated pathways for restart and repair. Mol Cell 37(4):492–502 8. Salic A, Mitchison TJ (2008) A chemical method for fast and sensitive detection of DNA synthesis in vivo. Proc Natl Acad Sci 105(7):2415 9. Daddacha W et al (2017) SAMHD1 promotes DNA end resection to facilitate DNA repair by homologous recombination. Cell Rep 20 (8):1921–1935 10. D’Alessandro G et al (2018) BRCA2 controls DNA:RNA hybrid level at DSBs by mediating RNase H2 recruitment. Nat Commun 9 (1):5376

11. Reid DA et al (2015) Organization and dynamics of the nonhomologous end-joining machinery during DNA double-strand break repair. Proc Natl Acad Sci U S A 112(20): E2575–E2584 12. Yin YD, Rothenberg E (2016) Probing the spatial organization of molecular complexes using triple-pair-correlation. Sci Rep 6:30819 13. Whelan DR, Holm T, Sauer M, Bell TDM (2014) Focus on super-resolution imaging with direct stochastic optical reconstruction microscopy (dSTORM). Aust J Chem 67 (2):179–183 14. Bermudez-Hernandez K et al (2017) A method for quantifying molecular interactions using stochastic modelling and superresolution microscopy. Sci Rep 7(1):14882 15. Schnitzbauer J et al (2018) Correlation analysis framework for localization-based superresolution microscopy. Proc Natl Acad Sci 115 (13):3219 16. Sage D et al (2019) Super-resolution fight club: assessment of 2D and 3D single-molecule localization microscopy software. Nat Methods 16(5):387–395 17. Durisic N, Cuervo LL, Lakadamyali M (2014) Quantitative super-resolution microscopy: pitfalls and strategies for image analysis. Curr Opin Chem Biol 20:22–28 18. Whelan DR, Bell TDM (2015) Superresolution single-molecule localization microscopy: tricks of the trade. J Phys Chem Lett 6 (3):374–382 19. Whelan DR, Bell TDM (2015) Image artifacts in single molecule localization microscopy: why optimization of sample preparation protocols matters. Sci Rep 5:7924

Chapter 25 The Analysis of Recombination-Dependent Processing of Blocked Replication Forks by Bidimensional Gel Electrophoresis Karol Kramarz, Anissia Ait Saada, and Sarah A. E. Lambert Abstract The perturbation of the DNA replication process is a threat to genome stability and is an underlying cause of cancer development and numerous human diseases. It has become central to understanding how stressed replication forks are processed to avoid their conversion into fragile and pathological DNA structures. The engineering of replication fork barriers (RFBs) to conditionally induce the arrest of a single replisome at a defined locus has made a tremendous impact in our understanding of replication fork processing. Applying the bidimensional gel electrophoresis (2DGE) technique to those site-specific RFBs allows the visualization of replication intermediates formed in response to replication fork arrest to investigate the mechanisms ensuring replication fork integrity. Here, we describe the 2DGE technique applied to the site-specific RTS1RFB in Schizosaccharomyces pombe and explain how this approach allows the detection of arrested forks undergoing nascent strands resection. Key words Bidimensional gel electrophoresis, Replication and recombination intermediates, Schizosaccharomyces pombe, Site-specific replication fork barrier, Psoralen cross-links, Benzoylated naphthoylated DEAE-cellulose

1

Introduction The analysis of replication intermediates by bidimensional gel electrophoresis(2DGE) has proved to be a powerful technique to investigate the dynamics of DNA replication at specific loci in a cell population [1]. This technique allows the separation of DNA molecules according to their mass, during the first dimension, and their shape, during the second dimension. This technique exploits the fact that a branched DNA molecule has a reduced rate of migration in agarose gels compared to a linear DNA molecule of the same mass [2]. The 2DGE was first applied to a Saccharomyces cerevisiae replicative plasmid by Bonita Brewer and Walton Fangman to map replication origin on plasmids [3] and then to map replication origins and replication fork barrier (RFB) at the genome

Andre´s Aguilera and Aura Carreira (eds.), Homologous Recombination: Methods and Protocols, Methods in Molecular Biology, vol. 2153, https://doi.org/10.1007/978-1-0716-0644-5_25, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Schematic examples of replication and recombination intermediates detected by bidimensional gel electrophoresis (2DGE) within a defined restriction fragment. Blue: bubble arc corresponding to the firing of a replication origin. Dark: small and large Y arc molecules. Purple: forks converging toward each other. Red: forks arrested by a site-specific obstacle. Green: X-spike structures corresponding to joint molecules. R: restriction site; LMW: Low molecular weight; HMW: Hight molecular weight

scale [4, 5]. Combined with site-specific RFBs and yeast genetics, 2DGE has become a popular technique to analyze in depth replication and recombination intermediates [1, 6, 7]. Those pioneering works have allowed to detect several replication intermediates within a defined restriction DNA fragment (Fig. 1): bubble arc corresponding to the firing of a defined replication origin, small and large Y arc corresponding to the progression of replication forks, and double Y arc corresponding to the progression of two converging forks. The accumulation of signal on the Y arc corresponds to replication forks arrested by a specific obstacle. Recombination intermediates have been further identified by 2DGE as X-shaped molecules [8]. Additional procedures to stabilize in vivo branched DNA molecules [cross-linking with trimethyl psoralen (TMP)] and to enrich replication intermediate on benzoylated naphthoylated DEAE-cellulose columns (BND) have considerably improved the detection of low abundant and transient DNA structures. Here, we describe a robust protocol to prepare replication intermediates from asynchronous Schizosaccharomyces pombe cells harboring an engineered site-specific and polar RFB called RTS1 (RTS1-RFB) [9] in order to detect replication and recombination intermediates formed upon activation of the RFB. The overall procedure consists on (Fig. 2):

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1. Set up a cell culture in which the RTS1-RFB is active or not (see Subheading 2.1 and [9]) (Subheading 3.1). 2. The freezing of replication intermediates using TMP + UV-A. This optional step allows to preserve in vivo replication intermediates and to prevent them to migrate in vitro (Subheading 3.2). 3. The restriction digestion of genomic DNA from cells embedded in agarose plugs (Subheading 3.3 and 3.4). 4. The enrichment of replication intermediates on BND column (Subheading 3.5). 5. The separation of replication intermediates by bidimensional gel electrophoresis (Subheading 3.6 and 3.7). 6. Southern blot to reveal the DNA fragment of interest (Subheading 3.8). Applying this 2DGE procedure to the fission yeast RTS1-RFB has allowed the identification of a novel replication intermediate corresponding to an arrested fork in which nascent strands undergo resection by the nuclease Exo1 [6]. As shown in Fig. 2 (Subheading 3.8), when the RTS1-RFB is inactive (RFB OFF), small and large Y arc were detected showing that replication forks progress homogenously through the locus analyzed. When the RTS1-RFB is active (RFB ON), the intensity of the large Y arc decreased and signal corresponding to arrested forks accumulated on the ascending Y arc. In addition, a tail signal emanating from arrested forks and descending toward the linear arc was detected (see red arrow on Fig. 2, Subheading 3.8). The migration pattern indicated that the corresponding DNA molecules were losing mass and shape, compared to arrested forks. This is consistent with nascent strands being resected (loss of mass) and becoming single stranded (loss of shape). Finally, this tail signal was abolished in the absence of the nuclease Exo1, further supporting that the corresponding DNA molecules are arrested forks undergoing nascent strands resection [6, 7].

2

Materials All solutions should be prepared using ultrapure, deionized water, and analytical grade reagents. After addition of sodium azide to cell cultures, all centrifugation steps must be performed at 4
C.

2.1 Strains and Cell Culture

1. Genotype of a WT strain harboring the RTS1-RFB (Fig. 2 top right panel of Subheading 3.1): sup35:nmt41:rtf1, ade6–704, leu1–32, t-ura4