Single Stranded DNA Binding Proteins (Methods in Molecular Biology, 2281) 1071612891, 9781071612897

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Methods in Molecular Biology 2281

Marcos T. Oliveira Editor

Single Stranded DNA Binding Proteins

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.

Single Stranded DNA Binding Proteins Edited by

Marcos T. Oliveira Jaboticabal, Brazil

Editor Marcos T. Oliveira Jaboticabal, Brazil

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1289-7 ISBN 978-1-0716-1290-3 (eBook) https://doi.org/10.1007/978-1-0716-1290-3 © 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. 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 There is no question that single-stranded DNA-binding proteins (SSBs) are important players in all processes required for genome maintenance. This fact does not seem to deaccelerate the rate with which research on SSBs is conducted across the globe, and in fact, I would argue that often working with SSBs represents the first step for many to start their careers in or move to the fields of DNA replication, repair, and/or recombination. This is exactly what happened to me in the beginning of my PhD: having had no prior experience other than what I learned in undergraduate biochemistry classes, my thesis advisor gave me the task of purifying a small series of mitochondrial SSB mutants and determining their single-stranded DNA-binding affinities. In her own words, “there is nothing like a toy protein for a beginner to practice protein biochemistry,” referring to the fact that SSBs are relatively more stable in solution that other replication proteins (I am actually not sure if those were her exact words, but I can certainly invoke poetic license in this context). To make a long story short, it turned out that the mitochondrial SSB and its variant forms had some particular properties, and their function in vitro (or lack thereof) varied according to changes in the protein preparation protocol. Most changes were artifacts that had to be circumvented before accurate functional details of the proteins could be described. I had to “play” a lot with that toy protein for us to figure what was going on, and unfortunately none of that has been recorded in my thesis or any of my publications. Others have also been “playing” with their favorite toy SSBs, and sometimes they invent their own “games.” This volume of Methods in Molecular Biology is a compilation of “games” (procedures) that researchers from almost all continents have developed and/or have been using to “play” (study) with their SSB of interest and are willing to share in a way that another researcher could use. Some of these “games,” such as single-molecule-based approaches, have definitely diversified and grown in use in the last 10 years, and have helped reveal new roles for SSB, representing a significant portion of this book. Other “games” represent more traditional biochemical, physiological, and/or structural approaches that can still provide valuable insights into SSB function and structure, especially for newly identified proteins. After the first two chapters, which are review articles that can help beginners start understanding SSB biology, this volume is organized in a taxonomical order, starting with protocols to study bacterial SSBs (Chapters 3–8), moving to eukaryotic (Chapters 8–15), mitochondrial (Chapters 16–20), and finalizing with viral SSBs (Chapters 21–23). I would like to thank all authors for their contribution, which is particularly important during such sensitive times that all countries are currently going through due to the spread of COVID-19. We have all been directly or indirectly affected by the changes in work and lifestyles we were rightfully forced to adopt, so I can say that finalizing this volume in a timely fashion was a challenge. I also would like to acknowledge the posthumous contribution of Dr. Margarita Salas to this book, who was a pioneer in studying bacteriophage ϕ29 DNA replication and has sadly passed away last year. I am glad that part of this book is now also a part of her scientific legacy. Jaboticabal, SP, Brazil

Marcos T. Oliveira

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

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1 The Essential, Ubiquitous Single-Stranded DNA-Binding Proteins . . . . . . . . . . . Marcos T. Oliveira and Grzegorz L. Ciesielski 2 Single-Stranded DNA-Binding Proteins in the Archaea . . . . . . . . . . . . . . . . . . . . . . Najwa Taib, Simonetta Gribaldo, and Stuart A. MacNeill 3 Single-Molecule Fluorescence Methods to Study Protein Exchange Kinetics in Supramolecular Complexes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Richard R. Spinks, Lisanne M. Spenkelink, and Antoine M. van Oijen 4 Comparing SSB-PriA Functional and Physical Interactions in Gram-Positive and -Negative Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yen-Hua Huang and Cheng-Yang Huang 5 In Vivo Binding of Single-Stranded DNA-Binding Protein to Stalled Replication Fork Helicases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cong Yu and Piero R. Bianco 6 Magnetic Tweezers-Based Single-Molecule Assays to Study Interaction of E. coli SSB with DNA and RecQ Helicase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Debjani Bagchi, Weiting Zhang, Samar Hodeib, Bertrand Ducos, Vincent Croquette, and Maria Manosas 7 High-Throughput Screening to Identify Inhibitors of SSB-Protein Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Andrew F. Voter 8 Single-Molecule Tethered Particle Motion Studies on the DNA Recombinase Filament Assembly and Disassembly . . . . . . . . . . . . . . . . . . . . . . . . . . Chih-Hao Lu, Wei-Hsuan Lan, and Hung-Wen Li 9 Generation of Fluorescent Versions of Saccharomyces cerevisiae RPA to Study the Conformational Dynamics of Its ssDNA-Binding Domains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sahiti Kuppa, Nilisha Pokhrel, Elliot Corless, Sofia Origanti, and Edwin Antony 10 RPA-1 from Leishmania sp.: Recombinant Protein Expression and Purification, Molecular Modeling, and Molecular Dynamics Simulations Protocols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Carlos A. H. Fernandes, Edna G. O. Morea, and Maria Isabel N. Cano 11 Single-Stranded DNA Curtains for Single-Molecule Visualization of Rad51-ssDNA Filament Dynamics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Upasana Roy and Eric C. Greene 12 Following Trypanosoma cruzi RPA-DNA Interaction Using Fluorescent In Situ Hybridization Coupled with Immunofluorescence (FISH/IF). . . . . . . . . Raphael S. Pavani and Maria Carolina Elias

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Quantifying the Affinity of Trypanosoma cruzi RPA-1 to the Single-Stranded DNA Overhang of the Telomere Using Surface Plasmon Resonance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Marcela de Oliveira Vitarelli and Maria Carolina Elias Expression, Purification, and Solution-State NMR Analysis of the Two Human Single-Stranded DNA-Binding Proteins hSSB1 (NABP2/OBFC2B) and hSSB2 (NAPB1/OBFC2A) . . . . . . . . . . . . . . . . . . . . . . . Serene El-Kamand, Mar-Dean Du Plessis, Teegan Lawson, Liza Cubeddu, and Roland Gamsjaeger Atomic Force Microscopy Reveals that the Drosophila Telomere-Capping Protein Verrocchio Is a Single-Stranded DNA-Binding Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alessandro Cicconi, Emanuela Micheli, Grazia Daniela Raffa, and Stefano Cacchione Analysis of Mitochondrial SSB-DNA Complexes and Their Effects on DNA Polymerase γ Activity by Electron Microscopy and Enzymatic Assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Oya Bermek and Grzegorz L. Ciesielski Optical Tweezers to Investigate the Structure and Energetics of Single-Stranded DNA-Binding Protein-DNA Complexes . . . . . . . . . . . . . . . . . Jose´ A. Morin, Fernando Cerro n, Francisco J. Cao-Garcı´a, and Borja Ibarra Measurements of Real-Time Replication Kinetics of DNA Polymerases on ssDNA Templates Coated with Single-Stranded DNA-Binding Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fernando Cerro n and Borja Ibarra Selective Suppression of Endogenous Gene Expression Using RNAi in Drosophila Schneider S2 Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yuichi Matsushima Stimulation of Variant Forms of the Mitochondrial DNA Helicase Twinkle by the Mitochondrial Single-Stranded DNA-Binding Protein. . . . . . . . . Ana P. C. Rodrigues and Marcos T. Oliveira Measuring the Complex Effects of the Single-Stranded DNA-Binding Protein gp2.5 on Primer Synthesis and Extension by the Bacteriophage T7 Replisome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alfredo J. Hernandez Strand Displacement and Unwinding Assays to Study the Concerted Action of the DNA Polymerase and SSB During Phi29 TP-DNA Replication. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alicia del Prado and Margarita Salas Structural Characterization of a Single-Stranded DNA-Binding Protein: A Case Study of the ORF6 Protein from Bacteriophage Enc34 . . . . . . . . . . . . . . . Elina Cernooka, Janis Rumnieks, and Andris Kazaks

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

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Contributors EDWIN ANTONY • Department of Biochemistry and Molecular Biology, Saint Louis University School of Medicine, St. Louis, MO, USA DEBJANI BAGCHI • Physics Department, Faculty of Science, Maharaja Sayajirao University of Baroda, Vadodara, Gujarat, India OYA BERMEK • National Institute of Environmental Health Sciences (NIEHS), Research Triangle Park, NC, USA PIERO R. BIANCO • Department of Pharmaceutical Sciences, College of Pharmacy, University of Nebraska Medical Center, Omaha, Nebraska, USA STEFANO CACCHIONE • Dipartimento di Biologia e Biotecnologie ‘C. Darwin’, Sapienza, ` di Roma, Rome, Italy Universita MARIA ISABEL N. CANO • Department of Chemical and Biological Sciences, Biosciences Institute, Sa˜o Paulo State University (UNESP), Botucatu, SP, Brazil FRANCISCO J. CAO-GARCI´A • Departamento Estructura de la Materia, Fı´sica Te´rmica y Electro nica, Universidad Complutense, Madrid, Spain; Instituto Madrilen ˜ o de Estudios Avanzados en Nanociencia, IMDEA Nanociencia, Madrid, Spain ELINA CERNOOKA • Latvian Biomedical Research and Study Centre, Riga, Latvia FERNANDO CERRO´N • Hospital Universitario de Canarias, Santa Cruz de Tenerife, Spain ALESSANDRO CICCONI • Department of Laboratory Medicine, Yale University School of Medicine, New Haven, CT, USA; Dipartimento di Biologia e Biotecnologie ‘C. Darwin’, ` di Roma, Roma, Italy Sapienza, Universita GRZEGORZ L. CIESIELSKI • Department of Chemistry, Auburn University at Montgomery, Montgomery, AL, USA ELLIOT CORLESS • Department of Biological Sciences, Marquette University, Milwaukee, WI, USA VINCENT CROQUETTE • Laboratoire de Physique Statistique, De´partement de physique de l’ENS, E´cole normale supe´rieure, PSL Research University, Universite´ Paris Diderot, Sorbonne Paris Cite´, Sorbonne Universite´s, UPMC University Paris 06, CNRS, Paris, France; IBENS, De´partement de biologie, E´cole normale supe´rieure, CNRS, INSERM, PSL Research University, Paris, France LIZA CUBEDDU • School of Science, Western Sydney University, Penrith, NSW, Australia; School of Life and Environmental Sciences, University of Sydney, Sydney, NSW, Australia BERTRAND DUCOS • Laboratoire de Physique Statistique, De´partement de physique de l’ENS, E´cole normale supe´rieure, PSL Research University, Universite´ Paris Diderot, Sorbonne Paris Cite´, Sorbonne Universite´s, UPMC University Paris 06, CNRS, Paris, France; IBENS, De´partement de biologie, E´cole normale supe´rieure, CNRS, INSERM, PSL Research University, Paris, France; High Throughput qPCR Core Facility of the ENS, qPCR-HD-GPC, Universite´ PSL, CNRS, IBENS, 46 rue d’Ulm, Paris, France MARIA CAROLINA ELIAS • Laboratorio de Ciclo Celular and Center of Toxins, Immune Response and Cell Signaling (CeTICS), Instituto Butantan, Sa˜o Paulo, SP, Brazil; Cell Cycle Laboratory, Butantan Institute, Sa˜o Paulo, SP, Brazil SERENE EL-KAMAND • School of Science, Western Sydney University, Penrith, NSW, Australia

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CARLOS A. H. FERNANDES • Department of Biophysics and Pharmacology, Biosciences Institute, Universidade Estadual Paulista “Ju´lio de Mesquita Filho”, Jaboticabal, SP, Brazil ROLAND GAMSJAEGER • School of Science, Western Sydney University, Penrith, NSW, Australia; School of Life and Environmental Sciences, University of Sydney, Sydney, NSW, Australia ERIC C. GREENE • Department of Biochemistry and Molecular Biophysics, Columbia University, New York, NY, USA SIMONETTA GRIBALDO • Unit Evolutionary Biology of the Microbial Cell, Department of Microbiology, Institut Pasteur, Paris, France ALFREDO J. HERNANDEZ • Department of Biological Chemistry and Molecular Pharmacology, Harvard Medical School, Boston, MA, USA SAMAR HODEIB • High Throughput qPCR Core Facility of the ENS, qPCR-HD-GPC, Universite´ PSL, CNRS, IBENS, Paris, France YEN-HUA HUANG • School of Biomedical Sciences, Chung Shan Medical University, Taichung City, Taiwan CHENG-YANG HUANG • School of Biomedical Sciences, Chung Shan Medical University, Taichung City, Taiwan; Department of Medical Research, Chung Shan Medical University Hospital, Taichung City, Taiwan BORJA IBARRA • Instituto Madrilen ˜ o de Estudios Avanzados en Nanociencia, IMDEA Nanociencia, Madrid, Spain ANDRIS KAZAKS • Latvian Biomedical Research and Study Centre, Riga, Latvia SAHITI KUPPA • Department of Biochemistry and Molecular Biology, Saint Louis University School of Medicine, St. Louis, MO, USA WEI-HSUAN LAN • Department of Chemistry, National Taiwan University, Taipei, Taiwan TEEGAN LAWSON • School of Science, Western Sydney University, Penrith, NSW, Australia HUNG-WEN LI • Department of Chemistry, National Taiwan University, Taipei, Taiwan CHIH-HAO LU • Department of Chemistry, National Taiwan University, Taipei, Taiwan; Department of Chemistry, Stanford University, Stanford, CA, USA MARIA MAN˜OSAS • Departament de Fı´sica de la Materia Condensada, Universitat de Barcelona, Barcelona, Spain STUART A. MACNEILL • Biomedical Sciences Research Complex, School of Biology, University of St Andrews, St Andrews, UK MARIA MANOSAS • Small Biosystems Lab, Departament de Fı´sica de la Mate`ria Condensada, Facultat de Fı´sica, Universitat de Barcelona, Barcelona, Spain YUICHI MATSUSHIMA • Department of Clinical Chemistry and Laboratory Medicine, Graduate School of Medical Sciences, Kyushu University, Fukuoka, Japan EMANUELA MICHELI • Dipartimento di Biologia e Biotecnologie ‘C. Darwin’, Sapienza, ` di Roma, Roma, Italy Universita EDNA G. O. MOREA • Department of Chemical and Biological Sciences, Biosciences Institute, Universidade Estadual Paulista “Ju´lio de Mesquita Filho”, Jaboticabal, SP, Brazil JOSE´ A. MORIN • Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, Germany ANTOINE M. VAN OIJEN • Molecular Horizons and School of Chemistry and Molecular Bioscience, University of Wollongong, Wollongong, NSW, Australia; Illawarra Health and Medical Research Institute, Wollongong, NSW, Australia

Contributors

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MARCOS T. OLIVEIRA • Departamento de Tecnologia, Faculdade de Cieˆncias Agra´rias e Veterina´rias, Universidade Estadual Paulista “Ju´lio de Mesquita Filho”, Jaboticabal, SP, Brazil MARCELA DE OLIVEIRA VITARELLI • Cell Cycle Laboratory, Butantan Institute, Sa˜o Paulo, SP, Brazil; Center of Toxins, Immune Response and Cell Signaling (CeTICS), Butantan Institute, Sa˜o Paulo, SP, Brazil; Laboratorio de Ciclo Celular and Center of Toxins, Immune Response and Cell Signaling (CeTICS), Instituto Butantan, Sa˜o Paulo, SP, Brazil SOFIA ORIGANTI • Department of Biology, Saint Louis University, St. Louis, MO, USA RAPHAEL S. PAVANI • Laboratorio de Ciclo Celular and Center of Toxins, Immune Response and Cell Signaling (CeTICS), Instituto Butantan, Sa˜o Paulo, SP, Brazil MAR-DEAN DU PLESSIS • School of Science, Western Sydney University, Penrith, NSW, Australia NILISHA POKHREL • Department of Biological Sciences, Marquette University, Milwaukee, WI, USA; New England Biolabs, Ipswich, MA, USA ALICIA DEL PRADO • Centro de Biologı´a Molecular Severo Ochoa, Consejo Superior de Investigaciones Cientı´ficas, Universidad Autonoma de Madrid, Madrid, Spain GRAZIA DANIELA RAFFA • Dipartimento di Biologia e Biotecnologie ‘C. Darwin’, Sapienza, ` di Roma, Rome, Italy Universita ANA P. C. RODRIGUES • Departamento de Tecnologia, Faculdade de Cieˆncias Agra´rias e Veterina´rias, Universidade Estadual Paulista “Ju´lio de Mesquita Filho”, Jaboticabal, SP, Brazil UPASANA ROY • Department of Biochemistry and Molecular Biophysics, Columbia University, New York, NY, USA JANIS RUMNIEKS • Latvian Biomedical Research and Study Centre, Riga, Latvia MARGARITA SALAS • Centro de Biologı´a Molecular Severo Ochoa (Consejo Superior de Investigaciones Cientı´ficas-Universidad Autonoma de Madrid), Universidad Autonoma, Madrid, Spain LISANNE M. SPENKELINK • Molecular Horizons and School of Chemistry and Molecular Bioscience, University of Wollongong, Wollongong, NSW, Australia; Illawarra Health and Medical Research Institute, Wollongong, NSW, Australia RICHARD R. SPINKS • Molecular Horizons and School of Chemistry and Molecular Bioscience, University of Wollongong, Wollongong, NSW, Australia; Illawarra Health and Medical Research Institute, Wollongong, NSW, Australia NAJWA TAIB • Unit Evolutionary Biology of the Microbial Cell, Department of Microbiology, Institut Pasteur, Paris, France; Hub Bioinformatics and Biostatistics, Department of Computational Biology, Institut Pasteur, Paris, France ANDREW VOTER • Department of Biomolecular Chemistry, University of Wisconsin School of Medicine and Public Health, Madison, WI, USA CONG YU • Department of Pharmaceutical Sciences, College of Pharmacy, University of Nebraska Medical Center, Omaha, Nebraska, USA WEITING ZHANG • Laboratoire de Physique Statistique, De´partement de physique de l’ENS, E´cole normale supe´rieure, PSL Research University, Universite´ Paris Diderot, Sorbonne Paris Cite´, Sorbonne Universite´s, UPMC University Paris 06, CNRS, Paris, France; IBENS, De´partement de biologie, E´cole normale supe´rieure, CNRS, INSERM, PSL Research University, Paris, France

Chapter 1 The Essential, Ubiquitous Single-Stranded DNA-Binding Proteins Marcos T. Oliveira and Grzegorz L. Ciesielski Abstract Maintenance of genomes is fundamental for all living organisms. The diverse processes related to genome maintenance entail the management of various intermediate structures, which may be deleterious if unresolved. The most frequent intermediate structures that result from the melting of the DNA duplex are single-stranded (ss) DNA stretches. These are thermodynamically less stable and can spontaneously fold into secondary structures, which may obstruct a variety of genome processes. In addition, ssDNA is more prone to breaking, which may lead to the formation of deletions or DNA degradation. Single-stranded DNA-binding proteins (SSBs) bind and stabilize ssDNA, preventing the abovementioned deleterious consequences and recruiting the appropriate machinery to resolve that intermediate molecule. They are present in all forms of life and are essential for their viability, with very few exceptions. Here we present an introductory chapter to a volume of the Methods in Molecular Biology dedicated to SSBs, in which we provide a general description of SSBs from various taxa. Key words Genome maintenance, Replication, Repair, Recombination, Single-stranded DNA intermediates

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The OB Fold and the General Mechanism of ssDNA Binding SSB proteins share a structural motif, called the oligosaccharide/ oligonucleotide-binding (OB) fold that enables binding to singlestranded (ss) DNA with high specificity. The OB fold consists minimally of a five-stranded β-sheet arranged as a β-barrel usually capped by a single α-helix [1, 2] (Fig. 1a). In the ssDNA-OB fold complex, the DNA bases are often in close contact with the protein, while the phosphodiester groups are mostly exposed to the solvent. Nucleotides interact with the protein primarily via stacking interactions with aromatic amino acid side chains and packing interactions with hydrophobic side chains or the aliphatic portions of more polar groups such as lysine and arginine. Such nonpolar interactions can involve both the ribose rings and the bases of the nucleic acid. Additionally, hydrogen-bond donor and acceptor

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Bacterial SSBs. Structural representation of E. coli SSB, the archetype of bacterial-like SSBs, showing the OB fold of a single subunit (a), the homotetramer (b), and how the protein interacts with ssDNA in the (SSB)65 binding mode (c, see text for details). The C-terminal domain is unstructured and not shown. Each subunit of the tetrameric protein is represented with distinct colors. The software Pymol (www.pymol.org) was used to analyze the structure deposited on PDB under the accession code 1EYG, and to create the figure

groups from polar side chains can satisfy hydrogen bonds, providing recognition of the edges of specific bases [1, 3]. Although strong electrostatic interactions between SSB proteins and ssDNA are necessary to stabilize the latter, the transient role of SSB proteins in DNA metabolic processes indicates that the binding is weak enough to enable efficient recycling and repositioning of SSB proteins [4]. In addition, studies demonstrated that SSB proteins diffuse (or “slide”) on ssDNA with efficiency comparable to transcription factors, as opposed to being inert as previously thought [5–7]. The diffusion along ssDNA is important in the fast redistribution of SSB proteins on ssDNA after initial binding to a random location [8]. Notably, “sliding” SSB proteins can destabilize short DNA hairpins, which likely contributes to efficient processing of the ssDNA template [6, 9].

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Classification of SSB Proteins The OB folds of different SSB proteins can range in length from 70 to 150 aa, and the loops connecting the β-sheets that are responsible for binding specificity vary in sequence, length, and conformation. Furthermore, most SSB proteins share a low degree (5–25%) of general sequence similarity [1, 3]. Therefore, comparative approaches to the characterization of SSB proteins, based on sequence similarity alone, are difficult. However, a general quaternary organization of SSB proteins is largely consistent within individual kingdoms of life, while it is strikingly different between the

The Essential, Ubiquitous Single-Stranded DNA-Binding Proteins

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kingdoms. Therefore, the majority of SSB proteins can be classified based on their phylogenetic distribution as “bacterial” or “eukaryotic” [10].

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Bacterial SSB Proteins The “bacterial” SSB proteins are predominantly homotetrameric [11] (Fig. 1). The monomers of the most studied bacterial SSB protein, that of Escherichia coli (EcSSB) [12, 13], are 18.8 kDa in size and are composed of an N- and a C-terminal domain. The N-terminal domain consists of approximately 116 residues and contains a single OB fold [14–16] (Fig. 1a). The monomers assemble into a tetramer via two distinct protein-protein interfaces (Fig. 1b). The first of these interfaces is formed between two monomers, whereas the second interface is formed between two dimers [11]. Each SSB monomer has the potential for extensive contact with DNA in the assembled tetramer [15]. The kinetics of ssDNA binding is characterized by fast association and slow dissociation rates [17]. EcSSB has been demonstrated to bind ssDNA predominantly in two binding modes, which are reversibly interconvertible depending on the salt concentration and type, as well as its density on the ssDNA [13, 18]. In the 65 nucleotide-binding mode, (SSB)65, the ssDNA fully wraps around the tetramer, interacting with all four subunits (Fig. 1c), and is favored at high salt concentrations and low protein-to-DNA ratios [18, 19]. In the 35 nucleotide-binding mode, (SSB)35, the ssDNA interacts with only two subunits of the tetramer and is favored in low salt concentrations and upon high protein-to-DNA ratios [20–22]. Furthermore, EcSSB binds ssDNA in a cooperative manner, which is promoted in (SSB)35 but limited in the (SSB)65 binding mode [13, 23]. In both binding modes, the EcSSB tetramer is able to diffuse along ssDNA without dissociating [8]. It has long been speculated that because the properties of these wrapping modes differ significantly, they may be used preferentially in various DNA metabolic processes [13, 21, 24]. These suggestions have recently been supported by a study that showed that the PriA helicase imposes the conversion of EcSSB from the (SSB)65 to (SSB)35 binding mode, which licenses DNA replication restart [25]. The C-terminal domain consists of an intrinsically disordered linker (IDL) region of approximately 54 amino acids and a wellconserved tail formed by the last 8–10 predominantly acidic residues, terminated in a hydrophobic tripeptide [11, 14, 26]. The acidic tail plays a major role in interacting with an array of at least 14 proteins, termed the SSB interactome, which, among others, include DNA polymerases II, III, and V; exonuclease I; the DNA primase DnaG; and the DNA helicase PriA [10, 13, 24, 27–

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32]. The interaction with SSB facilitates the activity of the partner proteins [10, 12]; for example EcSSB stimulates the polymerase activity of polymerases II and III [33–35], facilitates loading, and stimulates the activities of PriA helicase [31, 32, 36] and exonuclease I [27]. Mutations within the sequence of the acidic tail of EcSSB are detrimental to cell growth [12]. For example, the P176S replacement (known as the ssb113 mutation) confers temperaturesensitive lethality by producing an SSB variant that, although still competent in ssDNA binding, cannot support DNA replication at nonpermissive temperatures due to a failure to interact with the replication machinery [37]. Together these studies indicate that, in addition to ssDNA stabilization, EcSSB poses as a hub for the recruitment of ssDNA processing factors, and thus serves as a major regulator of genome maintenance. Notably, the C-terminal tail also seems to play a role in ssDNA binding [38, 39]. Its deletion increases the intrinsic affinity of EcSSB for ssDNA and enhances the negative cooperativity between ssDNA-binding sites, indicating that the C-termini exert an inhibitory effect on ssDNA binding [38, 40]. Several bacterial species contain more than one SSB-coding gene. In these cases, one of the proteins is typically essential for bacterial growth (usually denoted as SsbA), whereas the other is not (usually denoted as SsbB). The nonessential SSB paralogs typically participate in processes related to bacterial natural competence [41, 42]. In Bacillus subtilis, BsSsbB is involved in natural competence-associated recombination, and share with SsbA the ssDNA-binding activities required during genetic recombination, by modulating RecA nucleation [43]. In Streptococcus pneumoniae, SpSsbB maintains the reservoir of naturally internalized ssDNA, thereby increasing the likelihood of multiple chromosomal transformation events in the same cell [42]. In addition, SsbB proteins also appear to be implicated in bacterial virulence. The SsbB of Neisseria gonorrhoeae is engaged in the secretion of chromosomal DNA to the media [44], and the SsbB of Agrobacterium tumefaciens, VirE2, functions as a molecular motor facilitating the transfer of the Ti plasmid and its stability in the host plant cells [45]. The SSB paralogs also often exhibit variations in structural organization. For example, BsSsbB lacks the C-terminal acidic tail, which likely alters its protein-protein interaction specificities, whereas BsSsbA resembles EcSSB, as each of the monomers contains the N-terminal OB fold and the C-terminal tail, and they interact to assemble a tetramer capable of binding ssDNA [41]. Moreover, although crystal structures suggest that SsbB binds ssDNA in a manner similar to SsbA, their DNA-binding properties are different [43]. BsSsbB binds to ssDNA with lower affinity than BsSsbA [43], which may be indicative of the role of the C-terminus in ssDNA binding, as suggested for the EcSSB protein [40]. However, the SsbB of Streptomyces coelicolor, which also lacks the C-terminal

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tail, has higher DNA-binding affinity than ScSsbA [46]. Notably, the ssDNA-binding affinity of the SsbB paralogs is generally more sensitive to increased salt concentration and presents lower to no cooperativity [43, 44, 46, 47]. Recently, a third paralog of SSB in Staphylococcus aureus, SsbC, has been identified. An initial study indicated that, similarly to the SsbB protein, it lacks the C-terminal tail [47], but further studies exploring its biological roles are necessary for the understanding of SSB diversification. Whereas the majority of bacterial SSB family members function as homotetramers, dimeric SSB proteins were discovered in a distinct bacterial lineage of extremophiles, the Deinococcus-Thermus group [48–51]. Similarly to the homotetrameric SSB proteins, the dimeric bacterial SSB proteins (DraSSB in Deinococcus radiodurans and TthSSB in Thermus thermophilus) retain four OB folds, two per monomer. However, the two OB folds within each monomer have distinct amino acid sequences and the functional SSB homodimer possesses only two conserved C-terminal tails [48, 51]. Notably, in response to ionizing radiation, DraSSB levels increase in D. radiodurans from ~20,000 to 56,000 dimers per cell, whereas E. coli maintains 200–3000 EcSSB tetramers per cell under normal conditions and in response to DNA-damaging agents [52]. The large number of DraSSB dimers per cell and its rapid increase in response to ionizing radiation certainly contribute to the high tolerance of D. radiodurans to such DNA-damaging condition [53]. These results indicate that the homodimeric SSBs found in extremophiles may have an expanded role in DNA metabolism. Studies indicated that although the ssDNA wraps around homodimeric SSB proteins in a way similar to that seen in the EcSSBssDNA complex, their ssDNA-binding affinities are considerably lower than those of EcSSB (Kd < 100 nM for DraSSB versus Kd < 1 nM for EcSSB, in 1 M salt) [54–56]. Similarly to homotetrameric SSB proteins, the homodimeric SSB proteins (at least TthSSB) are capable of rapid diffusion along ssDNA, indicating that this activity is conserved across different types of SSB proteins [57].

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Eukaryotic SSB Proteins The “eukaryotic” SSB protein is commonly referred to as the replication protein A (RPA) and functions generally as a heterotrimer (Fig. 2a) [58, 59]. Representative for this group, the human RPA (HsRPA), comprises 70, 32, and 14 kDa subunits termed RPA70, RPA32, and RPA14, respectively [58]. HsRPA contains six OB folds in total, referred to as DNA-binding domains (DBDs) A–F [1, 10, 60, 61]. Only four of the six DBDs are engaged in ssDNA binding and three of those are located in the RPA70 subunit (DBDs A–C) [62–65]. The fourth DNA-binding DBD (DBD-D) is located in the RPA32 subunit [66]. Though usually

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Fig. 2 Eukaryotic SSBs. Structural representations of the fungus Ustilago maydis RPA bound to ssDNA (a), hSSB1 bound to ssDNA (b) and human mtSSB (c). The structure of U. maydis RPA comprises DBD-A, DBD-B, and DBD-C of the RPA70 subunit; DBD-D of the RPA32 subunit; and DBD-E of the RPA14 subunit (see text for details). The structure of hSSB1 bound to a 35 nt ssDNA was extracted from the crystal structure of SOSS1 (see text for details). Each subunit of RPA and mtSSB is represented with distinct colors. The software Pymol (www.pymol.org) was used to analyze the structures deposited on PDB under the accession codes 4GNX, 4OWW, and 6RUP, respectively, and to create the figure

considered a nonspecific ssDNA-binding protein, HsRPA displays a slight preference for polypyrimidine tracts [67]. Binding is believed to occur in a sequential fashion [61, 68–70]: it is initiated by interaction of DBD-A and DBD-B with 8–10 nucleotides (nt) at the 50 -end of ssDNA [62], followed by a more stable intermediate 13–22 nt binding mode occurring with the additional involvement of DBD-C [63, 64]. Finally, the cooperative binding of RPA DBDs A–D occludes about 30 nt of ssDNA [67, 68]. The interaction of RPA with ssDNA is dynamic, and similarly to bacterial SSB proteins, RPA diffuses along the ssDNA [6]. This is facilitated by rapid binding and dissociation cycles of DBD-A and DBD-D, while RPA as a whole remains bound to ssDNA [70]. Furthermore, RPA can unwind double-stranded (ds) DNA substrates, in a salt-sensitive manner [6, 71, 72]. HsRPA contains a zinc finger motif on RPA70 that regulates ssDNA binding in a redox-dependent manner [73, 74]. ssDNA binding by RPA is enhanced over tenfold in reducing conditions [74], and because the cytoplasm is generally more of a reducing environment than the nucleus, it is possible that this increased DNA binding by RPA serves as a defense mechanism against foreign DNA, specifically viral DNA [10]. DBD-E and DBD-F primarily facilitate protein-protein interactions. DBD-E, the only OB fold in the RPA14 subunit, contributes to RPA multimerization, forming the trimerization core together with DBD-C and DBD-D [64, 75]. This structural role

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of RPA14 is essential for proper RPA function and its deletion is lethal in yeast [76]. In addition, DBD-E has been shown to weakly but specifically interact with single-stranded G-rich telomeric DNA sequences [77]. DBD-F, which is located at the N-terminus of RPA70 and is connected to the high-affinity DNA-binding core (DBD-A and -B) by an 67-residue flexible linker [78], contains a basic cleft that warrants the RPA’s ability to interact with partner proteins [79, 80]. These interactions are often aided by the C-terminal domain of the RPA32 subunit [81]. The RPA interactome (also referred to as RPA-interacting proteins, RIPs) includes factors engaged in DNA repair, recombination, and replication [10, 60, 82, 83]. RPA contributes to nucleotide excision repair by recruiting the XPA, XPG, and XPF-ERCC1 endonucleases to the lesion site [84–86]. In base excision repair (BER), RPA interacts with human uracil-DNA glycosylase UNG2 [81, 87, 88], and in the homologous recombinational repair of DNA double-strand breaks (DSBs), RPA interacts with and modulates the activities of Rad51 and Rad52 [89–94]. HsRPA has also been reported to interact with the breast cancer susceptibility proteins BRCA1 and BRCA2, two probable recombination mediators, as well as with the tumor suppressor p53 [61, 95–97]. In DNA replication, RPA participates in the initiation and elongation steps by enhancing the assembly and recruitment of DNA polymerases α, δ, and ε, by promoting polymerase switch on the lagging strand, and by coordinating the processing of Okazaki fragments [98–101]. During the different DNA metabolic processes, recruited factors displace RPA from ssDNA gradually, through modulation of the DNA-binding affinity of individual DBDs aligned sequentially along the DNA strand [60, 70]. In vivo, the function of RPA appears to be regulated by phosphorylation [102, 103]. The phosphorylation domain is unstructured and located at the N-terminus of the RPA32 subunit [75, 81, 104]. Phosphorylation or mutations that add multiple negative charges to the N-terminal phosphorylation domain of RPA decrease interactions with p53, DNA polymerase α, and other factors [105]. It also appears that phosphorylation reduces the DNA binding [106] and helix destabilization [107] activities, and promotes dissociation of the RPA complex [108]. However, these observations are not consistent as results indicating opposite effects have also been reported [105, 109, 110]. In humans and yeast, RPA proteins are phosphorylated in a cell cycle-dependent fashion from the G1/S transition throughout mitosis [111, 112]. Nevertheless, the relevance of these posttranslational modifications is in question, as the efficiency of DNA replication is not affected by mutations in the phosphorylation sites [113]. Interestingly, RPA becomes hyper-phosphorylated during mitosis in response to DNA-damaging agents such as UV or ionizing radiation, during treatment with the replication inhibitor hydroxyurea, and during

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cellular apoptosis [103, 112]. This appears to be a mechanism for inactivating RPA, as hyper-phosphorylated RPA does not bind to chromatin and the hyper-phosphorylation state disappears after the cell exits mitosis. Consistent with this, addition of Cdc2, the kinase acting on RPA, to purified interphase chromatin results in RPA foci disassembly [114]. Nevertheless, the roles of RPA phosphorylation clearly warrant further studies.

5

The “Other” Eukaryotic SSB Proteins

5.1 Single-Stranded DNA-Binding Proteins 1 and 2

RPA was thought to be the only SSB used for the maintenance of the nuclear genome, until two genes encoding “bacterial”-like SSBs (aside from the well-known mitochondrial SSB gene, see Subheading 5.2) were found [121, 123]. In humans, the two proteins were named hSSB1 and hSSB2 (or hNABP2 and 1, respectively), shown to share higher sequence and structural similarities with a group of archaeal SSBs (see Subheading 6 below and [172]), and to be involved in replicative and non-replicative DNA repair, and cell cycle checkpoints [115–118]. As expected, their N-terminal OB fold binds ssDNA and, although crystal structure analysis did not indicate that the C-terminal tail participates in this interaction [119] (Fig. 2b), its deletion decreases ssDNA binding affinity and protein stability of hSSB1 [120]. The available crystal structure was determined for a heterotrimeric complex containing hSSB1, INTS3, and C9orf80 [119], named sensor of singlestranded DNA complex 1 (SOSS1), which is recruited to DSBs in a Mre11/Rad50/Nbs1 (MRN) complex-dependent manner [115]. A second complex, named SOSS2, may also be formed with INTS3 and C9orf80 and recruited to DSBs, but in this case hSSB1 is replaced with hSSB2. SOSS2 may have a more tissuespecific role based on the restricted expression pattern of hSSB2 in immune cells and the testes [121]. hSSB1 also functions independently of SOSS1, as the protein is able to form homotetramers (and other oligomers) in solution under oxidative conditions [117, 122], which is a necessary structural arrangement for the protein to functionally interact with the human oxoguanine glycosylase hOGG1 in BER [116]. hSSB2 can also multimerize on ssDNA, a property that is dependent on its C-terminal tail [120], but that undoubtedly warrants further investigation. A lot more is known about the C-terminal tail of hSSB1, specifically the fact that this is the site for posttranslational phosphorylation and acetylation in the protein. Through the action of two important kinases (ATM and DNA-PK), phosphate groups can be attached to residues T117 and S134, respectively, to stabilize the protein and facilitate its function in ionizing radiation-induced DSB repair and in rescue of stalled replication forks [123– 125]. Acetylation of hSSB1 residue K94 has also been reported to

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stabilize the protein in response to DNA damage events following ionizing radiation exposure [126]. We expect to see an increasing number of studies detailing hSSB1 and hSSB2 function in all DNA repair pathways in the near future; however their roles in DNA replication per se will most likely remain absent. In DNA replication, RPA is still “the” eukaryotic SSB. 5.2 Mitochondrial SSB Proteins

Mitochondrial (mt) SSB proteins retain a homotetrameric organization (Fig. 2c), which is reminiscent of their bacterial evolutionary origin [127]. Structural aspects and the general mechanism of DNA binding by mtSSB proteins are also similar to those of bacterial SSB proteins [13]. In fact, residues of EcSSB relevant for DNA binding are conserved in the mtSSB proteins in humans, frogs, rats, flies, and lower metazoans [13, 128]. Also similar to EcSSB, the number of nucleotides occluded upon ssDNA binding depends on the ionic conditions and the protein-to-DNA ratio, such that at low salt concentrations and low relative SSB abundance, the number of nucleotides bound is about half of that bound at higher concentrations (~60) [17, 129, 130]. We and others have shown that mtSSB proteins stimulate the activity of DNA polymerase gamma (Pol γ) and the mtDNA helicase Twinkle [131–133]. The stimulatory effect is not species specific, and in the case of Pol γ-mtSSB interactions, it likely results from a specific organization of the ssDNA template by mtSSB [134], whereas physical interactions may still at least partially explain Twinkle-mtSSB interactions [173]. Notably, recent advances in the characterization of human mtSSB provided direct evidence that interconversion between the DNA-binding modes (i.e., 60 to 30 nt binding mode) warrants DNA synthesis [129]. The transition can be imposed on DNA-bound mtSSB proteins by progressing Pol γ, through repulsive electrostatic interactions [135]. A notable difference between the bacterial and mitochondrial SSB proteins is the lack of the C-terminal domain in the latter [14, 133]. Interestingly, all features provided by such domain in EcSSB appear to be missing in mtSSBs or have not yet been described: (1) mtSSB proteins exhibit low to no cooperative ssDNA binding when in the low nucleotide-binding mode, which supports the predicted role of the C-terminal domain in bacterial SSB binding cooperativity [17, 130], and (2) unlike EcSSB which physically interacts with many genome maintenance proteins via its C-terminal domain, direct binding of mtSSB proteins to other mitochondrial replicative factors has not been reported to date. The human mtSSB has a short, unstructured C-terminal tail instead, whose deletion grants the protein an increased ability to stimulate DNA synthesis by Pol γ on a singly primed ssDNA template [133]. Despite no reports of physical interactions, we have shown that the loops between the OB β-sheets and the

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capping α-helix play important roles in stimulating Pol γ and Twinkle [133], and speculate that these may partially provide the functions that the EcSSB C-terminal tail exerts in bacterial DNA replication. Recently, it has been shown that human mtSSB can be phosphorylated at residue Y73, which results in a decrease in mitochondrial DNA copy number. This is the first demonstration of a posttranslational regulatory mechanism acting on mtSSB function to ensure proper mitochondrial DNA levels [136].

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Archaeal SSB Proteins Archaeal SSB proteins are not as well studied as their bacterial and eukaryotic counterparts. In this section, we intend to provide only a brief overview of these proteins, and would like to refer the reader to the extensive review on archaeal SSBs presented in Chapter 2 of this volume [172]. SSB proteins appear to have evolved distinctively in the two main archaeal phyla: the Euryarchaeota phylum contains more eukaryotic-like RPA proteins, whereas the SSBs in the Crenarchaea phylum tend to have a domain organization that resembles that of the bacterial proteins [137, 138]. However, the OB folds of both euryarchaeal and crenarchaeal SSB proteins are structurally more similar to those of eukaryotic SSBs [139, 140]. SSB proteins of both archaeal subdivisions interact with DNA-processing protein factors. The heterotrimeric RPA-like SSB of Pyrococcus furiosus (Pf) co-precipitates with RadA from cell extracts, and stimulates its strand exchange activity in vitro, facilitating resolution of Holliday junction intermediates [141]. In addition, PfRPA may interact with other recombination proteins such as Hjc, and with DNA polymerases and primase [141]. Interactions with partner proteins of the crenarchaeal SSB proteins are often facilitated by a C-terminal extension, similarly to that of their bacterial homologs [28, 139]. The C-terminal tail of Sulfolobus solfataricus SSB, for example, promotes the interactions of this protein with the cognate RNA polymerase, stimulating its activity [142]. Archaea also has some unique cases of SSBs with unusual properties. In Nanoarchaeum equitans, the only known representative of the phylum Nanoarchaeota [143], the SSB protein (NeqSSB-like protein) is biologically active as a monomer with a single OB fold, resembling SSB-like proteins of some viruses. Although it displays ssDNA binding properties and 32% sequence similarity with the aforementioned S. solfataricus SSB protein, the NeqSSB-like protein also binds to mRNA and, surprisingly, various dsDNA forms, with no apparent structure-dependent preferences. With a binding site size of 7  1 nt and no salt-dependent binding mode transition [144], the NeqSSB-like protein probably binds

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ssDNA in a manner different than those shown for other SSB proteins. In Thermoproteales, a clade of hyperthermophilic Crenarchaea, 10 out of 11 representative species lack the canonical OB fold-containing SSB protein [145, 146]. Instead, they encode a distinct protein termed “ThermoDBP,” which binds specifically ssDNA with low sequence specificity. Crystal structure analysis revealed a unique fold for ssDNA binding, consisting of an extended cleft lined with hydrophobic phenylalanine residues and flanked by basic amino acids. Two ssDNA-binding domains are linked by a C-terminal leucine zipper dimerization domain [146]. The ThermoDBP proteins likely displaced the canonical SSBs during the diversification of the Thermoproteales, which is a highly unusual example of loss of a ubiquitous protein during evolution. Interestingly in Sulfolobus acidocaldarius, robust cell growth was not affected by deletion of the gene encoding the only canonical SSB protein, demonstrating that SSBs are not vital in some archaeal species [147]. This is highly surprising considering the vast number of biochemical and genetic studies that have demonstrated the importance of functional SSBs for all life forms. Nonetheless, we reckon that “essential” and “ubiquitous” are still the best adjectives to describe SSBs in the title of scientific articles, such as this one.

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Viral SSB Proteins The existence of viruses is contingent on the efficient replication of their genome in a relatively hostile environment inside the host cell. Mechanisms of viral genome replication are highly diverse and depend on the host proteome to a variable degree. In the context of SSB, viruses can be divided into those that rely on the host SSB proteins and those that encode and utilize their own SSB homolog. The former may be exemplified by the classical replication system of bacteriophage λ, which utilizes the bacterial host DNA replication factors almost entirely, including the SSB protein [148, 149]. Similarly, many viruses of eukaryotes “hijack” the host SSB proteins: the large antigen of the simian virus 40 (SV-40), the bovine papillomavirus E2 protein, and the Epstein–Barr virus EBNA-1 antigen all interact with human RPA [10]. In fact, RPA was initially identified because it is essential for SV-40 replication [150]. The gp2.5 protein of bacteriophage T7 typifies viral SSB proteins (Fig. 3a). The crystal structure of gp2.5 shows a dimer in which each monomer adopts the OB fold consisting of a fivestranded antiparallel β-barrel capped by an α-helix on one end (αA). This αA helix is uniquely longer than the α-helix in other homologous OB folds, and also interacts with the β-barrel differently [151, 152], which perhaps allows gp2.5 to selectively stimulate the activity of the T7 DNA polymerase [153]. Although there

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Fig. 3 Viral SSBs. Structural representations of the homodimeric gp2.5 of the bacteriophage T7 (a) and the monomeric gp32 of the bacteriophage T4 (b). Each subunit of T7 gp2.5 is represented with distinct colors. The structure of T4 gp32 comprises only the core domain (see text for details). The software Pymol (www.pymol. org) was used to analyze the structures deposited on PDB under the accession codes 1JE5 and 1GPC, respectively, and to create the figure

is little amino acid sequence similarity between gp2.5 and other SSB proteins, superposition of its OB fold structure with that of EcSSB and HsRPA70 showed that the aromatic residues that stack against the bases in ssDNA are conserved [151], which suggests that gp2.5 binds to ssDNA similarly to other SSB proteins [152]. Like the NeqSSB-like protein, gp2.5 monomers bind ~7 nucleotides of ssDNA in a noncooperative and salt-sensitive manner [154, 155]. Interestingly, the protein comprises an acidic 28-amino acid-long C-terminal tail, which functionally resembles that of bacterial SSBs. The tail is necessary for phage growth, and modulates the oligomerization of gp2.5 and its interaction with ssDNA, T7 DNA polymerase, and T7 helicase-primase [152, 156]. These interactions are essential for establishing the coordinated synthesis of leading and lagging DNA strands of the viral genome [157]. A large number of SSB proteins encoded in the genomes of other bacteriophages share significant similarity with gp2.5, including the distinctive αA-helix, important ssDNA-binding residues, and highly acidic C-terminus [152]. Interestingly, computational studies identified SSB-encoding genes homologous to gp2.5 in viral genomes of four out of five families of the eukaryote-infecting nucleocytoplasmic large DNA viruses (NCLDVs), which is consistent with the presumed bacteriophagal evolutionary origin of eukaryotic dsDNA viruses [158]. Viral SSB proteins in fact exhibit a large diversity of structure and function overall, some of which are monomeric in nature. The best known example of a monomeric viral SSB protein is gp32 of bacteriophage T4 (Fig. 3b), which is a 34 kDa, 301-amino acidlong, zinc metalloprotein with three functional domains: the

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N-terminal basic (B) domain (residues 1–21), the core domain (residues 22–254), and the C-terminal acidic (A) domain (255–301) [159]. The core domain contains the ssDNA-binding site with a single OB fold [1, 160]. The acidic C-terminus mediates critical interactions with cognate replisome proteins such as the Dda helicase and the gp43 polymerase [161, 162]. The B domain mediates gp32 monomer self-association in solution and the highly cooperative ssDNA-binding mode of the protein [159, 163, 164]. The gp5 protein of bacteriophage φ29 and ICP8 of the herpes simplex virus are other examples of monomeric viral SSB proteins, in prokaryotes and eukaryotes, respectively. Both SSBs also bind ssDNA in a cooperative fashion and efficiently stimulate the activity of their cognate polymerases [165, 166]. ICP8, however, is considerably larger than bacteriophagal SSB proteins (with 128 kDa in size) and can form double-helical filaments in the absence of DNA, which likely facilitates the formation of replication compartments by mediating interactions with other components of the replication machinery [167]. As in the archaeal Thermoproteales group, SSB-like proteins that lack OB fold have been identified in adenoviruses. The adenovirus type 5 DNA-binding protein (Ad DBP) is an ~59 kDa multifunctional protein essential for the virus replication, which binds ssDNA in a cooperative manner with a binding site size of ~13 nt [168]. The cooperative binding promotes template DNA unwinding in an ATP-independent fashion during elongation of the adenovirus DNA replication [169]. The protein contains two zinc atoms, which appear to be required for the stability of the protein fold, rather than being involved in direct contact with the DNA [170]. The crystal structure shows that the protein contains a 17-amino acid-long C-terminal extension, which hooks onto a second molecule, thereby forming a protein chain; deletion of this C-terminal tail decreases cooperative DNA binding [170, 171]. In addition, Ad DBP can also bind uncooperatively dsDNA and RNA [170]. Identification of SSB-like proteins with no apparent OB fold suggests distinct evolutionary origins and specialization through convergent evolution, which in turn emphasizes the essential role of this type of protein in genome maintenance and stability.

Acknowledgments G.L.C. was partially supported by a grant from the Auburn University at Montgomery Research Grant-in-Aid Program. M.T.O. would like to acknowledge funding from the Fundac¸˜ao de Amparo a` Pesquisa do Estado de Sa˜o Paulo (FAPESP, grant number 2017/04372-0) and the Conselho Nacional de Desenvolvimento Cientı´fico e Tecnolo´gico (CNPq, grant numbers 424562/ 2018-9 and 306974/2017-7).

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133. Oliveira MT, Kaguni LS (2011) Reduced stimulation of recombinant DNA polymerase γ and mitochondrial DNA (mtDNA) helicase by variants of mitochondrial single-stranded DNA-binding protein (mtSSB) correlates with defects in mtDNA replication in animal cells. J Biol Chem 286:40649–40658 134. Ciesielski GL, Bermek O, Rosado-Ruiz FA, Hovde SL, Neitzke OJ, Griffith JD, Kaguni LS (2015) Mitochondrial single-stranded DNA-binding proteins stimulate the activity of DNA polymerase γ by organization of the template DNA. J Biol Chem 290:28697–28707 135. Cerro´n F, de Lorenzo S, Lemishko KM, Ciesielski GL, Kaguni LS, Cao FJ, Ibarra B (2019) Replicative DNA polymerases promote active displacement of SSB proteins during lagging strand synthesis. Nucleic Acids Res 47:5723–5734 136. Djeungoue-Petga MA, Lurette O, Jean S, Hamel-Coˆte´ G, Martı´n-Jime´nez R, Bou M, Cannich A, Roy P, Hebert-Chatelain E (2019) Intramitochondrial Src kinase links mitochondrial dysfunctions and aggressiveness of breast cancer cells. Cell Death Dis 10:940 137. Che´din F, Seitz EM, Kowalczykowski SC (1998) Novel homologs of replication protein A in archaea: implications for the evolution of ssDNA-binding proteins. Trends Biochem Sci 23:273–277 138. Wadsworth RI, White MF (2001) Identification and properties of the crenarchaeal singlestranded DNA binding protein from Sulfolobus solfataricus. Nucleic Acids Res 29:914–920 139. Kerr ID, Wadsworth RI, Cubeddu L, Blankenfeldt W, Naismith JH, White MF (2003) Insights into ssDNA recognition by the OB fold from a structural and thermodynamic study of Sulfolobus SSB protein. EMBO J 22:2561–2570 140. Kelly TJ, Simancek P, Brush GS (1998) Identification and characterization of a singlestranded DNA-binding protein from the archaeon Methanococcus jannaschii. Proc Natl Acad Sci U S A 95:14634–14639 141. Komori K, Ishino Y (2001) Replication protein A in Pyrococcus furiosus is involved in homologous DNA recombination. J Biol Chem 276:25654–25660 142. Richard DJ, Bell SD, White MF (2004) Physical and functional interaction of the archaeal single-stranded DNA-binding protein SSB with RNA polymerase. Nucleic Acids Res 32:1065–1074

143. Huber H, Hohn MJ, Rachel R, Fuchs T, Wimmer VC, Stetter KO (2002) A new phylum of Archaea represented by a nanosized hyperthermophilic symbiont. Nature 417:63–67 144. Olszewski M, Balsewicz J, Nowak M, Maciejewska N, Cyranka-Czaja A, ZalewskaPia˛tek B, Pia˛tek R, Kur J (2015) Characterization of a single-stranded DNA-binding-like protein from Nanoarchaeum equitans—a nucleic acid binding protein with broad substrate specificity. PLoS One 10:e0126563 145. Luo X, Schwarz-Linek U, Botting CH, Hensel R, Siebers B, White MF (2007) CC1, a novel crenarchaeal DNA binding protein. J Bacteriol 189:403–409 146. Paytubi S, McMahon SA, Graham S, Liu H, Botting CH, Makarova KS, Koonin EV, Naismith JH, White MF (2012) Displacement of the canonical single-stranded DNA-binding protein in the Thermoproteales. Proc Natl Acad Sci U S A 109:E398–E405 147. Suzuki S, Kurosawa N (2019) Robust growth of archaeal cells lacking a canonical singlestranded DNA-binding protein. FEMS Microbiol Lett 366:fnz124 148. Wold MS, Mallory JB, Roberts JD, LeBowitz JH, McMacken R (1982) Initiation of bacteriophage lambda DNA replication in vitro with purified lambda replication proteins. Proc Natl Acad Sci U S A 79:6176–6180 149. Dodson M, McMacken R, Echols H (1989) Specialized nucleoprotein structures at the origin of replication of bacteriophage lambda. Protein association and disassociation reactions responsible for localized initiation of replication. J Biol Chem 264:10719–10725 150. Borowiec JA, Dean FB, Bullock PA, Hurwitz J (1990) Binding and unwinding—how T antigen engages the SV40 origin of DNA replication. Cell 60:181–184 151. Hollis T, Stattel JM, Walther DS, Richardson CC, Ellenberger T (2001) Structure of the gene 2.5 protein, a single-stranded DNA binding protein encoded by bacteriophage T7. Proc Natl Acad Sci U S A 98:9557–9562 152. Hernandez AJ, Richardson CC (2019) Gp2.5, the multifunctional bacteriophage T7 single-stranded DNA binding protein. Semin Cell Dev Biol 86:92–101 153. Nakai H, Richardson CC (1988) The effect of the T7 and Escherichia coli DNA-binding proteins at the replication fork of bacteriophage T7. J Biol Chem 263:9831–9839 154. Kim YT, Tabor S, Bortner C, Griffith JD, Richardson CC (1992) Purification and

The Essential, Ubiquitous Single-Stranded DNA-Binding Proteins characterization of the bacteriophage T7 gene 2.5 protein. A single-stranded DNA-binding protein. J Biol Chem 267:15022–15031 155. Shokri L, Rouzina I, Williams MC (2009) Interaction of bacteriophage T4 and T7 single-stranded DNA-binding proteins with DNA. Phys Biol 6:025002 156. Lee SJ, Marintcheva B, Hamdan SM, Richardson CC (2006) The C-terminal residues of bacteriophage T7 gene 4 helicaseprimase coordinate helicase and DNA polymerase activities. J Biol Chem 281:25841–25849 157. Lee SJ, Richardson CC (2011) Choreography of bacteriophage T7 DNA replication. Curr Opin Chem Biol 15:580–586 158. Koonin EV, Yutin N (2010) Origin and evolution of eukaryotic large nucleo-cytoplasmic DNA viruses. Intervirology 53:284–292 159. Williams KR, Sillerud LO, Schafer DE, Konigsberg WH (1979) DNA binding properties of the T4 DNA helix-destabilizing protein. A calorimetric study. J Biol Chem 254:6426–6432 160. Shamoo Y, Friedman AM, Parsons MR, Konigsberg WH, Steitz TA (1995) Crystal structure of a replication fork single-stranded DNA binding protein (T4 gp32) complexed to DNA. Nature 376:362–366 161. Burke RL, Alberts BM, Hosoda J (1980) Proteolytic removal of the COOH terminus of the T4 gene 32 helix-destabilizing protein alters the T4 in vitro replication complex. J Biol Chem 255:11484–11493 162. Krassa KB, Green LS, Gold L (1991) Proteinprotein interactions with the acidic COOH terminus of the single-stranded DNA-binding protein of the bacteriophage T4. Proc Natl Acad Sci U S A 88:4010–4014 163. Giedroc DP, Khan R, Barnhart K (1990) Overexpression, purification, and characterization of recombinant T4 gene 32 protein 22-301 (g32P-B). J Biol Chem 265:11444–11455 164. Casas-Finet JR, Fischer KR, Karpel RL (1992) Structural basis for the nucleic acid binding cooperativity of bacteriophage T4 gene 32 protein: the (Lys/Arg)3(Ser/Thr) 2 (LAST) motif. Proc Natl Acad Sci U S A 89:1050–1054

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Chapter 2 Single-Stranded DNA-Binding Proteins in the Archaea Najwa Taib, Simonetta Gribaldo, and Stuart A. MacNeill Abstract Single-stranded (ss) DNA-binding proteins are found in all three domains of life where they play vital roles in nearly all aspects of DNA metabolism by binding to and stabilizing exposed ssDNA and acting as platforms onto which DNA-processing activities can assemble. The ssDNA-binding factors SSB and RPA are extremely well conserved across bacteria and eukaryotes, respectively, and comprise one or more OB-fold ssDNA-binding domains. In the third domain of life, the archaea, multiple types of ssDNAbinding protein are found with a variety of domain architectures and subunit compositions, with OB-fold ssDNA-binding domains being a characteristic of most, but not all. This chapter summarizes current knowledge of the distribution, structure, and biological function of the archaeal ssDNA-binding factors, highlighting key features shared between clades and those that distinguish the proteins of different clades from one another. The likely cellular functions of the proteins are discussed and gaps in current knowledge identified. Key words Archaea, SSB, RPA, OB fold, Phylogenetics, Replication protein A, Single-stranded DNA, ssDNA, DNA replication, DNA repair

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Introduction Single-stranded DNA-binding proteins play essential roles in almost all aspects of DNA metabolism in all three domains of life: bacteria, eukaryotes, and archaea [1]. In bacteria, the major singlestranded DNA-binding protein is SSB, whereas in eukaryotes, this role is performed by the trimeric replication protein A (RPA) complex. SSB and RPA are characterized by the presence of one or more OB (oligosaccharide-oligonucleotide binding)-fold domains [2]. OB-fold domains span 75–150 amino acids and consist of a five-stranded β-sheet that is coiled to form a closed β-barrel structure, often capped by an α-helix. The heterogeneity in length reflects the presence of variable loop regions generally located between the β-strands. Single-stranded DNA binding by the SSB and RPA OB folds is not sequence specific, reflecting the need for the SSB/RPA proteins to engage, transiently, with single-stranded DNA in a wide variety of functional contexts [2].

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_2, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Bacterial SSB. (a) Domain organization of the archetypal E. coli SSB protein. (b) Crystal structure of the SSB tetramer bound to single-stranded DNA (PDB: 1EYG) [63]. The structure was obtained using SSB protein lacking the unstructured C-terminal tail (amino acids 1–116 of 178)

In bacteria, the archetypal Escherichia coli SSB is a 255-amino acid protein that comprises a single N-terminal OB-fold domain followed by a relatively short, flexible C-terminal tail [3] that mediates protein-protein interactions (Fig. 1). Individual bacterial SSB proteins of this type (i.e., with a single OB-fold domain per protein) assemble into homotetrameric complexes. In contrast to the situation in E. coli, in certain other bacterial lineages (Deinococcus and Thermus being the best studied) SSB proteins with a tandem pair of OB-fold domains are found [4]. The second OB-fold domain is followed by a flexible C-terminal tail related in sequence to that of E. coli SSB. These proteins assemble to form homodimers that, like E. coli SSB, contain four OB-fold domains in total [5]. In eukaryotes, replication protein (RPA) is the major singlestranded DNA-binding factor [6, 7]. RPA is a heterotrimeric complex made up of the RPA1, RPA2, and RPA3 proteins (also known as RPA70, RPA32, and RPA14, respectively). Figure 2a shows the domain organization of the well-studied RPA from the budding yeast Saccharomyces cerevisiae. RPA1 contains four OB-fold domains (designated DBD-A, DBD-B, DBD-C, and DBD-F, where DBD is short for DNA-binding domain), RPA2 contains one OB fold (DBD-D), and RPA3 also one OB fold (DBD-E). DNA-binding domains DBD-A, DBD-B, DBD-C, and DBD-D are primarily responsible for ssDNA binding by RPA. DBD-E plays a structural role at the heart of the RPA complex while

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Fig. 2 Eukaryotic RPA. (a) Domain organism of budding yeast S. cerevisiae RPA proteins RPA1 (RPA70), RPA2 (RPA32), and RPA3 (RPA14) with OB-fold motifs labeled DBD-A through DBD-F and RPA2 C-terminal winged helix domain labeled wH. (b) Structure of the fungus U. maydis RPA1 DBD-A, DBD-B, and DBD-C, RPA2 DBD-D, and RPA3 DBD-E bound to single-stranded DNA (PDB: 4GNX) [9]

DBD-F functions as a site of protein-protein interaction. RPA2 also possesses an extended N-terminal domain that is the target for regulatory phosphorylation and a C-terminal winged helix (wH) domain, both of which are also involved in protein-protein interactions [8]. A number of partial RPA structures have been solved, both with and without bound ssDNA, such as the Ustilago maydis DBD-A/B/C/D/E structure shown in Fig. 2b [9]. RPA is essential for eukaryotic chromosome replication and plays important roles in a variety of DNA repair processes [6, 7]. In addition to RPA, variant RPA-like complexes are found in a number of lineages [10–12], including mammals and plants, and the related CST (Cdc13-Stn1-Ten1) complex plays an important role in telomere maintenance [13]. Single OB-fold SSBs (hSSB1 and hSSB2) have also been identified and characterized in mammals and shown to have roles in DNA damage repair [14–16]. This chapter provides an overview of the current state of knowledge of the biology of single-stranded DNA-binding proteins in the archaea. Archaeal organisms make up the third domain of life on Earth, are ubiquitous in nature, make up an estimated 20% of

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the planet’s biomass, and play important roles in biosphere and atmosphere. OB fold-containing RPA- and/or SSB-like ssDNAbinding factors have been identified in a number of well-studied archaeal species, with the RPA-like proteins displaying a range of domain architectures with varying numbers of OB folds (Figs. 3 and 4). Biochemical characterization of these proteins has allowed conclusions to be drawn regarding their ssDNA-binding properties in vitro, while genetic analysis has inferred vital roles for the proteins in archaeal DNA replication and repair in vivo. Here, we present an up-to-date phylogenetic analysis of the distribution of RPA/SSB proteins across the archaeal domain and provide a comprehensive review of current knowledge of the properties of these proteins. Taken together, the data provides an excellent foundation for further studies of the biology of ssDNA binding in the archaea.

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Phylogenetic Distribution of Archaeal SSB/RPA Proteins Figure 3 illustrates the phylogenetic distribution of OB foldcontaining SSB/RPA proteins across the archaea [17] and provides a number of important insights into single-stranded DNA-binding capacity in different lineages. Individual SSB/RPA proteins can be classified into four groups (shown in schematic form in Fig. 3a): RPA41 and RPA32 proteins, readily recognizable by the presence of characteristic C-terminal zinc finger OB fold and winged helixturn-helix (wH) domains, respectively; single OB-fold proteins, including both RPA14 and SSB proteins; and a broad class of multiple OB-fold RPA proteins containing neither a zinc finger nor a wH domain. The archaeal RPA41 proteins are structurally and evolutionarily related to eukaryotic RPA1 and are defined by the presence of one or several OB folds and a C-terminal C4 or C3H zinc finger motif. The first archaeal RPA to be characterized, from Methanocaldococcus jannaschii [18], was a member of this group and was originally reported to comprise four tandem OB-fold domains followed by a C4 zinc finger motif [18, 19]. More recent analysis has shown that the zinc finger is in fact embedded in a fifth OB fold that is related to the DBD-C OB fold in eukaryotic RPA1 (SM, unpublished). The presence of a DBD-C-like OB fold is a defining characteristic of all archaeal RPA41 proteins. To date, RPA41 proteins have been found to be encoded by almost all archaeal lineages, with exceptions being the Micrarchaeota, the closely related Nanoarchaeota and Parvarchaeota, and almost all lineages within the TACK superphylum (originally named for the Thaumarchaeota, Aigarchaeota, Crenarchaeota, and Korarchaeota) (Fig. 3b). The archaeal RPA32 proteins are structurally and evolutionarily related to eukaryotic RPA2 and are defined by a structure that comprises one or two OB folds followed by the winged helix-

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Fig. 3 Domain structures and phylogenetic distribution of archaeal SSB/RPA proteins. (a) Schematic showing representative structures of various archaeal SSB/RPA proteins. The numbers of OB folds in the RPA41,

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turn-helix (wH) domain. Very few of these proteins have been characterized biochemically and in those that have, the presence of the C-terminal wH domain and the similarity to RPA2 went entirely unnoticed [20, 21]. Like the RPA41 proteins, RPA32 proteins are widespread across the archaea, although it is noticeable that a small number of lineages that encode RPA41 do not encode RPA32 and that an even smaller number of lineages (specifically, the Nanoarchaeota and Parvarchaeota) appear to encode an RPA32 but not an RPA41 (Fig. 3b). These observations suggest that the RPA41 and RPA32 proteins are each capable of functioning independently as single-stranded DNA-binding proteins, even if they are likely to form dimeric RPA41-RPA32 complexes in most species (discussed further below). Single OB-fold proteins are also widespread and fall into at least two groups (Fig. 3). A number of species of the Thermococcales have been shown to encode trimeric RPA complexes that are very similar in organization to eukaryotic RPA (see Subheading 3.4 below). In these cases, the RPA41, RPA14, and RPA32 proteins (akin to eukaryotic RPA1, RPA3, and RPA2, respectively) are encoded by adjacent genes and co-transcribed [22, 23]. This genetic organization is seen in the Theionarchaea and in some species of Methanococcales (Subheading 3.1.1), suggesting that these organisms also encode a trimeric RPA (Fig. 3). However, in addition to this, many archaeal lineages carry genes that encode single OB-fold proteins that are not linked genetically to those encoding RPA41 and RPA32 (and where the genes encoding RPA41 and RPA32, if both are present, are not necessarily linked to one another). In the absence of biochemical evidence, it is not possible to be determined with certainty if these single OB-fold proteins are the RPA14 component of a trimeric RPA, or if they act independently as monomeric or possibly homomultimeric SSBs. In the Crenarchaeota, which lack both RPA41 and RPA32, the single OB-fold SSB protein has been extensively characterized (see Subheading 4). SSB proteins of this type appear to be a shared feature of the TACK archaea (with the exception of the Korarchaeota) that lack RPA41 and RPA32. The final type of archaeal RPA proteins possess multiple OB folds and can be distinguished from the RPA41 and RPA32 proteins by the absence of a C-terminal DBD-C-like zinc fingerä Fig. 3 (continued) RPA32, and multiple OB-fold RPA proteins vary from lineage to lineage. (b) Taxonomic distribution and diversity of SSB/RPA across the archaeal domain. HMM profiles were built for each component using reference sequences and searched against a local data bank of 258 archaeal genomes using HMMSEARCH. The hits were manually checked using domain composition and structure prediction. The presence/absence in each phylum is indicated using a color gradient according to the number of occurrences within the group. White is complete absence and dark red is presence in all the genomes. The full dataset can be found in ref. [17]

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containing OB fold or a winged helix (wH) domain, respectively. These proteins are scattered across a range of species (Fig. 3), with no obvious pattern in relation to the presence or absence of the other three types of RPA/SSB (RPA41, RPA32, or single OB-fold proteins). The following sections summarize current knowledge of the biochemical properties and biological function of studied archaeal RPA/SSB proteins. With one exception (see Subheading 3.1.1), we refer to these proteins by their original given names (for example MthRPA or HvoRpap1) rather than renaming them to take into account their newfound relatedness to RPA41 or RPA32.

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Euryarchaeal RPA Proteins

3.1 RPA Proteins in Class I Methanogens

The Euryarchaeota represent a broad range of archaeal species with highly diverse characteristics, including methanogens, thermophiles, and halophiles (Fig. 3a). The class I methanogens form the Methanomada superclass [24]. These organisms are obligate anaerobes. RPA proteins from three different class I methanogens, representing three different taxonomic groupings, have been characterized biochemically: Methanocaldococcus jannaschii (formerly Methanococcus jannaschii), a member of the order Methanococcales and the first archaeal organism to have its genome completely sequenced [25]; Methanothermobacter thermautotrophicus (formerly Methanobacterium thermoautotrophicum), a member of the Methanobacteriales; and Methanopyrus kandleri, a member of the Methanopyrales. Although broadly similar in structure, these RPA proteins display 100  C [29]. MkaRPA is a 432-amino acid RPA41-type protein comprising three OB folds, with the most C-terminal being of the eukaryotic RPA1 DBD-C type and containing an integral zinc finger (see Fig. 4) (SM, unpublished). MkaRPA has been shown to bind single-stranded DNA with high affinity in electrophoretic mobility shift assays (EMSAs), with FRET analysis suggesting two different binding modes (compacting then stretching) as the protein-to-DNA concentration increases. Gel filtration analysis of recombinant MkaRPA indicates that the protein is a trimer in solution. Similar to the situation with MthRPA, full-length MkaRPA (but not a C-terminally truncated protein that removes the DBD-C-like OB fold) inhibits the primer extension activity of the M. kandleri family B DNA polymerase MkaPolBI in vitro [29]. As in the case of M. jannaschii, M. kandleri encodes an RPA32-type protein [17] that may interact with MkaRPA to form a dimeric complex in vivo but which has not been studied.

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3.2 RPA Proteins in the Class II Methanogens

The class II methanogens form the Methanomicrobia class and are phylogenetically distinct from the class I methanogens such as M. jannaschii and M. thermautotrophicus [24]. Indeed, the class II methanogens are more closely related to the non-methanogenic Halobacteria (which, despite their name, are not bacteria, but archaea), something that is borne out by consideration of their RPA proteins. To date, with the exception of a single structural study, investigation of the properties of these proteins has been confined to a single representative species, Methanosarcina acetivorans [29–33].

3.2.1 Methanosarcina acetivorans

M. acetivorans encodes three RPA homologs designated MacRPA1, MacRPA2, and MacRPA3 [29–31]. MacRPA1 is a 484-amino acid protein containing four tandem OB-fold domains, but no zinc finger motif (Fig. 4), and is therefore related to the Haloferax volcanii Rpa2 protein and Halobacterium salinarum Rfa1 proteins discussed below, with the absence of the zinc finger being a shared feature (see Subheading 3.3). Interestingly, the RPA1 protein of the closely related species Methanococcoides burtonii (MbuRPA1) contains only three, not four, OB-fold domains [33]. Multiple sequence alignments point to the protein having arisen due to deletion in sequences encoding the second and third OB folds of a MacRPA1-like ancestral RPA with four OB folds. Biochemical analysis suggests that recombinant MacRPA1 can exist as both a dimer and a tetramer in solution [30]. Deletion of either one or two OB folds from the N-terminal or C-terminal end of the MacRPA1 protein reduces, to varying degrees, but does not abolish, ssDNA binding in EMSA and/or fluorescence polarization assays, while removal of a third OB fold (leaving only the most N-terminal or most C-terminal fold, OB1 and OB4, in Fig. 4, respectively) abolishes ssDNA binding altogether. There is clearly a degree of redundancy among the OB folds in these in vitro assays, something that is borne out by in vivo assays of HvoRpa2 function in H. volcanii (discussed in Subheading 3.3.1 below). In vitro, MacRPA1 is capable of inhibiting the activity of the M. acetivorans flap endonuclease (Fen1) homolog MacFEN1 to cleave 20-nucleotide 50 -flap structures but cannot inhibit cleavage of 5-nucleotide flaps [32]. The ability to inhibit MacFEN1 activity is not shared by truncated MacRPA1 proteins lacking more than just the N- or C-terminal OB fold, or by MacRPA2 or MacRPA3. MacRPA2 and MacRPA3 are RPA41-type proteins are 417 and 450 amino acids in length and closely related to one another. Each protein comprises an N-terminal tandem pair of OB-fold domains followed by a C-terminal DBD-C-type OB fold with an integral C3H zinc finger (Fig. 4) (SM, unpublished). There is no structural information for either of the MacRPA2 or MacRPA3 proteins, but the structure of the first OB-fold domain of the M. mazei RPA2

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protein (MmaRPA2), which is 96% identical to MacRPA2, has been determined by NMR (Fig. 3c, Table 1). Recombinant MacRPA2 and MacRPA3 have been reported to exist as dimers in solution, to contain bound zinc, as would be expected, and to specifically bind ssDNA over dsDNA, with MacRPA3’s affinity for ssDNA being greater than that of MacRPA2 [30]. Both proteins, as well as MacRPA1, are capable of stimulating primer extension by M. acetivorans DNA polymerase B (MthPolB). Mutating individual cysteines in the MacRPA3 DBD-C zinc finger to alanine reduces single-stranded DNA binding and the ability of the proteins to stimulate MacPolBI activity in vitro but also causes significant structural changes [31], while removing the entire DBD-C region does not greatly alter ssDNA binding [29, 30]. Although there is abundant published information regarding the biochemical properties of recombinant MacRPA2 and MacRPA3 proteins in isolation, M. acetivorans also encodes unstudied RPA32 proteins [17] and it is highly likely that MacRPA2 and MacRPA3 form heterodimeric RPA41-RPA32 complexes in vivo. As a result, as with M. jannaschii and M. kandleri, discussed above, the earlier biochemical data obtained with recombinant MacRPA2 and MacRPA3 in isolation should be treated with a degree of caution when considering the in vivo behavior of these proteins. 3.3 RPA Proteins in the Halobacteriales

The Halobacteria are relatively closely related to Methanosarcinales such as M. acetivorans, M. burtonii, and M. mazei (Fig. 3) and comprise a range of euryarchaeal organisms that inhabit hypersaline environments such as salt lakes and solar salterns and which have found an important place as model systems in archaeal research due to ease with which they can be grown in the lab and their tractability to molecular genetic analysis [34].

3.3.1 Haloferax volcanii

There is substantial similarity between the RPA proteins encoded by the Halobacteriales and the Methanosarcinales. The H. volcanii Rpa1 and Rpa3 proteins (hereafter referred to as HvoRpa1 and HvoRpa3, also known as RpaA1 and RpaB1) are RPA41-type proteins and homologs of M. acetivorans RPA2 (MacRPA2) and RPA3 (MacRPA3), respectively, while Rpa2 (HvoRpa2) is a homolog of M. acetivorans RPA1 (MacRPA1). HvoRpa1 is a 427-amino acid protein that comprises three putative OB folds, with the most C-terminal being the eukaryotic DBD-C type with integral C3H zinc finger, while at 311 amino acids, HvoRpa3 is somewhat shorter, comprising only two OB folds, the second being the DBD-C type with C3H zinc finger (SM, unpublished) (Fig. 4). At the chromosome level, the rpa1 gene overlaps with rpap1, which encodes a 623-amino acid RPA32 protein containing an N-terminal OB fold and a C-terminal winged helix (wH) domain, while rpa3 is located adjacent to rpap3, which

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encodes a 196-amino acid RPA32 protein with an N-terminal OB fold and a C-terminal wH domain [17]. Consistent with their classification as RPA41 and RPA32 proteins, HvoRpa1 and HvoRpap1 can be co-purified from native extracts prepared from overexpressing cells, as can HvoRpa3 and HvoRpap3 [21]. There is no evidence for the existence of an RPA14 protein associated with these RPA41-RPA32 complexes, as is the case in P. furiosus and T. kodakarensis (see Subheading 3.4 below), and the current assumption is that the H. volcanii RPAs are heterodimers. No structural information is available for any part of any of the four proteins that make up the two heterodimers, but HvoRpa3—in isolation, i.e., without the corresponding RPA32 protein HvoRpap3—has been purified in recombinant form and shown to be able to bind DNA in high-salt conditions mimicking the internal salt concentrations of the H. volcanii cell [35]. The N-terminal OB fold alone has reduced affinity for ssDNA compared to the full-length HvoRpa3 protein, whereas the C-terminal DBD-C type OB fold did not bind ssDNA under the conditions used. Reverse genetic analysis has allowed initial insight into the cellular functions of the HvoRpa1-HvoRpap1 and HvoRpa3HvoRpap3 complexes [20, 21]. The rpa1-rpap1 and rpa3-rpap3 operons can be deleted from the chromosome individually, but not at the same time, indicating that HvoRpa1-HvoRpap1 and HvoRpa3-HvoRpap3 share an essential function in the cell. Consistent with this, repression of rpa3-rpap3 expression in Δrpa1rpap1 cells leads to a significant growth retardation [20]. Cells lacking the HvoRpa3-HvoRpap3 complex (Δrpa3-rpap3) display increased sensitivity to UV light and to the DNA-damaging agent mitomycin C, while Δrpa1-rpap1 cells lacking HvoRpa1HvoRpap1 retain full repair capacity [21]. UV exposure causes formation of cyclobutane pyrimidine dimers (CPDs) and pyrimidine 6–4 photoproducts (6-4PPs) in DNA, whereas mitomycin C (MMC) produces three types of DNA damage: monoadducts, and intra- and inter-strand cross-links. The observed sensitivity of Δrpa3-rpap3 cells is therefore indicative of the HvoRpa3HvoRpap3 complex playing a role in ssDNA binding during the repair of these DNA lesions, even if the molecular mechanisms of UV and MMC damage repair in H. volcanii are not fully understood, while the shared essentiality of the HvoRpa1-HvoRpap1 and HvoRpa3-HvoRpap3 complexes most likely implies a role in chromosomal DNA replication. The H. volcanii Rpa2 protein (HvoRpa2, also known as RpaC) is an ortholog of M. acetivorans RPA1 (MacRPA1) (Fig. 4). HvoRpa2 is a 483-amino acid protein and like MacRPA1 comprises a short, conserved N-terminal domain, followed by four OB-fold domains, the most C-terminal of which was not immediately apparent at the time of initial publication, but no zinc finger [20]. No structural information is available for any part of HvoRpa2.

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Using reverse genetic methodology, HvoRpa2 has been shown to be essential for cell viability: the rpa2 gene cannot be deleted from the chromosome and downregulation of rpa2 expression in otherwise wild-type cells results in significant growth retardation [20]. Interestingly, this slow-growth phenotype can be partially rescued by increased expression of rpa3-rpap3, though not rpa1rpap1, indicating that HvoRpa3-HvoRpap3 can partly substitute for Rpa2 in vivo. Overexpression of HvoRpa2 leads to enhanced resistance to DNA damage caused by exposure to UV light, 4NQO (4-nitroquinoline 1-oxide), the alkylating agent MMS, and the antibiotic phleomycin, which causes DNA strand breaks. The Rpa2 protein has been used as a platform for in vivo analysis of the role of individual OB-fold domains, with various truncation and deletion derivatives of the Halogeometricum borinquense Rpa2 protein (HgmRpa2) being tested for their ability to rescue the slow growth defect resulting from repression of H. volcanii rpa2 expression [20]. The HgmRpa2 protein is 75% identical to HvoRpa2 and is able to rescue to near-wild-type levels. Deletion of the C-terminal OB fold (OB4 in HvoRpa2 in Fig. 3) did not affect the ability of the HgmRpa2 protein to rescue the growth defect in the absence of exogenous DNA damage, but did result in increased sensitivity to UV irradiation and MMS exposure. Deletion of OB folds OB1 and OB2 led to somewhat reduced ability to rescue the growth defect, while deletion of OB3 significantly impaired rescue ability. Some rescue was seen when OB1 and OB2 were deleted together, but none when OB1 and OB3, or OB2 and OB3, were deleted. Taken together, these results indicate a particularly important role for OB3 in Rpa2 function, assisted by OB1, OB2, and OB4 [20]. It remains to be seen how these genetic observations correlate with the actual ssDNA-binding dynamics. To gain further insights into the biological role of the essential HvoRpa2 protein, H. volcanii cells have been engineered to express a GFP-Rpa2 fusion protein from the native chromosomal locus, with the gfp:rpa2 open reading frame replacing the endogenous rpa2 gene [36]. Microscopic examination of either exponentially growing or stationary-phase cells reveals that the GFP-Rpa2 protein is found at a small number of discrete foci, even in the absence of exogenous DNA damage, with the fact that slightly more foci are observed in exponentially growing cells leading to the suggestion that at least some of these may correspond to activated DNA replication forks [36]. UV treatment of cells leads to a marked reduction in the number of GFP-Rpa2 foci, with single large foci being observed in cells treated with higher UV doses. In contrast, treatment of cells with the DNA polymerase inhibitor aphidicolin leads to a doubling in the number of GFP-Rpa2 foci, while treatment with the DNA-damaging agent phleomycin leads to an approximate fivefold increase. It is assumed that the patterns observed with different damaging agents reflect the amount of ssDNA available to be bound by HvoRpa2 at each dose [36].

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3.3.2 Halobacterium salinarum

RPA function has also been investigated in the closely related organism Halobacterium salinarum (also known as Halobacterium sp. NRC-1). H. salinarum encodes homologs of HvoRpa1, HvoRpap1, HvoRpa3, and HvoRpap3, designated HsaRfa2, HsaRfa7, HsaRfa3, and HsaRfa8, respectively, as well as an HvoRpa2 homolog, HsaRpa1. The proteins are very similar in size and composition across the species, with the exception of the RPA32 protein HsaRfa7 which is ~160 amino acids shorter in its central section (between the OB-fold and the wH domain) than its H. volcanii counterpart HvoRpap1 (Fig. 3). Biochemical analysis shows that both HsaRfa3 and HsaRfa8 can be purified on the basis of one or other of the proteins, or both, having affinity for ssDNA, but falls short of proving that the two proteins interact with one another, as would be expected by analogy with HvoRpa3 and HvoRpap3 in H. volcanii or RPA41-RPA32 more generally [37]. A number of studies have shown that the rfa3-rfa8 operon is upregulated in response to DNA damage caused by ionizing radiation (IR) and UV exposure and that this results in a concomitant increase in Rfa3 and Rfa8 protein levels [38–40]. Upregulation is also seen in H. salinarum strains selected for their enhanced resistance to IR [37, 39, 41] and has subsequently been shown to be sufficient in itself for increased IR resistance [37]. Gene deletion analysis [42] has further underlined the importance of the Rfa3 and Rfa8 proteins in H. salinarum, as strains deleted for either rpa3 or rpa8 are sensitive to both IR and UV exposure, with the UV sensitivity mirroring what is seen with Δrpa3-rpap3 strains in H. volcanii (where IR sensitivity has not been tested) [21]. In contrast to the situation in H. volcanii, deletion of the H. salinarum rfa1 gene, encoding the homolog of the essential HvoRpa2 protein, is viable, although slow growing and severely sensitive to ionizing radiation, UV, and mitomycin C, while deletion of rfa2, but not rfa7, appears to be lethal, despite the products of these genes likely forming a dimeric complex. This raises the possibility that Rfa2 may have a function distinct from that of the presumed Rfa2-Rfa7 complex. In support of this notion, rfa2 expression is markedly upregulated in response to UV and MMC treatment whereas rfa7 expression is not [42]. Further analysis will be required to confirm whether this is the case.

3.4 RPA Proteins in the Thermococcales

RPA proteins have been characterized from two well-studied representatives of the Thermococcales, P. furiosus [22] and T. kodakarensis [23]. Both organisms encode RPA41, RPA14, and RPA32 proteins that form a heterotrimeric RPA complex, similar to eukaryotic RPA. In P. furiosus, the RPA41 protein is 360 amino acids in length and contains two OB folds, the most C-terminal of which is of the DBD-C type with an integral C3H zinc finger (Fig. 4) (SM, unpublished). The RPA32 protein is 273 amino acids in length

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and contains an N-terminal OB fold and a C-terminal wH domain, while the RPA14 protein is 122 amino acids in length and contains a single OB fold (Fig. 4). The T. kodakarensis proteins have identical domain organization and share 60–70% sequence identity with the P. furiosus archetypes. In both species, the three proteins are encoded from a single operon, with the genes arranged in the order rpa41-rpa14-rpa32 [22, 23]. Biochemically, PfuRPA has been shown to bind ssDNA with high affinity and specificity, to stimulate RadA-mediated strand exchange in vitro, and to co-immunoprecipitate from P. furiosus cell extracts with PfuRadA, mimicking the RPA-Rad51 interaction seen in eukaryotes [22]. Weaker interactions with other recombination and replication proteins, such as the Holliday junction resolvase Hjc, replication factor C, and DNA polymerase PolD, were also seen. In the related species P. abyssi, RPA was also reported to interact on ssDNA with PolD and primase, and to interact with and stimulate RNA polymerase [43]. TkoRPA has also been shown to bind ssDNA with high affinity and specificity, and to be able to relieve DNA polymerase pausing in in vitro reactions by resolving DNA template secondary structure while at the same time reducing DNA polymerase processivity [23]. TkoRPA is also reported to be able to interact in solution with both T. kodakarensis DNA polymerases TkoPolB and TkoPolD. Like PfuRPA and PabRPA, TkoRPA may also interact with primase, TkoRadA, TkoRad50, and reverse gyrase [23, 44]. 3.5 RPA Proteins in the Thermoplasmatales

Ferroplasma acidarmanus is a representative of the Thermoplasmatales, a clade of acidophilic Euryarchaea. F. acidarmanus was originally reported to encode two OB fold-containing ssDNA-binding proteins, designated FacRPA1 and FacRPA2 [29]. FacRPA1 is a 369-amino acid RPA41 protein possessing three OB folds, the third of which is of the DBD-C type with integral zinc finger, while FacRPA2 resembles a bacterial SSB protein, with a single OB fold and short C-terminal tail (discussed further in Subheading 4 below) (Fig. 4). As with several of the species discussed above, F. acidarmanus also encodes an unstudied RPA32 protein, with a single OB fold and C-terminal wH domain, encoded by the gene downstream of that encoding FacRPA1, suggesting the presence of a heterodimeric RPA complex (SM, unpublished). Biochemical analysis has shown that both FacRPA1 (in isolation, i.e., in the absence of its cognate RPA32) and FacRPA2 are able to bind ssDNA with high affinity and specificity. FacRPA1, but not FacRPA2, is also able to stimulate the in vitro activity of the M. acetivorans PolB polymerase to overcome pausing on DNA templates [29] and both proteins, but particularly FacRPA2, have been shown to be able to stimulate efficient unwinding of forked DNA substrates by the F. acidarmanus XPD helicase [45].

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Crenarchaeal SSB Proteins The Crenarchaeota are a constituent phylum of the TACK superphylum [46] (Fig. 3a). Perhaps the best characterized crenarchaeal organisms belong to the Sulfolobales, an order that includes the genus Sulfolobus. Various Sulfolobus species have been used as models for diverse aspects of archaeal biology, but S. solfataricus in particular has found a role as a key model organism [34].

4.1 Sulfolobus solfataricus

The S. solfataricus SSB protein (SsoSSB) was the first crenarchaeal single-stranded DNA-binding protein to be biochemically characterized [47, 48] and also the first to have its three-dimensional structure determined [49, 50]. SsoSSB was initially identified in S. solfataricus cell extracts on the basis of its ability to bind to ssDNA and is an abundant, 148-amino acid protein comprising an N-terminal OB-fold domain (residues 1–114) and a flexible C-terminal tail rich in glycine, arginine, and glutamate residues [48]. Overall, this simple structure resembles that of the bacterial SSB proteins. Indeed, despite having very limited primary sequence similarity to E. coli SSB, the SsoSSB protein is able to support growth of a temperature-sensitive E. coli ssb-1 strain at its restrictive temperature, and is capable of stimulating DNA strand exchange by E. coli RecA [47]. The oligomeric state of SsoSSB has been the subject of much discussion, with both monomeric [50, 51] and tetrameric [47, 52] forms being proposed and no straightforward way to reconcile the differences in the published observations. It is agreed however that SsoSSB binds ~5 nucleotides of ssDNA per monomer and with high selectively for ssDNA over dsDNA. Interestingly, recent results indicate that in addition to binding ssDNA, SsoSSB is also able to bind with high affinity to RNA [53], although the biological significance of this remains to be elucidated. The structure of the N-terminal OB-fold domain of SsoSSB has been solved by X-ray crystallography [49, 50] and in complex with ssDNA by NMR (Fig. 6, Table 1) [51, 54]. Structural comparisons with E. coli SSB and eukaryotic RPA point to the SsoSSB OB fold being more similar to the latter, specifically to the DBD-B OB fold (Fig. 2) [50]. Binding relies upon base stacking involving three conserved aromatic residues Trp56, Trp75, and Phe79, and is unidirectional, with each SsoSSB protein-binding ssDNA with the same polarity with respect to the 50 and 30 ends of the DNA [51, 54]. A detailed biophysical analysis of SsoSSB-ssDNA binding has been reported, again highlighting similarities in the dynamics of binding by SsoSSB and DBD-B, rather than E. coli SSB [55]. In common with other SSBs, SsoSSB is able to destabilize double-stranded DNA, a property that is enhanced by the presence in the DNA of single mismatches or more complex lesions such as

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Fig. 6 Solution structure of the SsoSSB bound to single-stranded DNA. The structure corresponds to amino acids 1–117 of S. solfataricus SSB (PDB: 2MNA) bound to single-stranded DNA (shown in red) [51, 54]

cyclobutene pyrimidine dimers (CPDs), the result of UV damage to double-stranded DNA [56]. These observations suggest a potential role for SsoSSB in recognizing DNA lesions in vivo, melting the damaged DNA, and acting as a platform for recruitment of necessary repair factors. Several proteins have been identified as interacting with SsoSSB-coated ssDNA including the XPB helicase Rad50, topoisomerase 1, RNA polymerase (RNAP), and reverse gyrase, although some of these proteins may be interacting with the ssDNA rather than with SsoSSB [56]. Both RNA polymerase and reverse gyrase had previously been shown to interact directly with SsoSSB. Interaction with RNAP stimulates transcription [57], while interaction with reverse gyrase stimulates all the steps in in vitro reverse gyrase activity assays: DNA binding, cleavage, strand passage, and ligation [58]. Given the undoubted importance of ssDNA-binding proteins in all three domains of life, it would be reasonable to expect that the crenarchaeal SSB would be essential for cell viability. The essentiality of SSB has been examined in two closely related Sulfolobus species, S. islandicus and S. acidocaldarius, but with sharply contradictory results. S. islandicus ssb gene deletions are absent from genome-wide transposon insertion libraries and the gene cannot be deleted in three different S. islandicus strains [59]. These results strongly suggest that SisSSB is essential for cell viability, as would be expected. In sharp contrast however, recent evidence suggests that the S. acidocaldarius SSB protein (SacSSB) is not required for cell viability as the ssb gene can apparently be deleted from the chromosome and viable Δssb progeny obtained [60]. Cells deleted for ssb grow indistinguishably from wild type at temperatures ranging from 60 to 80  C, but display cold sensitivity at 50–55  C, heightened sensitivity to novobiocin at 55  C, and sensitivity to heat

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shock at 90  C. If confirmed, the S. acidocaldarius observations raise very significant questions about the cellular functions of the SacSSB protein and how cells are able to cope in its absence. 4.2 Displacement of SSB in the Thermoproteales

The Thermoproteales are a clade of the Crenarchaeota that includes well-studied archaeal organisms such as Thermoproteus tenax and Pyrobaculum aerophilum. Remarkably, unlike all other members of the Crenarchaeota, the majority of sequenced Thermoproteales genomes do not encode a recognizable crenarchaeal-like SSB or euryarchaeal-like RPA. Instead, these organisms encode a distinct ssDNA-binding protein known as ThermoDBP, exemplified by the T. tenax protein referred to here as TteThermoDBP [61]. The gene encoding the 196-amino acid ThermoDBP protein appears to have replaced the ancestral Thermoproteales SSB gene in an example of non-orthologous gene displacement. First identified in a screen for ssDNA-binding proteins in T. tenax cell extracts, recombinant TteThermoDBP has been shown to bind strongly to ssDNA, weakly to ssRNA, and not at all to duplex DNA, consistent with it acting as an SSB in vivo [61]. The structure of the N-terminal ssDNA-binding domain (amino acids 10–148) of TteThermoDBP has been solved by X-ray crystallography, revealing a compact globular domain comprising four α-helices and a four-stranded antiparallel β-sheet (Fig. 7a, Table 1). A putative DNA-binding cleft has also been identified, consisting of a solvent-exposed cleft lined along its length by hydrophobic residues and with positively charged residues at its outer edge [61], and ideally structured for nucleobase and backbone interactions, respectively. Absent from the crystal structure is the C-terminal region of the protein which is helical in nature, possibly forms a basic leucine zipper structure, and is responsible for the dimerization of the recombinant protein [61]. In addition to ThermoDBP, two groups of ThermoDBPrelated proteins (ThermoDBP-RPs) have been identified on the basis of sequence similarity to the N-terminal ssDNA-binding domain of TteThermoDBP [61]. Representatives of the first group, termed ThermoDBP-RP1s, are found in diverse crenarchaeal species including S. solfataricus and euryarchaeal species such as P. furiosus and T. kodakarensis. Representatives of the second group, termed ThermoDBP-RP2s, are found in crenarchaea such as Aeropyrum pernix and Euryarchaea such as Archaeoglobus profundus. Biochemical analysis of purified recombinant ThermoDBPRP1 and ThermoDBP-RP2 proteins has shown that A. pernix ThermoDBP-RP2 binds with high affinity to short (21 nt) mixed sequence or homo-pyrimidine ssDNAs, while not binding, or binding very weakly, to homo-purine ssDNA, dsDNA, or single- or double-stranded RNAs of a similar length [62]. P. furiosus

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Fig. 7 Structure of ThermoDBP and ThermoDBP-related proteins. (a) Structures of the N-terminal domain (amino acids 10–148) of T. tenax ThermoDBP (PDB: 3TEK) [61] and conserved N-terminal domains of P. furiosus ThermoDBP-RP1 (PDB: 4PSL) and A. pernix ThermoDBP-RP2 (PDB: 4PSO) [62]. (b, c) Structures of P. furiosus ThermoDBP-RP1 and A. pernix ThermoDBP-RP2 tetramers. In each case, individual monomers are shown in different colors, with the conserved N-terminal domain of one monomer colored as in part A. Single-stranded DNA bound to ApeThermoDBP-DP2 is colored in dark blue

ThermoDBP-RP1 failed to bind to any of these short substrates, but bound weakly to a longer (45 nt) ssRNA. While earlier results had suggested that S. solfataricus ThermoDBP-RP1 protein was associated with the box C/D small RNAs and the 30S ribosome subunit, in vitro interaction of P. furiosus ThermoDBP-1 with box C/D small RNAs and ribosomes was weak at best, leaving the cellular role of this protein unclear [62]. The three-dimensional structures of the PfuThermoDBP-RP1 and ApeThermoDBP-RP2 protein have been determined by X-ray crystallography, in both cases revealing an N-terminal domain structure similar to that of T. tenax ThermoDBP [61] (Fig. 7a). In the case of PfuThermoDBP-RP1, the N-terminal domain is followed by amphipathic α-helix. PfuThermoDBP-RP1 is a tetramer in solution: in the crystal structure, four of these helices are seen to come together to form an antiparallel four-helix bundle with a pair of N-terminal domains located at either end of this central rodlike structure (Fig. 7b) [62]. The structure of the A. pernix ThermoDBP-RP2 is also tetrameric, comprising intertwined dimers (Fig. 7c). Each monomer contains a ThermoDBP-like N-terminal domain, but also a globular C-terminal domain comprising a five-stranded antiparallel

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β-sheet flanked by two α-helices, connected to the N-terminal domain by a bent linker helix. As with PfuThermoDBP-RP1, ApeThermoDBP-RP2 is a tetramer in solution. The structure of the ApeThermoDBP-RP2 protein bound to ssDNA has also been determined. Binding occurs within a central tunnellike structure that spans multiple subunits and results in significant distortion of the DNA backbone without any change to protein conformation (Fig. 7c). The internal dimensions of the tunnel and the required distortion of the ssDNA backbone are not compatible with dsDNA binding, nor with binding to homo-purine ssDNA (see above). Superimposition of the PfuThermoDBP-RP1 and ApeThermoDBP-RP2 structures reveals that the potential DNA-binding surface is conserved but that in PfuThermoDBPRP1 is occluded by the C-terminal helical tail of another monomer, providing an explanation for the lack of ssDNA binding observed in vitro and at the same time suggesting a simple mechanism for activation of PfuThermoDBP-RP1 ssDNA binding by reorientation of the C-terminal helical tail [62]. Despite this detailed structural knowledge, the cellular functions of the ThermoDBP-RPs, and how these relate to the function of the archetypal ThermoDBP (presumed to be a stand-in for the absent SSB), remain unclear. It is not known if the ThermoDBPRPs are essential for cell viability, for example, whether they play nonessential roles in cellular DNA transactions, or whether they act in concert with other cellular factors. Further work is clearly required to answer these questions to get a full understanding of ThermoDBP and ThermoDBP-RP functions.

5

Summary Since the discovery of the first archaeal RPA-like proteins in 1998 [19], considerable progress has been made in identifying and characterizing ssDNA-binding proteins with diverse structural characteristics in a broad range of archaeal organisms. Biochemical analysis, primarily of proteins expressed and purified in recombinant form, has identified core ssDNA-binding properties, while molecular genetic analysis, most notably in the halophilic archaea, has led to significant new insights into biological function. In the future, technological advances, such as the application of superresolution microscopy to archaeal systems [36], will allow a more detailed analysis of RPA/SSB behavior in vivo, ultimately leading to a comprehensive understanding of the cellular roles of these proteins across the third domain of life.

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Dassarma S (2014) Bioengineering radioresistance by overproduction of RPA, a mammalian-type single-stranded DNA-binding protein, in a halophilic archaeon. Appl Microbiol Biotechnol 98:1737–1747 38. McCready S, Muller JA, Boubriak I, Berquist BR, Ng WL, Dassarma S (2005) UV irradiation induces homologous recombination genes in the model archaeon, Halobacterium sp. NRC-1. Saline Syst 1:3 39. Webb KM, Yu J, Robinson CK, Noboru T, Lee YC, DiRuggiero J (2013) Effects of intracellular Mn on the radiation resistance of the halophilic archaeon Halobacterium salinarum. Extremophiles 17:485–497 40. Whitehead K, Kish A, Pan M, Kaur A, Reiss DJ, King N, Hohmann L, DiRuggiero J, Baliga NS (2006) An integrated systems approach for understanding cellular responses to gamma radiation. Mol Syst Biol 2:47 41. DeVeaux LC, Muller JA, Smith J, Petrisko J, Wells DP, DasSarma S (2007) Extremely radiation-resistant mutants of a halophilic archaeon with increased single-stranded DNA-binding protein (RPA) gene expression. Radiat Res 168:507–514 42. Evans JJ, Gygli PE, McCaskill J, DeVeaux LC (2018) Divergent roles of RPA homologs of the model archaeon Halobacterium salinarum in survival of DNA damage. Genes (Basel) 9:223 43. Pluchon PF, Fouqueau T, Creze C, Laurent S, Briffotaux J, Hogrel G, Palud A, Henneke G, Godfroy A, Hausner W et al (2013) An extended network of genomic maintenance in the archaeon Pyrococcus abyssi highlights unexpected associations between eucaryotic homologs. PLoS One 8:e79707 44. Li Z, Santangelo TJ, Cubonova L, Reeve JN, Kelman Z (2010) Affinity purification of an archaeal DNA replication protein network. MBio 1:e00221–e00210 45. Pugh RA, Lin Y, Eller C, Leesley H, Cann IK, Spies M (2008) Ferroplasma acidarmanus RPA2 facilitates efficient unwinding of forked DNA substrates by monomers of FacXPD helicase. J Mol Biol 383:982–998 46. Guy L, Ettema TJ (2011) The archaeal ’TACK’ superphylum and the origin of eukaryotes. Trends Microbiol 19:580–587 47. Haseltine CA, Kowalczykowski SC (2002) A distinctive single-strand DNA-binding protein from the archaeon Sulfolobus solfataricus. Mol Microbiol 43:1505–1515 48. Wadsworth RI, White MF (2001) Identification and properties of the crenarchaeal single-

Archaeal RPA/SSB Proteins stranded DNA binding protein from Sulfolobus solfataricus. Nucleic Acids Res 29:914–920 49. Kerr ID, Wadsworth RI, Blankenfeldt W, Staines AG, White MF, Naismith JH (2001) Overexpression, purification, crystallization and data collection of a single-stranded DNA-binding protein from Sulfolobus solfataricus. Acta Crystallogr D Biol Crystallogr 57:1290–1292 50. Kerr ID, Wadsworth RI, Cubeddu L, Blankenfeldt W, Naismith JH, White MF (2003) Insights into ssDNA recognition by the OB fold from a structural and thermodynamic study of Sulfolobus SSB protein. EMBO J 22:2561–2570 51. Gamsjaeger R, Kariawasam R, Gimenez AX, Touma C, McIlwain E, Bernardo RE, Shepherd NE, Ataide SF, Dong Q, Richard DJ et al (2015) The structural basis of DNA binding by the single-stranded DNA-binding protein from Sulfolobus solfataricus. Biochem J 465:337–346 52. Rolfsmeier ML, Haseltine CA (2010) The single-stranded DNA binding protein of Sulfolobus solfataricus acts in the presynaptic step of homologous recombination. J Mol Biol 397:31–45 53. Morten MJ, Gamsjaeger R, Cubeddu L, Kariawasam R, Peregrina J, Penedo JC, White MF (2017) High-affinity RNA binding by a hyperthermophilic single-stranded DNA-binding protein. Extremophiles 21:369–379 54. Gamsjaeger R, Kariawasam R, Touma C, Kwan AH, White MF, Cubeddu L (2014) Backbone and side-chain 1H, 13C and 15N resonance assignments of the OB domain of the single stranded DNA binding protein from Sulfolobus solfataricus and chemical shift mapping of the DNA-binding interface. Biomol NMR Assign 8:243–246 55. Morten MJ, Peregrina JR, Figueira-GonzalezM, Ackermann K, Bode BE, White MF, Penedo JC (2015) Binding dynamics of a monomeric SSB protein to DNA: a singlemolecule multi-process approach. Nucleic Acids Res 43:10907–10924

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56. Cubeddu L, White MF (2005) DNA damage detection by an archaeal single-stranded DNA-binding protein. J Mol Biol 353:507–516 57. Richard DJ, Bell SD, White MF (2004) Physical and functional interaction of the archaeal single-stranded DNA-binding protein SSB with RNA polymerase. Nucleic Acids Res 32:1065–1074 58. Napoli A, Valenti A, Salerno V, Nadal M, Garnier F, Rossi M, Ciaramella M (2005) Functional interaction of reverse gyrase with single-strand binding protein of the archaeon Sulfolobus. Nucleic Acids Res 33:564–576 59. Zhang C, Phillips APR, Wipfler RL, Olsen GJ, Whitaker RJ (2018) The essential genome of the crenarchaeal model Sulfolobus islandicus. Nat Commun 9:4908 60. Suzuki S, Kurosawa N (2019) Robust growth of archaeal cells lacking a canonical singlestranded DNA-binding protein. FEMS Microbiol Lett 366:fnz124 61. Paytubi S, McMahon SA, Graham S, Liu H, Botting CH, Makarova KS, Koonin EV, Naismith JH, White MF (2012) Displacement of the canonical single-stranded DNA-binding protein in the Thermoproteales. Proc Natl Acad Sci U S A 109:E398–E405 62. Ghalei H, von Moeller H, Eppers D, Sohmen D, Wilson DN, Loll B, Wahl MC (2014) Entrapment of DNA in an intersubunit tunnel system of a single-stranded DNA-binding protein. Nucleic Acids Res 42:6698–6708 63. Raghunathan S, Kozlov AG, Lohman TM, Waksman G (2000) Structure of the DNA binding domain of E. coli SSB bound to ssDNA. Nat Struct Biol 7:648–652 64. Zimmermann L, Stephens A, Nam SZ, Rau D, Kubler J, Lozajic M, Gabler F, Soding J, Lupas AN, Alva V (2017) A completely reimplemented MPI Bioinformatics Toolkit with a new HHpred server at its core. J Mol Biol 430:2237–2243

Chapter 3 Single-Molecule Fluorescence Methods to Study Protein Exchange Kinetics in Supramolecular Complexes Richard R. Spinks, Lisanne M. Spenkelink, and Antoine M. van Oijen Abstract Recent single-molecule studies have demonstrated that the composition of multi-protein complexes can strike a balance between stability and dynamics. Proteins can dynamically exchange in and out of the complex depending on their concentration in solution. These exchange dynamics are a key determinant of the molecular pathways available to multi-protein complexes. It is therefore important that we develop robust and reproducible assays to study protein exchange. Using DNA replication as an example, we describe three single-molecule fluorescence assays used to study protein exchange dynamics. In the chase exchange assay, fluorescently labeled proteins are challenged by unlabeled proteins, where exchange results in the disappearance of the fluorescence signal. In the FRAP exchange assay, fluorescently labeled proteins are photobleached before exchange is measured by an increase in fluorescence as non-bleached proteins exchange into the complex. Finally, in the two-color exchange assay, proteins are labeled with two different fluorophores and exchange is visualized by detecting changes in color. All three assays compliment in their ability to elucidate the dynamic behavior of proteins in large biological systems. Key words Exchange, Single-molecule, Replication, Fluorescence, DNA, Supramolecular complexes, Compositional dynamics

1

Introduction Traditional in vitro biochemical assays are invaluable to study protein function and determine kinetic parameters that govern protein binding and enzymatic activity. Methods such as spectrophotometry, surface plasmon resonance, and isothermal titration calorimetry are well-established standards used to measure molecular kinetics. These methods, however, provide measurements of molecular properties that represent the average behavior of many asynchronous components. Therefore, short-lived intermediates and stochastic dynamics are difficult, if not impossible, to observe using these traditional biochemical assays. Single-molecule detection techniques offer the potential to circumvent ensemble averaging by visualizing individual proteins.

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Single-molecule experiments can reveal rare substates, alternative reaction pathways, or interaction hot spots that are otherwise masked in ensemble experiments [1]. Using single-molecule approaches, stochastic processes can be detected as they occur, opening the door to previously inaccessible kinetic information. In particular, the dynamic exchange of proteins is one type of molecular behavior that has not been accessible through ensemble-averaging methods. A variety of multi-protein complexes have been shown to exhibit protein exchange behavior that is dependent on protein concentration [2–5]. Specifically, a series of single-molecule studies have demonstrated that the highly stable DNA replication machinery is capable of exchanging proteins for competitors from the environment in a concentration-dependent manner [6–12]. Concentration-dependent protein exchange, at first glance, seems contradictory to the principle of dissociation rate constants being independent of concentration. Aberg et al. [13] and Sing et al. [14], however, have modeled large multi-protein complexes and how the kinetics of a protein can satisfy both stability and exchange mechanisms. For multi-protein complexes supported by many weak binding interactions, the intricate choreography of bonds being rapidly made and unmade provides the ability to balance stability with plasticity. The observation of protein exchange suggests that supramolecular protein complexes have a degree of plasticity allowing the complex to overcome challenges in a constantly changing molecular world [15–17]. Exchange dynamics play a key role in determining the molecular pathways that are available to multi-protein complexes. It is therefore important that we develop robust and reproducible assays to study this phenomenon. In this chapter, we describe the experimental protocols underpinning three complementary singlemolecule fluorescence techniques that all allow the visualization and quantification of protein exchange dynamics. Specifically, we focus on the use of a single-molecule rolling-circle replication assay [18–20] to study exchange in the multi-protein DNA-replication complex. In this assay, DNA templates are tethered to the surface of microfluidic flow cells and stained with fluorescent intercalators to allow the real-time visualization of flow-stretched DNA-replication products [21]. Simultaneously, we use fluorescently labeled replication proteins to detect the dynamics of protein exchange. The three assays herein, termed the “chase exchange assay,” the “FRAP exchange assay,” and the “two-color exchange assay,” have all been employed to examine exchange kinetics of proteins in the DNA replication machineries of T7 phage [9], E. coli [7, 8], and S. cerevisiae yeast [6]. These three types of single-molecule exchange assays, although similar in their methodology, each provides a unique insight into the mechanisms of protein exchange. Each assay reveals something

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different; the chase exchange assay detects the unbinding of initially bound proteins by the disappearance of fluorescence. The FRAP exchange assay measures appearance of fluorescence, thereby revealing association of proteins with the replisome. The two-color exchange assay uses two fluorescent signals to visualize proteins as they transiently associate and disassociate. When used together they provide a comprehensive characterization of exchange kinetics. While our examples described in this chapter relate to DNA replication as our biological system of choice, the exchange assays could be easily adapted to almost any biological system. Most experimental aspects described below are directly applicable to the observation of any process supported by multi-protein complexes on DNA, while the general approach is broadly applicable to multiprotein complexes that can be surface immobilized and observed by single-molecule fluorescence imaging approaches.

2

Materials

2.1 Glass-Surface Functionalization

1. 24
24 mm Glass microscope coverslips. 2. Plastic coverslip washing container. 3. 100% Anhydrous ethanol. 4. 1 M Potassium hydroxide (KOH). 5. Acetone. 6. (3-Aminopropyl)triethoxysilane (APTES). 7. Biotin-PEG and MPEG (MW 5000) bundle (store under N gas at 20  C). 8. PEGylation buffer: 100 mM NaHCO3, pH 8.2 (prepare fresh). 9. Oven (110  C). 10. Bath sonicator. 11. Compressed nitrogen gas.

2.2 Microfluidic Flow Cell Device

1. PDMS mold: Metal mold that is laser-engraved with a ridge measuring 0.1
0.5
19 mm. 2. Microscope-compatible flow cell holder. 3. Polydimethylsiloxane. 4. 184 Silicone Elastomer. 5. PE60 Tubing (0.76 inner diameter, 1.22 mm outer diameter). 6. NeutrAvidin protein solution. 7. PDMS lid-cleaning solutions: 0.5% Triton, 1 M NaHCO3, 70% ethanol.

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8. Milli-Q water. 9. Compressed nitrogen gas. 10. Vacuum desiccator. 2.3 Exchange Assay Imaging

1. SYTOX Orange nucleic acid stain. 2. 100 μM dNTP bundle, mix all 4 equally to make a 25 μM dNTP solution. 3. 100 μM NTP bundle, mix all 4 equally to make a 25 μM NTP solution. 4. Biotinylated rolling-circle DNA template. 5. Heating block. 6. Single-molecule (SM) reaction buffer: 30 mM Tris–HCl pH 7.5, 12 mM magnesium acetate, 50 mM potassium glutamate, 0.5 mM EDTA, 0.0025% Tween20, 0.5 mg/mL BSA (store at 4  C). 7. Blocking solution: 1
SM reaction buffer, 2% Tween20 (store away from light for up to 2–3 weeks). 8. DNA solution: 1
SM reaction buffer, 10 mM dithiothreitol, 1 mM ATP, 150 mM SYTOX Orange, 20 pM biotinylated rolling-circle DNA template. 9. Replication solution: 1x SM reaction buffer, 10 mM dithiothreitol, 1 mM ATP, 125 μM dNTPs, 250 μM NTPs, replication proteins, fluorescent protective reagents. 10. Typical E. coli replication proteins: 30 nM DnaB6(DnaC)6, 30 nM αεθ pol III core, 10 nM τ3δδ’χψ clamp loader complex, 30 nM β2 clamp, 75 nM DnaG, 200 nM SSB. 11. Fluorescently labeled protein: Any protein can be substituted for its labeled counterpart. In our example experiments, we use pol III core labeled with Alexa Fluor 647 or Alexa Fluor 488 via SNAP-tag [8], and SSB labeled with Alexa Fluor 647 via cysteine-maleimide coupling [7]. 12. Fluorescence protective reagents: 1 mM UV-aged Trolox, 10% w/v glucose, 0.45 mg/mL glucose oxidase, and 21 μg/mL catalase (these reagents increase the lifetime of the fluorophores and reduce blinking). 13. Single-molecule inverted TIRF microscope: Nikon optical microscope body, 100
TIRF objective (N.A. ¼ 1.49, oil), EMCCD camera, heated-stage insert, lasers matching the excitation wavelength of the fluorophores used (we used 647 nm for Alexa Fluor 647, 488 nm for Alexa Fluor 488, and 568 nm for SYTOX Orange), excitation and emission filters appropriate for each laser line, DV2 dual-view apparatus, SyringeONE syringe pump with 5 mL syringe.

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14. Acquisition software: Nikon Elements Advanced Research. 15. Syringe pump software: SyringePumpPro. 2.4

Data Analysis

1. Movie analysis software: ImageJ/FIJI (version 1.52e), with the TrackMate plug-in (version 3.6.0) [22], as well as custom plugins designed to expedite the analysis process (the plug-ins are freely available on GitHub, https://github.com/ SingleMolecule and https://github.com/LMSpenkelink/ SingleMoleculeReplication). 2. Data plotting and fitting software: MATLAB 2016b.

3

Methods

3.1 Chase Exchange Assay

In this section, we describe the experimental details of the chase exchange assay and explain how it is used to visualize protein exchange. Firstly, replication is initiated with the fluorescently labeled protein of interest. The reaction solution is then rapidly switched for a solution containing an unlabeled version of the same protein. Protein exchange can be identified by a disappearance of the fluorescence signal as unlabeled proteins exchange for labeled proteins in the replisome. Because of its relative simplicity, this assay is ideal to provide an initial characterization of exchange mechanisms. However, discriminating between the signals of photobleaching and exchange can be challenging. The following chase exchange assay protocol describes how to prepare the functionalized coverslip, assemble the microfluidic device, carry out the fluorescence microscopy, and finally analyze the resulting data.

3.1.1 Glass-Surface Functionalization

A functionalized glass surface is employed to reduce non-specific surface interactions and allow specific tethering of the DNA replication template within the flow cell channel. The procedure described here is an adapted version of the functionalization method described by Geertsema et al. [20]. In order to achieve functionalization, the coverslips are cleaned, and the silicon surface covalently bonded to aminosilane, which is in turn reacted with succinimidyl polyethylene glycol (PEG). The PEG is a mixture of unreactive, methylated PEG and biotinylated PEG that can bind the DNA template via a biotin–streptavidin–biotin link (Fig. 1, inset). 1. Using the plastic washing container, sonicate glass coverslips in anhydrous ethanol for 30 min to liberate any hydrophobic contaminants. Dispose of the ethanol appropriately and rinse the coverslips with water. Then sonicate coverslips in 1 M KOH

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Fig. 1 Experimental setup used for the three different single-molecule exchange assays. Reaction solutions are loaded into the channel of a microfluidic device by hydrodynamic flow applied via a syringe pump. The microfluidic device is positioned on the microscope stage and laser light of a specific excitation wavelength is coupled through the microscope objective to illuminate the channel via total internal reflected fluorescence (TIRF). The excited proteins fluoresce at a specific emission wavelength. This signal is captured by an EMCCD camera and analyzed using custom-built software. (Inset) On the molecular scale, DNA templates are immobilized on the surface of the coverslip via a biotin–streptavidin–biotin link and the exchange reaction components are sequentially assembled in situ by loading different solutions

for 30 min to remove any hydrophilic contaminants. Dispose of the KOH appropriately and rinse the coverslips again. Repeat both the ethanol washing step and KOH washing step. 2. Prior to silanization, all water needs to be removed by washing three times with acetone. Sonicate in acetone on the last wash. Prepare the solution of 3-aminopropyl triethoxysilane (see Note 1) and introduce it to the coverslips. Gently agitate the coverslips by hand or on an orbital shaker for 10 min, sonicate

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for 1 min, and then return them to the shaker for another 10 min. The repeat sonication maximizes surface group density and uniformity. Quench the silanization reaction by flushing the container with 10–15 volumes of water (see Note 2). Dry the silanized coverslips using dry-compressed N2 gas. 3. Remove the m-PEG and biotin-PEG from the freezer and allow them to warm to room temperature on the benchtop before use (see Note 3). Mix the m-PEG and biotin-PEG in a ratio of 25:1 in the PEGylation buffer and to a final concentration of 0.2% biotin-PEG (for example, 75 mg m-PEG and 3 mg biotin-PEG in 500 mL buffer). Swiftly after dissolving the PEG, apply ~50 μL to the surface of a coverslip laid flat inside a humidity chamber (see Note 4). Place a second silanized coverslip on top of the first coverslip to make a “coverslip sandwich” (this conserves space and PEG solution). Make more coverslip sandwiches with the remaining silanized coverslips. Incubate the coverslips in the humidity chamber for 3 h at room temperature and away from light. Rinse the coverslips with water (making note of which is the functionalized side; see Note 5) and dry with N2 gas. Following that, repeat the PEGylation process with a fresh PEG solution so the coverslips become doubly PEGylated. This second batch of coverslip sandwiches can be left overnight. Rinse and dry the coverslips again and store them under vacuum for up to 2 weeks. Any longer storage sees degradation in surface passivation. 3.1.2 Constructing a Microfluidic Flow Cell Device

A microfluidic flow cell device that is microscope compatible is essential to assemble the reaction components in situ and to provide a continuous solution flow. The flow cell device consists of a combination of a functionalized coverslip, an imprinted PDMS lid, and inlet and outlet tubing (Fig. 1). 1. Polydimethylsiloxane (PDMS) is a silicon-based polymer that is generally inert, nontoxic, and relatively easy to use to create custom structures. To make our flow cell lid, pour PDMS mixed with a silicone-curing agent (1:10 ratio) into a metal mold that is laser-engraved with a ridge measuring 0.1
0.5
19 mm (see Note 6). This ridge imprints a channel on the bottom surface of the PDMS block (see Note 7). Place the PDMS in a vacuum desiccator to remove air bubbles for 1 h, and then bake in the oven at 70  C for 2 h. Once set, cut the block from the mold and then pierce 0.6 mm diameter holes through the PDMS to connect to each end of the channel. From here, the PDMS block is reusable following a wash procedure (see Note 8). 2. The following step should be done immediately prior to conducting the actual exchange assay. Spread 125 μL of 0.2 mg/ mL NeutrAvidin solution over a previously functionalized glass

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coverslip and incubate in a humidity chamber at room temperature for 30 min. Rinse and dry the coverslip before pressing the PDMS block onto the functionalized side ensuring that the seal is airtight. To prevent the conjoined coverslip and PDMS block from separating during the experiment they are secured within a flow cell holder that is also compatible to fit with the microscope stage. To complete the assembly of the flow cell, insert 11 cm inlet and outlet tubing halfway into each hole using tweezers. Use a 21-gauge needle and a syringe to slowly draw degassed blocking solution into the flow channel and leave for 20 min to further minimize nonspecific interactions with the surface. Afterwards, the flow cell is ready for use. 3.1.3 Imaging the Chase Exchange Assay

The imaging component of the chase exchange assay involves the use of a single-molecule total internal reflection fluorescence (TIRF) microscope to image individual proteins. The imaging can be divided into two phases: first, replication is initiated on an anchored rolling-circle DNA template with a replication solution that includes a fluorescently labeled protein of interest. Second, at a defined change point, the active solution is switched to an identical solution that contains unlabeled proteins. A decrease of the initial fluorescence signal after the solution switch indicates that the unlabeled proteins have exchanged into the replisome (exchange rate must be faster than the photobleaching rate to be detected) (see Note 9). 1. Position the assembled flow cell on the microscope stage. Connect the outlet tubing to a syringe pump and leave the inlet tubing in the current reservoir containing the blocking solution (Fig. 1). Turn on the appropriate laser line(s) and TIRF microscope and make sure that the laser incident angle is at TIRF. Focus the objective to the glass and position the stage to ensure that you are viewing the flow cell channel. 2. DNA solution and replication solution should be prepared in degassed SM reaction buffer to prevent formation of air bubbles during the experiment. 3. When the flow cell channel is ready, pause the flow from the syringe pump and pinch the inlet tubing (either by hand or bull clip) to transfer it to the DNA solution. Next, rapidly flow (e.g., ~100 μL/min) DNA solution equal to the volume of the inlet tubing (e.g., 11 cm of PE-60 tubing gives ~50 μL) and then flow solution at a slow rate (5–20 μL/min) to allow the DNA template molecules to bind the surface (Fig. 1, inset). Once optimum DNA template density is achieved (determined by visualizing the DNA using fluorescent intercalator) transfer the inlet tube to the first replication solution containing the fluorescently labeled protein.

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4. Start flowing the first replication solution at a steady rate of 10 μL/min and start the acquisition (typical acquisition parameters are 2–6 min length, an exposure time of 200 ms, and a laser intensity of 80–800 mW/cm2). Under laser excitation, foci of active proteins should be observable as the anchored DNA templates start to replicate (see Note 10). 5. As the experiment is approaching the defined change point, switch the solution to the second replication solution (e.g., change point ¼ 3 min, switch at 2 min), accounting for the time it takes for the new solution to flow through the inlet tube (flow at 50 μL/min for 50 μL taking 1 min). Then continue flowing at the normal rate for the duration of the experiment (Fig. 2a, top). 6. Once the experiment is complete, disassemble the flow cell. The coverslip and tubing are discarded and the PDMS block is reused after a strict washing procedure (see Note 8). 3.1.4 Data Quantification

In order to ascertain whether protein exchange has occurred in the chase exchange experiment, we need to quantify the lifetime of the fluorescently labeled protein at the replication fork and compare this to the average photobleaching lifetime of the fluorophore. There are two major steps in the foci quantification procedure: first, the raw movie is corrected for background intensity, and second, individual foci are tracked over the course of the movie and their intensity measured in each frame. We use ImageJ with custom plug-ins to expedite the analysis process with reproducible speed and accuracy. These plug-ins are specifically made for our type of data; however, the analysis principles of the plug-ins are applicable to all kinds of exchange data potentially captured by the chase exchange assay. 1. Open the movie in ImageJ and convert the stack to 32-bit to preserve data quality. To background correct the image stack, first create a beam profile image by averaging the whole stack and then applying a smoothing Gaussian filter with a large sigma value (e.g., sigma ¼ 80, or until the image shows a uniform profile). Correct each frame of the stack by first subtracting the camera electronic offset value for each pixel, and then dividing the whole frame by the normalized beam profile image. 2. Next, draw a region of interest (ROI) box around the target molecule and use the TrackMate plug-in [22] to track this focus spatially through the movie. Use the TrackMate output to create a ROI centered about the molecule for each frame and add these ROIs to the ROI manager. For each ROI, integrate the pixel intensity and subtract the local background generated

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Fig. 2 The typical experimental pipeline for each single-molecule exchange assay. (a) Chase exchange assay. (Top) Schematic representation of the assay. (Middle) Simulated kymograph and graph of intensity over time showing the expected outcome. The fluorescence signal disappears when unlabeled proteins exchange for the existing labeled ones. (Bottom) Simulated average intensity over time for a large number of single molecules (typically n ¼ 20–40 molecules). The average signal decreases faster than photobleaching occurrence. (b) FRAP exchange assay. (Top) Schematic representation of the assay. (Middle) Simulated kymograph and graph of intensity over time showing the expected outcome. After each FRAP pulse (indicated by the vertical dashed lines), all the fluorophores have bleached. The fluorescence intensity recovers as unbleached proteins exchange into the system. (Bottom) Simulated averaged normalized intensity over time after a FRAP pulse. This curve can be fit to provide a characteristic exchange time. (c) Two-color exchange assay. (Top) Schematic representation of the assay. (Middle) Simulated kymograph and graph of intensity over time showing the expected outcome. The fluorescence intensity of the two colors alternates as the proteins exchange. (Bottom) Simulated average autocorrelation signal. A single exponential fit to the autocorrelation can reveal the characteristic timescale of exchange

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by integrating the intensity of an ROI twice the size. Save the measured intensity values and then plot the intensity over time for that particular focus in MATLAB (or your preferred graphing software) (Fig. 2a, middle, and Fig. 3a, top). 3. Repeat the signal quantification for all target foci and then create an averaged graph of intensity over time. This is representation of the average lifetime of the fluorescently labeled proteins and any decrease after the defined change point is indicative of protein exchange. 4. To further validate this experiment, the average fluorescent lifetime measurement can be compared to the average photobleaching lifetime calculated in the same way and under identical conditions. 5. In the presence of protein exchange, the active labeled protein lifetime should be significantly different from the photobleaching lifetime after the defined change point (Fig. 2a, bottom, and Fig. 3a, bottom). For a more quantitative determination of the exchange rate, see the FRAP exchange assay in Subheading 3.2. 3.2 FRAP Exchange Assay

The classical ensemble averaging approach to fluorescence recovery after photobleaching (FRAP) involves purposely bleaching a sample with high laser intensity and using the recovery of fluorescence as a reporter of diffusion kinetics [23]. We adapted the FRAP concept to the single-molecule scale to quantify exchange rates for individual multi-protein complexes. By bleaching the fluorescence signal of single-protein molecules, we can monitor the recovery of the fluorescence signal as unbleached proteins from solution exchange for the bleached proteins. In this section, we detail the technical steps involved in the FRAP exchange assay. Please note that the FRAP exchange assay uses the same surface functionalization method as described in Subheading 3.1.1 and the same design of the microfluidic flow cell device as described in Subheading 3.1.2. We use SSB exchange during DNA replication as an example; however, like the other exchange assays stated in this chapter, this assay has the potential for broader application.

3.2.1 Imaging the FRAP Exchange Assay

The FRAP exchange assay is considered relatively accessible, as only the imaging steps are different compared to the chase assay, and no requirements are placed on the availability of additional reagents (e.g., labeled proteins). Similar to the other assays in this chapter, replication is initiated, and imaging begins. Subsequently, at a specified time point, a FRAP pulse of high laser power is used to purposely bleach all fluorescent proteins in the field of view (Fig. 2b, top). Any recovered fluorescence observed thereafter is a direct result of exchange.

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Fig. 3 Data and analysis for each of the three single-molecule exchange assays. (a) Example chase exchange experimental data. (Top) Representative kymograph showing Pol III*-AF647 on an individual DNA molecule. The Pol III* moves with the direction of flow as the replisome elongates a DNA molecule. Initially visible as bright magenta spot, the intensity of the Pol III* signal decreases after the buffer change point. This decrease indicates that exchange with unlabeled Pol III* has occurred. (Bottom) The average intensity of Pol III*-AF647 (n ¼ 280) after the change point is clearly decreasing faster than average photobleaching lifetime (n ¼ 296) at the same laser power (colored area represents standard deviation; note: previously unpublished data). (b) Example FRAP exchange experimental data. (Top) A representative kymograph of SSB-AF647 as part of a replicating DNA molecule in a FRAP exchange experiment. After each high-intensity FRAP pulse all SSB molecules have bleached. The fluorescence intensity recovers as unbleached SSB exchanges into the replisome. (Bottom) The average recovered intensity of SSB-AF647 for each time period was fit with formula 1 to provide a characteristic exchange time. Comparison of exchange times across different concentrations of SSB shows a decrease in exchange time as concentration increases (2 nM SSB: τ ¼ 20  7 s, N ¼ 20; 10 nM SSB: τ ¼ 10  1 s, N ¼ 24; 20 nM SSB: τ ¼ 5.0  1.8 s, N ¼ 21; 100 nM SSB: τ ¼ 2.9  1.7 s, N ¼ 18) (adapted from Spenkelink et al. [7] with permission under the terms of the Creative Commons CC BY license). (c) Example two-color exchange experimental data. (Top) Kymograph of two colors of Pol III* (magenta ¼ AF647 labeled and green ¼ AF488 labeled) shows that polymerases are stochastically exchanged at the fork of a replicating DNA molecule. Quantifying the intensity of each color of Pol III* reveals that they are sometimes exchanged but also sometimes colocalized (adapted from Lewis et al. [8] with permission under the terms of the Creative Commons Attribution License)

1. Position the blocked flow cell on the microscope stage and focus the objective up to the glass to view the channel. Define appropriate imaging parameters including the parameters that describe the FRAP pulse (typical parameters: 2–6 min length, 200 ms exposure, 80–800 mW/cm2 laser intensity for visualization, with FRAP pulses of 300 W/cm2 for 5 s every 30 s; see Note 11).

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2. Prepare the DNA solution and replication solution in degassed buffer. When the channel is ready, introduce the DNA solution to the channel by first rapidly flowing (e.g., ~100 μL/min), and then flowing at a slow rate (e.g., 5–20 μL/min). Once optimum DNA template density is achieved, initiate the replication reaction. 3. In contrast to the chase exchange assay in which a constant laser intensity is used, the laser intensity is periodically increased to rapidly photobleach all fluorescence in the field of view. Exchange can be observed as a recovery of the fluorescence signal (Fig. 2b, middle). Start the imaging procedure. Single SSBs should be observable as the anchored DNA templates are replicated. If SSB exchange is relatively frequent, then more frequent FRAP pulses can be applied in one experiment to increase the amount of available exchange data (Fig. 3a, top). 4. Leave the imaging sequence to complete and then disassemble the flow cell. 3.2.2 Data Quantification

The output of the FRAP exchange assay is easily quantified as it follows the same principles of conventional FRAP experiments. The averaged recovery intensity signal can be fitted with an exponential function to derive the characteristic exchange constant. 1. Use ImageJ to correct the image stack the same as in Subheading 3.1.4, step 1. 2. Identify foci of interest and use the same protocol as in Subheading 3.1.4, step 2, to calculate the intensity over time for each focus (the bleached, dark period of each focus should still be measured). 3. Create an averaged trace of recovered intensity over time by synchronizing multiple signals to the end of the FRAP pulse. Fit the averaged recovery curve with a FRAP recovery function correcting for photobleaching (formula 1, where τ is the characteristic exchange time and tb is the photobleaching lifetime) (Fig. 2b, bottom, and Fig. 3b, bottom). Evaluating the goodness of fit of the function to the data will provide the error of the derived measurement: I ðtÞ ¼ ae

3.3 Two-Color Exchange Assay

tt

b

þ bð1  e τ Þ t

ðformula 1Þ

Similar to the chase exchange assay, the two-color exchange assay also utilizes single-molecule fluorescence to visualize exchange of proteins in biological systems. In this case, exchange is detected when a fluorescently labeled protein is substituted for a protein labeled with a spectrally different fluorophore (Fig. 2c, top). Unlike the chase exchange assay, this method detects exchange as it occurs stochastically and is not reliant on a change in solution conditions.

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This section describes the protocol to complete the two-color exchange assay. The assay uses the same surface functionalization method as described in Subheading 3.1.1 and the same design of the microfluidic flow cell device as described in Subheading 3.1.2. Our technical description is based on polymerase exchange [8], although the two-color exchange assay has an array of potential applications. 3.3.1 Imaging the Two-Color Exchange Assay

In contrast to the chase exchange assay, the two-color exchange assay only has one phase. The reaction solution and imaging parameters, however, are more intricate. To begin the experiment, replication is initiated on an anchored DNA template with a replication solution that includes equal concentration of two colors of labeled polymerases. The corresponding two laser lines must be used to image both proteins simultaneously. During the experiment, as the two colors of polymerase exchange stochastically, there should be an obvious anticorrelation in the intensities (Fig. 2c, middle) (the exchange rate must be slower than the frame rate to be detected). 1. Position the blocked flow cell on the microscope stage and focus the objective up to the glass to view the channel (Fig. 1). Define appropriate imaging parameters to capture data from two emission sources (see Note 12). 2. Prepare the DNA solution and replication solution in degassed SM reaction buffer. When the channel is ready, introduce the DNA solution to the channel by first rapidly flowing (e.g., ~100 μL/min), and then flowing at a slow rate (e.g., 5–20 μL/min). Once optimum DNA template density is achieved, start flowing the replication solution to initiate the reaction (Fig. 1, inset). 3. Start the imaging procedure with the defined parameters (typical parameters: 2–6 min length, exposure time 200 ms, 80–800 mW/cm2 intensity for each laser). Single polymerases in each color should be observable as the anchored DNA templates are replicated (Fig. 3c, top). Allow the imaging sequence to complete and then disassemble the flow cell.

3.3.2 Data Quantification

It might be clear by eye when exchange has occurred in the two-color exchange experiment. To quantify this behavior more accurately we need to compare the signal intensities of two differently colored proteins colocalized at the same replication event. The analysis procedure to obtain signal intensity values is the same as the chase exchange assay (Subheading 3.1.4), with one exception: Each laser line generates its own beam profile, and therefore the correction needs to be applied separately to each color. 1. Use ImageJ to correct the image stack the same as in Subheading 3.1.4, step 1.

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2. Identify colocalized foci of interest and use the same protocol as in Subheading 3.1.4, step 2, to calculate the intensity over time for each focus in each color. 3. Plot the intensity signal of both colors of one molecule on the same graph. If there is interrelated changes in each color’s signal, then this is characteristic of protein exchange (Fig. 2c, middle, and Fig. 3c, bottom). 4. Cross-correlation analysis can accurately identify the existence of exchange. Depending on signal quality, it can also potentially derive the characteristic exchange time as a function of one exchange event relative to another (Fig. 2c, bottom).

4

Notes 1. The 3-aminopropyl triethoxysilane reagent is highly sensitive and should only be used for a few days after opening; store in dehumidified desiccator at room temperature; we find Alfa Aesar or Sigma suppliers provide a high-quality product. 2. The silane solution should be gone once excessive bubbles no longer form in the water. 3. Water from any condensation will hydrolyze part of the PEG and consequently hinder proper functionalization. 4. A humidity chamber is easily created from an Eppendorf tube rack with wells half-filled with water and a lid. 5. The coverslip sandwiches can be tricky to separate, and it is best done with tweezers, and gloved hands. 6. The PDMS and curing agent need to be mixed very well (vigorously stir for >3 min) to ensure uniform curing of the PDMS. 7. If multiple channels are imprinted side by side in the one PDMS block then multiple single-molecule experiments can be carried out in quick succession. 8. Once the single-molecule imaging is complete, the PDMS lid can be cleaned for future reuse. This is done by sonicating for 20 min with 0.5% Triton, followed by 20 min with 1 M NaHCO3, and finally 20 min in 70% ethanol. In between each step, the PDMS lid is washed 2–3 times with Milli-Q water. After the final wash, the PDMS lid is dried with compressed air and stored. 9. If the situation suits, the opposite experiment can be done where replication is initiated with unlabeled protein and switched for labeled protein. In this case, we are looking for the appearance of signal to indicate that exchange has occurred.

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10. Depending on the success of the blocking step, there may be some single molecules of your fluorescent protein non-specifically bound to the surface. Some non-specific binding is acceptable, however, as long as active molecules can still be detected among the noise. 11. The FRAP pulse should bleach the entire field of view in a few seconds. Hence, the time and intensity need to be carefully chosen. Overexposure will result in extra-unwanted reactive oxygen species in the reaction solution. Some trial-and-error experiments may help determine the best parameters. 12. We chose to image two colors by using a dual-view device to direct light from each color onto separate halves of the camera chip. This means that the data from each color are perfectly correlated in time. Alternatively, if you are not so concerned with time correlation, you can rapidly switch between laser lines and filter sets. Although not correlated in time it provides a larger detectable field of view. References 1. Monachino E, Spenkelink LM, van Oijen AM (2017) Watching cellular machinery in action, one molecule at a time. J Cell Biol 216 (1):41–51. https://doi.org/10.1083/jcb. 201610025 2. Chen T-Y, Santiago AG, Jung W, Krzemin´ski Ł, Yang F, Martell DJ, Helmann JD, Chen P (2015) Concentration- and chromosomeorganization-dependent regulator unbinding from DNA for transcription regulation in living cells. Nat Commun 6:7445–7445. https:// doi.org/10.1038/ncomms8445 3. Gibb B, Ye LF, Gergoudis SC, Kwon Y, Niu H, Sung P, Greene EC (2014) Concentrationdependent exchange of replication protein A on single-stranded DNA revealed by singlemolecule imaging. PLoS One 9(2): e87922–e87922. https://doi.org/10.1371/ journal.pone.0087922 4. Graham JS, Johnson RC, Marko JF (2011) Concentration-dependent exchange accelerates turnover of proteins bound to doublestranded DNA. Nucleic Acids Res 39 (6):2249–2259. https://doi.org/10.1093/ nar/gkq1140 5. Delalez NJ, Wadhams GH, Rosser G, Xue Q, Brown MT, Dobbie IM, Berry RM, Leake MC, Armitage JP (2010) Signal-dependent turnover of the bacterial flagellar switch protein FliM. Proc Natl Acad Sci U S A 107 (25):11347–11351. https://doi.org/10. 1073/pnas.1000284107 6. Lewis JS, Spenkelink LM, Schauer GD, Yurieva O, Mueller SH, Natarajan V, Kaur G,

Maher C, Kay C, O’Donnell ME, van Oijen AM (2020) Tunability of DNA polymerase stability during eukaryotic DNA replication. Mol Cell 77(1):17–25.e15. https://doi.org/10. 1016/j.molcel.2019.10.005 7. Spenkelink LM, Lewis JS, Jergic S, Xu ZQ, Robinson A, Dixon NE, van Oijen AM (2019) Recycling of single-stranded DNA-binding protein by the bacterial replisome. Nucleic Acids Res 47:4111–4123. https:// doi.org/10.1093/nar/gkz090 8. Lewis JS, Spenkelink LM, Jergic S, Wood EA, Monachino E, Horan NP, Duderstadt KE, Cox MM, Robinson A, Dixon NE, van Oijen AM (2017) Single-molecule visualization of fast polymerase turnover in the bacterial replisome. eLife 6:e23932. https://doi.org/10.7554/ eLife.23932 9. Geertsema HJ, Kulczyk AW, Richardson CC, van Oijen AM (2014) Single-molecule studies of polymerase dynamics and stoichiometry at the bacteriophage T7 replication machinery. Proc Natl Acad Sci U S A 111 (11):4073–4078. https://doi.org/10.1073/ pnas.1402010111 10. Beattie TR, Kapadia N, Nicolas E, Uphoff S, Wollman AJ, Leake MC, Reyes-Lamothe R (2017) Frequent exchange of the DNA polymerase during bacterial chromosome replication. eLife 6:e21763. https://doi.org/10. 7554/eLife.21763 11. Loparo JJ, Kulczyk AW, Richardson CC, van Oijen AM (2011) Simultaneous singlemolecule measurements of phage T7 replisome

Single-Molecule Fluorescence Visualization of Protein Exchange composition and function reveal the mechanism of polymerase exchange. Proc Natl Acad Sci U S A 108(9):3584–3589. https://doi. org/10.1073/pnas.1018824108 12. Liao Y, Li Y, Schroeder JW, Simmons LA, Biteen JS (2016) Single-molecule DNA polymerase dynamics at a bacterial replisome in live cells. Biophys J 111(12):2562–2569. https:// doi.org/10.1016/j.bpj.2016.11.006 13. Aberg C, Duderstadt KE, van Oijen AM (2016) Stability versus exchange: a paradox in DNA replication. Nucleic Acids Res 44 (10):4846–4854. https://doi.org/10.1093/ nar/gkw296 14. Sing CE, Olvera de la Cruz M, Marko JF (2014) Multiple-binding-site mechanism explains concentration-dependent unbinding rates of DNA-binding proteins. Nucleic Acids Res 42(6):3783–3791. https://doi.org/10. 1093/nar/gkt1327 15. Mueller SH, Spenkelink LM, van Oijen AM (2019) When proteins play tag: the dynamic nature of the replisome. Biophys Rev 11 (4):641–651. https://doi.org/10.1007/ s12551-019-00569-4 16. van Oijen AM, Duderstadt KE, Xiao J, Fishel R (2018) Plasticity of multi-protein complexes. J Mol Biol 430(22):4441–4442. https://doi. org/10.1016/j.jmb.2018.08.008 17. van Oijen AM, Dixon NE (2015) Probing molecular choreography through singlemolecule biochemistry. Nat Stuct Mol Biol 22 (12):948–952. https://doi.org/10.1038/ nsmb.3119

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18. Tanner NA, Loparo JJ, Hamdan SM, Jergic S, Dixon NE, van Oijen AM (2009) Real-time single-molecule observation of rolling-circle DNA replication. Nucleic Acids Res 37(4): e27. https://doi.org/10.1093/nar/gkp006 19. Monachino E, Ghodke H, Spinks RR, Hoatson BS, Jergic S, Xu ZQ, Dixon NE, van Oijen AM (2018) Design of DNA rolling-circle templates with controlled fork topology to study mechanisms of DNA replication. Anal Biochem 557:42–45. https://doi.org/10.1016/j.ab. 2018.07.008 20. Geertsema HJ, Duderstadt KE, van Oijen AM (2015) Single-molecule observation of prokaryotic DNA replication. Methods Mol Biol 1300:219–238. https://doi.org/10.1007/ 978-1-4939-2596-4_14 21. Kaur G, Lewis JS, van Oijen AM (2019) Shining a spotlight on DNA: single-molecule methods to visualise DNA. Molecules 24 (3):491. https://doi.org/10.3390/ molecules24030491 22. Tinevez J-Y, Perry N, Schindelin J, Hoopes GM, Reynolds GD, Laplantine E, Bednarek SY, Shorte SL, Eliceiri KW (2017) TrackMate: an open and extensible platform for singleparticle tracking. Methods 115:80–90. https://doi.org/10.1016/j.ymeth.2016.09. 016 23. Axelrod D, Koppel DE, Schlessinger J, Elson E, Webb WW (1976) Mobility measurement by analysis of fluorescence photobleaching recovery kinetics. Biophys J 16 (9):1055–1069

Chapter 4 Comparing SSB-PriA Functional and Physical Interactions in Gram-Positive and -Negative Bacteria Yen-Hua Huang and Cheng-Yang Huang Abstract Single-stranded DNA (ssDNA)-binding protein (SSB) is essential for DNA metabolic processes. SSB also binds to many DNA-binding proteins that constitute the SSB interactome. The mechanism through which PriA helicase, an initiator protein in the DNA replication restart process, is stimulated by SSB in Escherichia coli (EcSSB) has been established. However, some Gram-positive bacterial SSBs such as Bacillus subtilis SsbA (a counterpart of EcSSB), Staphylococcus aureus SsbA, SsbB, and SsbC do not activate PriA helicase. Here, we describe some of the methods used in our laboratory to compare SSB-PriA functional and physical interactions in Gram-positive and -negative bacteria. Key words SSB, ssDNA-binding protein, PriA, SsbA, Protein-protein interaction, ConSurf, SPR, ATPase, Stimulation

1

Introduction Single-stranded DNA-binding proteins (SSBs) are ubiquitous within all kingdoms of life and play essential roles in DNA replication, recombination, repair, and replication restart [1]. SSB exhibits high affinity for ssDNA with no considerable sequence specificity. In addition, SSB also binds to many nucleoproteins and enzymes associated with DNA metabolism that constitute the SSB interactome [2]. SSB is typically a homotetramer, in which four oligonucleotide/oligosaccharide-binding folds (OB folds) form a conserved N-terminal ssDNA-binding/oligomerization domain. A flexible, highly disordered C-terminal domain responsible for protein–protein interaction can be further subdivided into two sub-domains: a long proline- or glycine-rich hinge, also known as the intrinsically disordered linker, and the highly conserved acidic tail of the last six C-terminal amino acid residues of SSB (DDDIPF).

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_4, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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PriA is utilized during replication restart to reload the replicative DnaB helicase back onto the chromosome [3]. In Escherichia coli, the replication restart primosome consists of PriA, PriB, PriC, DnaB helicase, DnaC, DnaT, and DnaG primase. PriA recognizes stalled DNA replication forks with either duplex or SSB-coated ssDNA lagging strands and then processes for full primosome assembly [4]. PriA is a poor helicase but its activity can be stimulated by a physical contact with the C-terminus of SSB [5]. The complex crystal structure of Klebsiella pneumoniae PriA reveals that five residues, namely Trp82, Tyr86, Lys370, Arg697, and Gln701, participate in SSB recognition [6]. Different bacteria have different strategies for the functional loading of cellular replicative DNA helicases. Unlike E. coli, which contains only one type of SSB (EcSSB), some bacteria have more than one paralogous SSB. In Staphylococcus aureus, three SSBs are found, namely SaSsbA, SaSsbB, and SaSsbC. In addition, essential helicase-loading components PriB, PriC, DnaT, and DnaC proteins are not found in Bacillus subtilis [7] and S. aureus [8]. Instead, three other proteins DnaD, DnaB, and DnaI have been proven to be required for the replication restart of the Gram-positive bacteria. Similar to the case of EcSSB-EcPriA, SaDnaD can bind and significantly stimulate the ATPase activity of SaPriA. However, B. subtilis SsbA [9], SaSsbA [10], SaSsbB [11], and SaSsbC [12] do not activate SaPriA helicase. Whether SSB functions and participates in the Gram-positive bacterial PriA-directed primosome assembly in a manner different from that of Gram-negative E. coli should be confirmed. In this chapter, we focus on some of the assays used in our laboratory to assess SSB-PriA functional and physical interactions in Gram-positive and -negative bacteria (see Note 1). The alignment consensus of 417 sequenced PriA homologs by ConSurf [13] revealed that the conserved SSB binding site of KpPriA is not present in SaPriA. Only Trp88 in SaPriA (Trp82 in KpPriA) is conserved. The assays of surface plasmon resonance (SPR) and ATPase stimulation presented here had been used for analyzing different SSB-PriA physical and functional interactions. Results from these assays have indicated that the case of EcPriA-EcSSB is not applicable to SaPriA-SaSsbA because of inherent differences among the species.

2

Materials 1. The ConSurf web server: http://consurf.tau.ac.il. 2. Biacore T200 SPR instrument. 3. CM5 sensor S chips.

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4. HEPES running buffer (see Note 2): 20 mM HEPES, 200 mM NaCl, and 0.05% Tween-20 at pH 7.4. 5. Tris running buffer: 40 mM Tris, 200 mM NaCl, and 0.05% Tween-20 at pH 8.0. 6. Immobilization buffer 4.0: 10 mM Sodium acetate at pH 4.0. 7. Immobilization buffer 4.5: 10 mM Sodium acetate at pH 4.5. 8. Immobilization buffer 5.0: 10 mM Sodium acetate at pH 5.0. 9. Immobilization buffer 5.5: 10 mM Sodium acetate at pH 5.5. 10. 500 mL Bottle-top vacuum filter, 0.22 μm. 11. Syringe filter, 0.22 μm. 12. Amine coupling kit: 750 mg 1-Ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC), 115 mg N-hydroxysuccinimide (NHS), 10.5 mL ethanolamine-HCl (1.0 M at pH 8.5). 13. 0.4 M EDC. 14. 0.1 M NHS. 15. 1.0 M Ethanolamine-HCl. 16. Peptide SaSsbA-Ct: NANGPIDISDDDLPF (15 mer), solubilized with Tris running buffer before experiment. 17. Peptide KpSSB-Ct: PSNEPPMDFDDDIPF (15 mer), solubilized with Tris running buffer before experiment. 18. The double-stranded DNA substrate, prepared with oligos PS4 (30 -GGGCTTAAGCCTATCGAGCCATGGG-50 , 25 mer) and PS3-dT30 (50 - CCCGAATTCGGATAGCTCGGTACCC dT30-30 , 55 mer) at a 1:1 concentration ratio, in 20 mM HEPES (pH 7.0) and 100 mM NaCl, by brief heating at 95  C for 5 min followed by slow cooling to room temperature overnight. 19. SaPriA, purified as described in [14]. 20. SaSsbA, purified as described in [10]. 21. KpSSB, purified as described in [15]. 22. 5 Reaction buffer: 200 mM Tris–HCl, 50 mM NaCl, 10 mM DTT, 0.5 mM EDTA, and 7.5 mM MgCl2 at pH 8.0. 23. 5 ATP reaction solution: Add 1 μL of 100 mM ATP and 1 μL of [γ-32P]ATP (6000 Ci/mmol) in 48 μL Milli-Q water. 24. 5 dsDNA solution: 0.5 μM PS4/PS3-dT30 solution. 25. HEPES dialysis buffer: 20 mM HEPES and 100 mM NaCl, pH 7.0. 26. TLC solvent: 25 mL of 0.5 M formic acid and 0.25 M LiCl, freshly prepared before experiments.

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27. TLC plate: Cellulose MN 300 PEI, PEI-impregnated cellulose ion-exchange layers. 28. BAS-2500 bio-imaging analyzer. 29. Phosphorimager IP (Imaging Plate BAS-IIIS, 20  40 cm). 30. Software Image Reader BAS-2500 and Image Gauge.

3 3.1

Methods ConSurf Analysis

Based on the known nucleotide sequence, the predicted SaPriA monomer protein has a length of 802 amino acid residues (which is higher than that of EcPriA with 732 amino acid residues) and a molecular mass of 92.7 kDa (Fig. 1). SaPriA may differ from EcPriA in terms of sequence and structure (see Note 3). Here we describe the procedure for sequence analysis using ConSurf [13]. Starting from a query sequence, the server automatically collects homologs, infers their multiple sequence alignment, and reconstructs a phylogenetic tree that reflects their evolutionary relations. 1. Access the ConSurf web server (http://consurf.tau.ac.il). 2. Check “Analyze Nucleotides or Amino Acids?”, and choose Amino Acids. 3. Check “Is there a known protein structure?”, and choose NO. 4. Check “Do you have a Multiple Sequence Alignment (MSA) to upload? (If not, ConSurf will make an MSA for you.)”, and choose NO. 5. Paste the protein sequence in FASTA format (see Note 4). For SaPriA, the protein sequence can be found at https://www. ncbi.nlm.nih.gov/protein/446482985. 6. Choose parameters for homolog search algorithm: Homolog search algorithm: HMMER Number of iterations: 1 E-value cutoff: 0.0001 Protein database: UNIREF-90 Select homologs for ConSurf analyses: automatically (recommended). 7. Choose parameters to select the sequences for the analysis out of homolog search algorithm results. Select “417” sequences “that sample the list of homologues” to reference sequence. Maximal % ID between sequences: 95. Minimal % ID for homologs: 35. 8. Choose alignment method to build the multiple sequence alignment (MSA):

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Fig. 1 Sequence analysis of PriA. (a) Sequence analysis of SaPriA using ConSurf, http://consurf.tau.ac.il. (b) An alignment consensus of 417 sequenced PriA homologs by ConSurf reveals the degree of variability at each position along the sequence [10]. Highly variable amino acid residues are colored teal, whereas highly conserved amino acid residues are burgundy. A consensus sequence was established by determining the most commonly found amino acid residue at each position relative to the primary sequence of SaPriA. The putative SSB binding sites in SaPriA structurally corresponding with those in KpPriA are indicated by asterisks. These sites are not conserved in SaPriA. (c) The SSB binding sites in KpPriA are Trp82, Tyr86, Lys370, Arg697, and Gln701. (d) The putative SSB binding sites in SaPriA are Trp88, Thr92, V439, Glu767, and Leu771

Alignment method: MAFFT-L-INS-i. 9. Calculation method: Bayesian. 10. Evolutionary substitution model: Best model (default). 11. Enter a descriptive job title for your ConSurf query: SaPriA. 12. Enter the e-mail. The ConSurf web server will use this address to report you when the job has finished or if error has occurred.

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13. Click “Send a link to the results by e-mail”. 14. Select Submit. You will be redirected to ConSurf Job Status page. 15. ConSurf will be processed. The running progress includes the following: (1) find sequence homologs, (2) align sequences, (3) select best evolutionary model, (4) calculate conservation scores, (5) search for 3D structure for the protein sequence, and (6) project conservation scores onto the molecule. 16. When the ConSurf calculation is finished, you will receive running messages similar to these: “There are 11472 HMMER hits. 11267 of them are unique, including the query. The calculation is performed on a sample of 417 sequences that represent the list of homologues to the query”. 17. The final results can be downloaded, including (1) the query sequence colored according to the conservation scores (HTML) (PDF); (2) multiple sequence alignment colorcoded by conservation; (3) viewing of MSA and phylogenetic tree using WASABI and running of ConSurf on sub-tree; (4) multiple sequence alignment color-coded by conservation and the (neighbor-joining) ConSurf tree (download Chimera; required); and (5) amino acid conservation scores, confidence intervals, and conservation colors. Other information such as sequence data, alignment, and phylogenetic tree are also available. 3.2

SPR

3.2.1 Preparation of Reagents and Samples

SPR-based instruments allow real-time monitoring of molecular interactions [16]. Given that the binding site of KpPriA to SSB is not found in SaPriA, we used SPR to confirm whether SaPriA interacts with SaSsbA. Chemically synthesized peptides SaSsbA-Ct and KpSSB-Ct were also used for SaPriA-binding experiments. Here we describe the procedure for analyzing these physical interactions of SaPriA using SPR. The results showed that SaPriA can bind to SaSsbA, but cannot bind to SaSsbA-Ct and KpSSB-Ct. 1. Dialyze SaPriA protein solution in 1 L of HEPES running buffer on a stirrer at 4  C for 8 h (see Note 4). HEPES running buffer is changed to a fresh one twice with 8-h interval. 2. Dialyze SaSsbA protein solution in 1 L of Tris running buffer on a stirrer at 4  C for 8 h. Tris running buffer is changed to a fresh one twice with 8-h interval. 3. Determine protein concentration by standard BSA analysis. 4. Start the “Biacore T200 control software”. 5. Check that the waste bottle is empty (right tray). 6. Check that the Milli-Q water bottle is filled (right tray).

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7. Set the temperatures (default is 25  C). Select Tools: Set Temperature. 8. Undock the maintenance chip. Have a quick inspection of the sensor square and store it in its dedicated plastic box. Select Tools: Eject Chip. 9. Dock CM5 sensor S chip into the SPR instrument. Insert the cartridge into the docking bay by following the arrows. Close manually the compartment door. In the Insert Chip window, select New Chip, and in Chip Type menu choose the one corresponding to your sensor chip (CM5). Select Dock Chip. 10. Exchange the water bottle connected to line A (left tray) with 1 L of HEPES running buffer bottle. 11. Keep fresh Milli-Q water (500 mL) in right tray (line B). 12. Prime the system with HEPES running buffer. Select Tools: Prime. Select Start. 3.2.2 Immobilization of SaPriA

1. Prepare 300 μL of ligand solutions. SaPriA (10 μg/mL) was prepared by dilution with the different immobilization buffers at different pH values, 4.0, 4.5, 5.0, and 5.5 (see Note 5). 2. Inject ligand solutions and determine the optimum pH condition in which the highest response level is observed. The immobilization buffer 5.0 was chosen for immobilization of SaPriA (see Note 6). 3. Choose the File menu: “Open New wizard/template” and then “Immobilization”. Click “New”. 4. Select immobilize flow cells (1, 2) and enter (see Note 7). Method: Amine. Ligand: SaPriA. Check “Aim for immobilized level”. Target level: 950 RU. Wash solution: 50 mM NaOH. 5. Prime before run. 6. Analysis temperature: 25  C. 7. Sample compartment temperature: 4  C. 8. For immobilizing flow cell 2: 99 μL EDC to activate the surface. 99 μL NHS to introduce maleimide groups. 139 μL Ethanolamine to deactivate excess reactive groups. 166 μL SaPriA. 68 μL 50 mM NaOH to remove the last traces of electrostatically bound ligand. Mix EDC and NHS in an empty vial automatically.

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9. Choose “Eject rack” and place the vials according to the setup in the software. “Close” and “Next”. 10. Check that HEPES running buffer and water levels will be sufficient for the run. Click “Next”. 11. Save the wizard and click “Start” (see Note 8). 3.2.3 Binding of SaSsbA, SaSsbA-Ct, and KpSSB-Ct

1. Exchange the HEPES running buffer bottle connected to line A (left tray) with 1 L of Tris running buffer bottle (see Note 9). 2. Prime the system with this running buffer. Select Tools: Prime. Select Start. 3. Prepare 1 mL of 1000 nM SaSsbA protein solution by diluting with Tris running buffer (see Note 10). 4. Select Run. Click Manual Run. 5. Inject SaSsbA and check the interaction with SaPriA. 6. Run a sensorgram on one flow cell at 30 μL/min. 7. Select Flow Cell (1, 2). Select the rack type (Sample & Reagent Rack 1). Determine SaSsbA condition by injecting 60 μL of 2 M MgCl2 (or 1 M NaCl) (see Note 11). Select Start. 8. Select File. Open/New Method. Set flow rate: 30 μL/min. Contact time (s): 120. Dissociation (s): 300. 9. Set regeneration: 2 M MgCl2. Flow rate: 30 μL/min. Contact time (s): 60. 10. Dilute SaSsbA solution in Tris running buffer to final concentrations of 1000, 500, 250, 125, and 63 nM. 11. Chemically synthesized peptides SaSsbA-Ct and KpSSB-Ct were also diluted in Tris running buffer to final concentrations of 2000, 1500, 1000, 500, 250, 125, and 63 nM. 12. Choose “Eject rack” and place the vials. Then “Close” and “Next”. 13. Check that Tris running buffer and water levels will be sufficient for the run. Click “Next”. 14. Save the wizard and click “Start”.

3.2.4 Analysis of the Sensorgrams

The estimated dissociation equilibrium constants (Kd values) can be derived by steady-state analysis by fitting the association and dissociation signals with a 1:1 (Langmuir) model using the Biacore T200 Evaluation Software (see Note 12). The Kd value of SaPriA

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Fig. 2 SPR analysis of SaPriA interacting with SaSsbA. SaPriA was immobilized on the CM5 chip, and the binding experiments were carried out using a Biacore T200 [10]. SaSsbA (1000, 500, 250, 125, and 63 nM) was injected over the immobilized protein for 120 s at a flow rate of 30 μL/min. The estimated Kd value was derived by fitting the association and dissociation signals with a 1:1 (Langmuir) model using the Biacore T200 Evaluation Software

bound by SaSsbA, calculated from the equilibrium binding isotherms, was 4.6  0.5  108 M (Fig. 2). However, the binding response for SaPriA did not change when injecting SaSsbA-Ct or KpSSB-Ct. Accordingly, we concluded that SaPriA can bind to SaSsbA, but cannot bind to SaSsbA-Ct and KpSSB-Ct. 3.3 ATPase Stimulation Assay

Although the binding site of KpPriA to SSB is not found in SaPriA (Fig. 1), SaPriA still can bind to SaSsbA through an unknown mechanism (Fig. 2). ATPase stimulation assay using thin-layer chromatography (TLC) was further used for analyzing different SSB-PriA functional interactions. TLC is a direct and flexible method for detecting ATPase activity using radiolabeled ATP. Here we describe the procedure for analyzing the ATPase stimulation effect of SaPriA using TLC. The results showed that KpSSB, but not SaSsbA, can stimulate the ATPase activity of SaPriA (Fig. 3).

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Fig. 3 ATPase stimulation assay. (a) TLC is used for analyzing different SSB-PriA functional interactions. Mark the origin line for the reaction spots. (b) The chamber (beaker) is used for developing a TLC plate. (c) SaPriA ATPase assay was performed with 0.4 mM [γ-32P]ATP, 0.12 μM SaPriA, and 0.1 μM PS4/PS3-dT30 DNA substrate for 1 h. To study the effect, 10 μM SaSsbA and 10 μM KpSSB were individually added into the assay solution. Aliquots were taken and spotted onto a TLC plate, which was subsequently developed in 0.5 M formic acid and 0.25 M LiCl for 15 min. Reaction products were visualized by autoradiography and quantified with a phosphorimager

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1. Dilute SaPriA solution using HEPES dialysis buffer to a final concentration of 0.6 μM. 2. Dilute SaSsbA and KpSSB solutions using HEPES dialysis buffer to a final concentration of 50 μM. 3. Mix 1 μL of 5 reaction buffer, 1 μL of 0.6 μM SaPriA solution, 1 μL of HEPES dialysis buffer (or 1 μL of 50 μM SaSsbA/KpSSB), 1 μL of 5 ATP reaction solution, and 1 μL of 5 dsDNA solution. Mix these solutions by pipetting up and down gently. Avoid introducing bubbles. 4. Transfer the reaction tubes to a 37  C heat block and incubate them for 1 h (see Note 13). 5. Spot 2–5 μL of each reaction mixture onto TLC plate in a straight line at least 1.5 cm away from the bottom edge (see Note 14). After spotting, dry the TLC plates in the oven. 6. Transfer the TLC plates to the TLC solvent-containing 1 L beaker using tweezers. 7. Cover the beaker with plastic wrap and develop until the front of the liquid phase reaches the top of the TLC plates (see Note 15). 8. Dry the TLC plates thoroughly in the oven. Wrap the TLC plates with plastic wrap and expose them to imaging plate (see Note 16). 9. Power on FUJIFILM BAS-2500 bio-imaging analyzer and the computer. The reader will take several minutes to go through its initialization routine. 10. The Image Reader BAS-2500 software: Sampling area: free. Image area: full grid (1–8 X A-P) for large plates. Gradation: 65536 (16 bit). Resolution: 200. Dynamic range: 10,000. 11. Load the exposed image plate into the reader. The Phosphorimager IP (20  40 cm) is placed directly into the reader. 12. Click the “Read” button and select the destination folder. 13. After the reader reads the image plate, the software will save and display the image. 14. Export your files and/or folders after you finish data. 15. Quantify an image using Image Gauge software in Quant mode. 16. Select the desired region where you want to measure. 17. Select Quant Result from the Windows menu. Quantification results will be displayed.

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Notes 1. B. subtilis SsbA [9], S. aureus SsbA [10], SsbB [11], and SsbC [12] did not affect the activity of PriA. Based on these results, one might conclude that Gram-positive bacterial SSBs cannot stimulate PriA activity. In fact, however, we also found that the Gram-negative Pseudomonas aeruginosa SSB (PaSSB) did not enhance the PriA activity, probably due to the different glycinerich flexible region [17]. The region in PaSSB is indeed not glycine rich [17]. Given that the ssDNA-binding mode of PaSSB recently shown by the complex structures [18, 19] differs significantly from that of EcSSB, whether EcSSB, SaSsbA, and PaSSB function and participate in the PriA-directed primosome assembly in a similar manner [8] needs to be confirmed by further biochemical, structural, and cellular experiments. 2. All solutions were prepared with reagent-grade chemicals and Milli-Q water that was purified by reverse osmosis and subsequently treated with the Millipore reagent water system to attain a sensitivity of 18 MΩ-cm at 25  C. Prepare and store all reagents at room temperature (unless indicated otherwise). 3. Unlike EcPriA, SaPriA is quite different in terms of the length, amino acid sequence, and SSB binding site. The alignment consensus of 417 sequenced PriA homologs by ConSurf revealed that amino acid residues in the SSB binding site and the N-terminal region of SaPriA, especially aa 114–286, are variable. 4. Prepare systematically fresh buffers and never use solutions more than 48 h. Clean carefully the instrument before running the experiments. Solutions need to be filtered (0.22 μm) and contained in glass bottles. Ligand and analyte samples must be filtered (0.22 μm) before experiments. 5. Protein was covalently linked according to the thiol ligand coupling strategy. Different sodium acetate buffers (10 mM at pH 4.0, 4.5, 5.0, and 5.5) were selected separately as immobilization buffers using the pH scouting procedure. 6. When there is little or no difference, select higher pH in order to avoid ligand deterioration and to promote coupling reaction. 7. The Biacore T200 has four flow cells (Fc). Each experiment uses minimum two flow cells. Fc1 and Fc3 are used as references (with no immobilized ligand or dummy ligand). Fc2 and Fc4 would be the functionalized flow cells. 8. If you want to stop the experiment in this step, eject the sensor chip from the instrument and store it at 4  C in HEPES running buffer to prevent it from drying out.

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9. After testing, we found that the binding system of SaPriA using Tris running buffer is much more stable than that using HEPES running buffer. 10. The SaSsbA solution should not include any reagents which perturb the interaction between SaSsbA and SaPriA. 11. We found that 2 M MgCl2 could fully wash out SaSsbA from the complex of SaPriA-SaSsbA for reuse of the SaPriAimmobilized chip. 12. We determined the Kd values of SaPriA for SaSsbA using a 1:1 Langmuir binding model. However, how SaPriA monomer can bind to SaSsbA tetramer needs to be investigated by further biochemical and cellular experiments. 13. During incubation, freshly prepare the TLC solvent by mixing 12.5 mL of 1 M formic acid and 12.5 mL of 0.5 M LiCl in a 1 L beaker. Cover the beaker with plastic wrap to prevent evaporation. Usually, it will take 20–30 min to saturate the air in the beaker (the developing chamber) with solvent molecules. 14. Use a soft pencil to mark the origin line (the line you will spot the samples). 15. For developing in 0.5 M formic acid and 0.25 M LiCl, the liquid phase will take 15–20 min to reach the top of the TLC plates. 16. 10–20-min exposure is sufficient for phosphorimager image.

Acknowledgment This work was supported by a grant from the Ministry of Science and Technology, Taiwan (MOST 108-2320-B-040-010 to C.Y. Huang). References 1. Antony E, Lohman TM (2019) Dynamics of E. coli single stranded DNA binding (SSB) protein-DNA complexes. Semin Cell Dev Biol 86:102–111 2. Bianco PR (2017) The tale of SSB. Prog Biophys Mol Biol 127:111–118 3. Windgassen TA, Wessel SR, Bhattacharyya B, Keck JL (2018) Mechanisms of bacterial DNA replication restart. Nucleic Acids Res 46:504–519 4. Windgassen TA, Leroux M, Satyshur KA, Sandler SJ, Keck JL (2018) Structure-specific DNA replication-fork recognition directs helicase and replication restart activities of the PriA

helicase. Proc Natl Acad Sci U S A 115: E9075–e9084 5. Cadman CJ, McGlynn P (2004) PriA helicase and SSB interact physically and functionally. Nucleic Acids Res 32:6378–6387 6. Bhattacharyya B, George NP, Thurmes TM, Zhou R, Jani N, Wessel SR, Sandler SJ, Ha T, Keck JL (2014) Structural mechanisms of PriA-mediated DNA replication restart. Proc Natl Acad Sci U S A 111:1373–1378 7. Bruand C, Ehrlich SD, Janniere L (1995) Primosome assembly site in Bacillus subtilis. EMBO J 14:2642–2650

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8. Huang YH, Huang CY (2014) Structural insight into the DNA-binding mode of the primosomal proteins PriA, PriB, and DnaT. Biomed Res Int 2014:195162 9. Polard P, Marsin S, McGovern S, Velten M, Wigley DB, Ehrlich SD, Bruand C (2002) Restart of DNA replication in Gram-positive bacteria: functional characterisation of the Bacillus subtilis PriA initiator. Nucleic Acids Res 30:1593–1605 10. Huang YH, Guan HH, Chen CJ, Huang CY (2017) Staphylococcus aureus single-stranded DNA-binding protein SsbA can bind but cannot stimulate PriA helicase. PLoS One 12: e0182060 11. Chen KL, Cheng JH, Lin CY, Huang YH, Huang CY (2018) Characterization of single-stranded DNA-binding protein SsbB from Staphylococcus aureus: SsbB cannot stimulate PriA helicase. RSC Adv 8:28367–28375 12. Huang YH, Huang CY (2018) SAAV2152 is a single-stranded DNA binding protein: the third SSB in Staphylococcus aureus. Oncotarget 9:20239–20254 13. Ashkenazy H, Erez E, Martz E, Pupko T, Ben-Tal N (2010) ConSurf 2010: calculating evolutionary conservation in sequence and structure of proteins and nucleic acids. Nucleic Acids Res 38:W529–W533

14. Huang YH, Huang CC, Chen CC, Yang KJ, Huang CY (2015) Inhibition of Staphylococcus aureus PriA helicase by flavonol kaempferol. Protein J 34:169–172 15. Huang YH, Huang CY (2012) Characterization of a single-stranded DNA-binding protein from Klebsiella pneumoniae: mutation at either Arg73 or Ser76 causes a less cooperative complex on DNA. Genes Cells 17:146–157 16. Jonsson U, Fagerstam L, Ivarsson B, Johnsson B, Karlsson R, Lundh K, Lofas S, Persson B, Roos H, Ronnberg I (1991) Realtime biospecific interaction analysis using surface plasmon resonance and a sensor chip technology. BioTechniques 11:620–627 17. Huang YH, Huang CY (2018) The glycinerich flexible region in SSB is crucial for PriA stimulation. RSC Adv 8:35280–35288 18. Huang YH, Lin ES, Huang CY (2019) Complexed crystal structure of SSB reveals a novel single-stranded DNA binding mode (SSB)3:1: Phe60 is not crucial for defining binding paths. Biochem Biophys Res Commun 520:353–358 19. Huang YH, Chen IC, Huang CY (2019) Characterization of an SSB-dT25 complex: structural insights into the S-shaped ssDNA binding conformation. RSC Adv 9:40388–40396

Chapter 5 In Vivo Binding of Single-Stranded DNA-Binding Protein to Stalled Replication Fork Helicases Cong Yu and Piero R. Bianco Abstract Understanding protein-protein interactions is key to unraveling protein function in vivo. Here we describe a dual/triple-plasmid system that enables co-expression of two, or three, recombinant proteins harboring different affinity tags in the same Escherichia coli cell. This novel protein expression system provides a platform to understand protein-protein interactions and enables researchers to study protein complex formation and in vivo localization. Key words Stalled fork rescue, Single-strand binding protein, SSB, DNA helicase, RecG, PriA

1

Introduction Escherichia coli is one of the most widely used model organisms, selected for its rapid growth and stable production of recombinant proteins to investigate cellular and molecular mechanisms in vivo and in vitro. In this chapter, we describe a novel technique to study protein-protein interactions in vivo. In short, we co-transform two or three recombinant plasmids into the same E. coli strain, induce expression, and detect potential protein interactions by SDS-PAGE following affinity chromatography. To achieve complex formation, one of the partners is cloned into an expression vector to add a tag either at its N- or C-terminus to allow binding to the affinity column. Depending on the protein complexes, sequential affinity chromatography can be used to examine complex formation using differential tagging of complex components. The level of protein binding and elution depends on several factors that include but not limited to the copy number of the expression vector, columnbinding capacities, salt concentration, and stoichiometry ratio of proteins within the complex. The single-stranded DNA-binding protein (SSB) and the DNA helicases RecG and PriA are partner proteins that act on stalled

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_5, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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replication forks to facilitate their rescue [1–7]. To understand the timing of protein loading onto the DNA fork, and thus to unravel the mechanism of how stalled replication forks are restarted, we constructed co-expression strains containing two or three separate vectors to generate the SSB-RecG, SSB-PriA, or SSB-RecG-PriA complexes [8]. For dual-expression strains, the detection of protein complexes in the eluted fractions is a result of interactions between the tagged and untagged partners. For the three-protein complex, either two or all three proteins are cloned with different tags to enable complex purification using three affinity columns sequentially. For the dual- and triple-tagged complexes, chromatography is followed by SDS-PAGE to determine the presence of one or more proteins in the eluted complexes (Fig. 1). Using the single-tagging approach, we observed SSB-RecG complexes when the histidine-tag was present on either RecG or SSB. When the histidine-tag was on SSB, a significantly higher yield of protein complex was eluted from the nickel column due to the availability of four RecG-binding sites per his-SSB tetramer, versus only one in his-RecG. However, when the histidine-tag was on either the N- or C-terminus of RecG, there was no difference in the amount of protein complex eluted from the column, indicating that the tag did not impair RecG protein folding or its column binding affinity.

Fig. 1 PriA and RecG bind to SSB in vivo. 12% SDS-PAGE gels of the purification of helicase-SSB complexes. Panels (a–c), co-purification of RecG and SSB; panels (d–f), co-purification of PriA and SSB. In (a) and (d), purification was done with buffer containing 600 mM NaCl. In panels (b) and (e), the final concentration of NaCl was 100 mM. Panel (c), purification of the his-RecG/Profinity-SSB complex; left panel, protein gel stained with Coomassie Brilliant Blue; right panel, protein gel stained with silver. Panel (f), purification of the his-SSB/PriAbiotin complex. The numbers at the top of each lane indicate fraction numbers from the peak in the corresponding elution profiles. M, molecular weight marker; CCL, clear cell lysate; FT, flow-through; Ni, pooled fractions from the nickel column; Av, pooled fractions from the avidin column; Pro, complex eluted from the Profinity eXact column (reproduced from [8] with permission from Genes to Cells)

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An unexpected benefit of using the co-expression system is that it can provide insight into protein stability provided by one of the interacting partners. For example, a typical RecG purification yields 1–2 mg of protein per liter of cells [9]. By using the dual-expression system, we consistently obtained more than 200 mg of RecG-SSB complexes from a single liter of cells with the helicase comprising at least 1/3 of the total protein. This surprising result suggested that SSB stabilizes the RecG helicase. Importantly, the purified complexes do not contain DNA when assessed using radioactive labeling. This is consistent with the proteins binding directly to each other to form a complex in vivo. Further evidence for complex formation came from mixing studies. Here, RecG and his-SSB were expressed in different strains and cell lysates were mixed just before loading onto the nickel column. The results showed that no complex eluted. Instead, only his-SSB was present in the eluted fractions, while RecG was present in the flowthrough and wash fractions. Thus, co-elution of co-expressed partners is consistent with complex formation occurring in vivo, before cell lysis. For strains harboring three expression plasmids, either two of the three plasmids or all three plasmids can contain different tags. For example, we have used histidine, biotin, and Profinity tags in various combinations to detect the triple-protein complex of RecGhis/Profinity-SSB/PriA-biotin [8, 10]. The proteins eluted from the first affinity column are loaded onto a second affinity column to achieve two sequential purifications steps. A third affinity column matching the final tag can be used if necessary. To verify complex size, the proteins harvested from the final affinity column can be applied to a gel filtration column with the elution profile compared to that of a standard curve [11]. The co-expression system is also a useful tool to determine protein intracellular location and co-localization by inserting autofluorescent protein genes into the recombinant plasmids (Fig. 2). Unlike co-expression followed by co-purification, expression of autofluorescent protein fusions is typically done in the absence of IPTG to reduce background noise [8, 12, 13]. Alternatively, fusion genes can be introduced into the chromosome using recombineering to attain a copy number of 1–2 genes per cell [14]. In this strain, that is, SSB-GFP, subunit mixing with wild-type subunits occurs, producing tetramers with different ratios of wild type:fusion protein [12, 13]. This produces an average ratio of SSB:GFP of 7.3 in LB + glucose as assessed by quantitative western blots [14].

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Fig. 2 SSB localizes PriA and RecG to the cell periphery. Representative images of cells co-expressing GFP-PriA, mCherry-RecG, and wild-type SSB. Panel (a) DIC; (b) fluorescence image captured with the GFP filter (PriA); (c) fluorescence image captured with the mCherry filter (RecG); (d) colocalization image of panels (b) and (c) (reproduced from [8] with permission from Genes to Cells)

2

Materials Prepare all solutions using reagent-grade chemicals and Nanopure water. Filter each solution through 0.2 μm pore size filters. Store at room temperature or 4
C. 1. Recombinant plasmids for protein expression: See Table 1 for a complete list. Plasmids can be provided upon request. 2. Hi-Trap nickel column (5 mL). 3. Monomeric avidin resin (5 mL). 4. Profinity eXact column (5 mL). 5. Superose 6, 10/300 GL column. 6. Tuner (λDE3) E. coli strain (see Note 1). 7. 2XYT: For 1 L 2XYT, dissolve 16 g Bacto Tryptone, 10 g yeast extract, 5 g NaCl, and 100 μL of 10 N NaOH in Nanopure water; autoclave to make basal medium; and add 10 mL of 1 M MgSO4 before use.

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Table 1 Plasmids used in this technique Protein Linker/tag

Vector

Cloning site

RecG

pET-28a (+) pET-28a (+) pET-28a (+)

Between NdeI and BamHI sites

N-terminal his6-tag C-terminal his6-tag N-terminal his-mcherry

PriA

SSB

N-terminal his6-linker

pET-28a (+) C-terminal his6-linker pET-28a (+) C-terminal biotinylation pET-28a (+) Low copy number his-GFP- pWKS30 PriA N-terminal Profinity tag N-terminal his6-linker C-terminal GFP C-terminal-YFP C-terminal-mcherry

pPAL7 pET-15b pET21a (+) pET21a (+)

Between NcoI and HindIII sites Insert mcherry into NdeI site Between ApaI and EcoRI sites Between NcoI and HindIII sites Between HindIII and XhoI sites Between BamHI and BglII sites Between SpeI and BamHI sites Between NdeI and BamHI sites GFP at C-terminal XhoI site YFP at C-terminal XhoI site mcherry at C-terminal XhoI site

8. 1 M Isopropyl-β-D-thiogalactoside (IPTG). 9. 10% Sodium dodecyl sulfate (SDS). 10. 0.5% Ammonium persulfate (APS). 11. 12% SDS-PAGE gels. 12. 5 mg/mL Lysozyme solution. 13. Benzonase nuclease. 14. 5% Sodium deoxycholate. 15. 2 M Imidazole. 16. 3 M Potassium chloride. 17. 5 M Sodium chloride. 18. NP-40 (Nonidet P-40 Substitute). 19. 50% Ammonium sulfate. 20. Lysis buffer: 50 mM Tris–HCl (pH 8.0), 20% sucrose. 21. Nickel column binding buffer: 30 mM Imidazole, 15.4 mM Na2HPO4, 4.5 mM NaH2PO4, 150 or 600 mM NaCl (pH 7.4). 22. Nickel column elution buffer: 250 mM Imidazole, 15.4 mM Na2HPO4, 4.5 mM NaH2PO4, 150 or 600 mM NaCl (pH 7.4).

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23. Histidine-tagged protein dialysis buffer: 20 mM Tris–HCl, pH 7.5; 1 mM EDTA; 500 mM NaCl. 24. Histidine-tagged protein storage buffer: 20 mM Tris–HCl, pH 7.5; 1 mM EDTA; 500 mM NaCl; 50% glycerol. 25. Avidin column binding buffer: 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4. 26. Avidin column elution buffer: 2 mM Biotin in PBS. 27. Profinity bind/wash buffer: 0.1 M Sodium phosphate, pH 7.2. 28. Profinity elution buffer: 0.1 M Sodium phosphate, 0.1 M sodium fluoride, pH 7.2. 29. Profinity storage buffer: 0.1 M Sodium phosphate, 0.02% sodium azide, pH 7.2. 30. Gel filtration buffer: 20 mM Tris–HCl, pH 8.0; 1 mM EDTA; and 100 mM NaCl. 31. Avidin and Profinity eXact columns: Prepare the avidin column and Profinity columns following the manuals of the suppliers. 32. Image-Pro Plus v6.1 (Media Cybernetics). 33. Fiji [15].

3

Methods

3.1 Dual/TriplePlasmid Transformation

1. Add 1–1.5 μL of each plasmid (Table 1) into a 1.5 mL microcentrifuge tube and add 50 μL of Tuner (λDE3)-competent cells. Mix gently by tapping the bottom of the tube and incubate on ice for 30 min. 2. Heat shock at 42
C for 2 min, and place on ice for 2 min. 3. Add 450 μL 2XYT and grow at 37
C for 60–75 min. 4. Plate 100 μL of cells on LB plate containing corresponding antibiotic(s) (50 μg/mL of carbenicillin, 250 μg/mL of ampicillin, 25 μg/mL of kanamycin, and/or 25 μg/mL of chloramphenicol) and 0.2% glucose. 5. Incubate the plate at 37
C overnight.

3.2 Dual/ Triple-Plasmid Coexpression and Cell Growth

1. For biotinylated protein, add a final concentration of 0.5 mM D-biotin to the LB media before autoclaving (see Note 2). 2. Select eight well-separated colonies from the transformation plate, and grow overnight at 37
C in 5 mL of LB containing antibiotics and 0.2% glucose to maintain transformed plasmids and minimize protein expression. 3. To verify protein expression level, use each of the overnights to separately inoculate eight 5 mL cultures. These contain fresh LB medium containing antibiotic(s) and are inoculated at 1:100 and grown for 2 h at 37
C (OD600 ¼ 0.4–0.5).

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4. Add IPTG to 100 or 300 μM (final), followed by an additional 3 h of growth. 5. Harvest 1 mL of cells by centrifugation at 5000 rcf for 1 min, and resuspend in 180 μL of autoclaved water followed by addition of 20 μL of 10% SDS. 6. Lyse the resuspended cells by boiling for 5 min at 100
C. 7. Take 25 μL of the lysate, mix with SDS loading dye, and subject the samples to electrophoresis in 12% SDS-PAGE gels. 8. Stain the gels with Coomassie Brilliant Blue. 9. Select the best expression clone which has comparable levels of either two or three proteins. Make a fresh overnight culture with antibiotics and 0.2% glucose. 10. The next day, inoculate 0.1, 1, or 10 L cultures containing antibiotic(s) using the fresh overnight at a ratio of 1:100. 11. Grow cells in flasks for 100 mL or 1 L cultures with vigorous shaking at 280 rpm under 37
C or in a 16 L fermentor for 5 or 10 L cultures. 12. Add IPTG at 100 or 300 μM (final) when the culture OD600 is between 0.4 and 0.5, and grow until the cell culture reaches the stationary phase (typically 3 h). 13. Harvest cells by centrifugation at 5000 rcf at 4
C, and resuspend cell pellets in ice-cold histidine-tagged protein lysis buffer at 2 mL/g by vortexing. Add additional lysis buffer to a total of 10 mL if the resuspension mixture is less than 10 mL. 14. The cell resuspension mixture can be stored at 20
C for short term and 80
C for long term, or stirred at 4
C overnight at 30–50 rpm for lysis the following day. 3.3 Protein Purification

3.3.1 Purification of Histidine-Tagged Protein Using Nickel Column Chromatography

All of the cell lysis, column preparation, protein loading, washing, and elution steps should be conducted at 4
C to diminish protein denaturation and achieve optimal purification. 1. Add lysozyme (1 mg/mL final) and benzonase (3 μL for 1 L and 10 μL for 5 L of culture, separately) to the lysis mixture and stir for 30 min. 2. Add deoxycholate (0.05% final) and stir for an additional 30 min. 3. Add imidazole to a final concentration of 30 mM and KCl or NaCl to a final concentration of 600 (high salt) or 150 (low salt) mM, respectively (see Note 3). 4. Centrifuge the whole-cell lysate at 37,000 rcf for 1 h. 5. Load the cleared cell lysate onto a nickel column using a flow rate of 2 mL/min.

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6. Apply three sequential washes after loading to remove both unbound and nonspecifically bound proteins: 50 column volume (CV) of binding buffer, 40 CV of binding buffer + 0.2% NP40, and 30 CV of binding buffer. 7. Mix nickel column binding and elution buffers to achieve a linear, imidazole gradient from 30–500 mM to elute proteins at a flow rate of 1 mL/min, and identify proteins using 12% SDS-PAGE. 8. After electrophoresis, stain the gels with Coomassie Brilliant Blue, destain, and photograph. 9. The presence of eluted complexes can be easily visualized in the destained gels (Fig. 1a, b, d, and e). 3.3.2 Purification of Biotinylated Protein After Nickel Column Chromatography

1. Prepare the avidin column (containing 5 mL resin) following the manufacturer’s instructions. 2. Collect fractions from the nickel column, combine, and dialyze against the avidin column binding buffer. 3. Load the proteins onto the avidin column using a flow rate of 1 mL/min. 4. Turn off the flow and incubate the column for 1 h to enhance protein binding. 5. Wash with 10 CV avidin column binding buffer (2 mL/min) until UV trace reaches the baseline before elution. 6. Elute proteins with 50 mL avidin column elution buffer at 1 mL/min. 7. Identify fractions containing complexes using 12% SDS-PAGE (Fig. 1f). 8. Pool fractions and precipitate proteins using 50% ammonium sulfate solution. 9. Recover purified proteins by centrifuging at 38,000 rcf for 90 min, and then resuspend in 1/10th volume Profinity bind/wash buffer. 10. Dialyze resuspended proteins against Profinity bind/wash buffer and store dialyzed protein in Profinity storage buffer.

3.3.3 Purification of Profinity-Tagged Complexes

1. Equilibrate the Profinity column with Profinity bind/wash buffer (10 CV) at 3 mL/min. 2. Apply dialyzed proteins to the 5 mL Profinity column at 2 mL/ min. 3. Wash with the Profinity bind/wash buffer to baseline at 3 mL/ min. 4. Cleave the complex by application of Profinity elution buffer containing 0.1 M sodium fluoride (see Note 5) at 2 mL/min.

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5. Identify complexes in eluted fractions by 12% SDS-PAGE (Fig. 1c). 6. If protein levels are low, it may be necessary to stain gels with silver. 3.3.4 Gel Filtration

1. Equilibrate the Superose 6, 10/300 GL, column in gel filtration (GL) buffer. 2. To determine the size of protein complexes, apply 250 μL of individual molecular weight markers to the Superose 6, 10/300 GL, to generate the calibration curve with a plot of Kav vs. log of molecular weight markers (see Note 6). 3. Elute proteins by the application of gel filtration buffer at 1 mL/min, and calculate elution volume by measuring the volume of the eluent from the point injection minus the center of the elution peak. 4. Determine the molecular weight of the protein sample depending on the Kav of the calibration curve.

3.4 Fluorescence Microscopy

1. Use a fresh overnight culture to inoculate 5 mL LB containing antibiotics and grow at 37
C for 3 h. Cultures are grown without IPTG to maintain the basal protein expression levels for imaging. 2. Harvest cells at 5000 rcf at room temperature and resuspend in 10 mM MgSO4. 3. Add 10 μL of resuspended cells to poly-L-lysine-coated glass coverslips and allow binding at room temperature for 10 min. 4. Wash coverslips twice with 10 mM MgSO4 to remove unbound cells. 5. Dry coverslips at room temperature for 10 min. 6. Place 15 μL of 10 mM MgSO4 onto a microscope slide, mount the coverslip on a microscope slide, and seal with nail polish. 7. Image using epifluorescence microscopy and appropriate filters for GFP and mcherry fluorescence (Fig. 2). 8. Analyze captured images using Image-Pro Plus v6.1 (Media Cybernetics) or Fiji (Schneider et al. 2012). 9. If background fluorescence is too high, fusion genes can be subcloned into low copy number expression vectors [8]. 10. If background fluorescence is still too high, grow cells in the presence of 0.2% glucose to lower the expression even further. 11. Alternatively, fusion genes can be introduced into the chromosome at ectopic loci using recombineering [14].

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Notes 1. Tuner (λDE3) used in the study contains a deletion of lacZY, which enables adjustable levels of protein expression in all cells by varying the concentration of the inducer, isopropyl β-D-1thiogalactopyranoside (IPTG). 2. The high temperature during autoclaving helps dissolve biotin in LB media.

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3. Salts are added at the end of lysis but before centrifugation to match the working salt concentration of the nickel column binding buffer. 4. Two types of nickel columns with similar chromatography conditions can be used depending on the total volume of the cleared cell lysate. When the cleared cell lysate is less than 30 mL (1 L cell culture), a 5 mL HisTrap FF column should be used. When the cleared cell lysate is equal or more than 30 mL (5 or 10 L cell culture), a 25 mL nickel column should be used (HisTrap FF manual, CytivaLifeSciences). 5. For the Profinity column, use sodium acetate (NaOAc) instead of NaCl in cell lysis buffer as chloride ions cleave the Profinity tag. 5. For Superose 6, 10/300 GL column, the void volume of the column is 7.2 mL, approximately 30% of the total column volume (24 mL). 7. Molecular weight markers of calibration curve include ferritin (440,000 Da), catalase (232,000 Da), aldolase (158,000 Da), alcohol dehydrogenase (150,000 Da), conalbumin (75,000 Da), ovalbumin (44,000 Da), carbonic anhydrase (29,000 Da), and ribonuclease A (13,700 Da). References 1. Lecointe F, Serena C, Velten M, Costes A, McGovern S, Meile JC, Errington J, Ehrlich SD, Noirot P, Polard P (2007) Anticipating chromosomal replication fork arrest: SSB targets repair DNA helicases to active forks. EMBO J 26(19):4239–4251 2. Shereda RD, Kozlov AG, Lohman TM, Cox MM, Keck JL (2008) SSB as an organizer/ mobilizer of genome maintenance complexes. Crit Rev Biochem Mol Biol 43(5):289–318. https://doi.org/10.1080/ 10409230802341296 3. Sun Z, Hashemi M, Warren G, Bianco PR, Lyubchenko YL (2018) Dynamics of the interaction of RecG protein with stalled replication forks. Biochemistry 57(13):1967–1976.

https://doi.org/10.1021/acs.biochem. 7b01235 4. Bianco PR, Lyubchenko YL (2017) SSB and the RecG DNA helicase: an intimate association to rescue a stalled replication fork. Protein Sci 26(4):638–649. https://doi.org/10. 1002/pro.3114 5. Bianco PR (2017) The tale of SSB. Prog Biophys Mol Biol 127:111–118. https://doi.org/ 10.1016/j.pbiomolbio.2016.11.001 6. Manosas M, Perumal SK, Bianco PR, Ritort F, Benkovic SJ, Croquette V (2013) RecG and UvsW catalyse robust DNA rewinding critical for stalled DNA replication fork rescue. Nat Commun 4:2368. https://doi.org/10.1038/ ncomms3368

SSB Binding to Fork Rescue Helicases 7. Buss J, Kimura Y, Bianco P (2008) RecG interacts directly with SSB: implications for stalled replication fork regression. Nucleic Acids Res 36(22):7029–7042 8. Yu C, Tan HY, Choi M, Stanenas AJ, Byrd AK, K DR, Cohan CS, Bianco PR (2016) SSB binds to the RecG and PriA helicases in vivo in the absence of DNA. Genes Cells 21(2):163–184. https://doi.org/10.1111/gtc.12334 9. Slocum SL, Buss JA, Kimura Y, Bianco PR (2007) Characterization of the ATPase activity of the Escherichia coli RecG protein reveals that the preferred cofactor is negatively supercoiled DNA. J Mol Biol 367(3):647–664. https://doi.org/10.1016/j.jmb.2007.01.007 10. Handa N, Bianco PR, Baskin RJ, Kowalczykowski SC (2005) Direct visualization of RecBCD movement reveals cotranslocation of the RecD motor after chi recognition. Mol Cell 17(5):745–750. https://doi.org/10.1016/j. molcel.2005.02.011 11. Biosciences G (2002) Gel filtration. Principles and methods. GE Healthcare Bio-Sciences AB, Uppsala 12. Bianco PR, Stanenas AJ, Liu J, Cohan CS (2012) Fluorescent single-stranded

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DNA-binding proteins enable in vitro and in vivo studies. Methods Mol Biol 922:235–244. https://doi.org/10.1007/ 978-1-62703-032-8_18. 13. Liu J, Choi M, Stanenas AG, Byrd AK, Raney KD, Cohan C, Bianco PR (2011) Novel, fluorescent, SSB protein chimeras with broad utility. Protein Sci 20(6):1005–1020. https://doi. org/10.1002/pro.633 14. Zhao T, Liu Y, Wang Z, He R, Xiang Zhang J, Xu F, Lei M, Deci MB, Nguyen J, Bianco PR (2019) Super-resolution imaging reveals changes in Escherichia coli SSB localization in response to DNA damage. Genes Cells 24 (12):814–826. https://doi.org/10.1111/gtc. 12729 15. Schindelin J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch T, Preibisch S, Rueden C, Saalfeld S, Schmid B, Tinevez JY, White DJ, Hartenstein V, Eliceiri K, Tomancak P, Cardona A (2012) Fiji: an opensource platform for biological-image analysis. Nat Methods 9(7):676–682. https://doi.org/ 10.1038/nmeth.2019

Chapter 6 Magnetic Tweezers-Based Single-Molecule Assays to Study Interaction of E. coli SSB with DNA and RecQ Helicase Debjani Bagchi, Weiting Zhang, Samar Hodeib, Bertrand Ducos, Vincent Croquette, and Maria Manosas Abstract The ability of magnetic tweezers to apply forces and measure molecular displacements has resulted in its extensive use to study the activity of enzymes involved in various aspects of nucleic acid metabolism. These studies have led to the discovery of key aspects of protein-protein and protein-nucleic acid interaction, uncovering dynamic heterogeneities that are lost to ensemble averaging in bulk experiments. The versatility of magnetic tweezers lies in the possibility and ease of tracking multiple parallel single-molecule events to yield statistically relevant single-molecule data. Moreover, they allow tracking both fast millisecond dynamics and slow processes (spanning several hours). In this chapter, we present the protocols used to study the interaction between E. coli SSB, single-stranded DNA (ssDNA), and E. coli RecQ helicase using magnetic tweezers. In particular, we propose constant force and force modulation assays to investigate SSB binding to DNA, as well as to characterize various facets of RecQ helicase activity stimulation by SSB. Key words SSB, RecQ helicase, Magnetic tweezers, Force, Single molecule, DNA

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Introduction Studying life processes at high resolution both spatially and temporally has become possible nowadays thanks to the development of super-resolution fluorescence imaging and force spectroscopy techniques (such as atomic force microscopy, optical and magnetic tweezers), which used separately or in combination yield to an unprecedented view of cellular processes [1–8]. While fluorescence methods are limited to the study of fast dynamics only, because of photobleaching of fluorescent probes, force spectroscopy methods allow studying slow dynamics (spanning hours), as well as millisecond-scale fast dynamics [9, 10]. Moreover, the latter allow the application and measurement of pico-Newton (pN) forces, the typical range of forces found in biomolecular processes involving

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_6, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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few kBT of energy (1 kBT ¼ 4.28 pN.nm at T ¼ 310 K). Force acts as an additional thermodynamic variable enabling mechanobiology assays [10]. In a magnetic tweezer assay, the biomolecule of interest is tethered between a micron-sized superparamagnetic bead and the glass surface of a microchannel with the help of antigen-antibody interactions. The magnetic field is applied by two small NdFeB permanent magnets having vertical magnetization with opposite direction and with a 1 mm gap in between. The bead magnetization follows the magnetic field and its dipole moment gets oriented horizontally while the field gradient is vertical. Force modulation from ~102 pN to ~20–30 pN is achieved by precise and controlled motion of a motor which can change the height of the magnets with respect to the microchannel containing the beads, thereby resulting in a near-perfect force clamp [9, 11]. Moreover, magnetic tweezers can also generate torsion with ease, simply by rotation of the magnets. Using magnetic tweezers, real-time monitoring of DNA extension can detect small changes in DNA conformation (in the nm scale), such as the opening of a few base pairs or the binding of ligands that induces a change in DNA elasticity. Several magnetic tweezers-based studies on protein-DNA interactions have revealed intermediates in crucial kinetic pathways and heterogeneities in dynamics of proteins involved in DNA metabolism (DNA replication, repair, and recombination) as well as in RNA metabolism [12–20]. This chapter revolves around the different ways in which magnetic tweezers can be employed to study the interaction between E. coli single-stranded binding protein (SSB), DNA, and E. coli RecQ helicase with high statistical accuracy as a result of real-time multi-bead tracking. SSB plays a key role in genomic maintenance [21]. During DNA replication, repair, and recombination, doublestranded DNA (dsDNA) has to be converted to the single-stranded form (ssDNA). The ssDNA, once produced, is highly susceptible to damage and degradation by chemicals, radiation, or nucleolytic enzymes. SSBs cooperatively bind to ssDNA and stabilize it preventing its degradation. Moreover, SSB-DNA complexes act as substrates for the loading of crucial enzymes such as polymerases, exonucleases, topoisomerases, and helicases, enhancing their activity [21]. SSB exists as a homo-tetramer, due to interactions between the N-terminal domains of each SSB monomer. The C-terminal domain of SSB mediates interactions with the proteins involved in genomic maintenance [21]. Incorporating a fluorescent tag or fluorescent protein at the SSB C-terminal or N-terminal domains has been found to impair tetramer formation or its interaction with other proteins, limiting fluorescence-based assays [22, 23], and stressing the need for other single-molecule assays which do not require protein modifications.

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This chapter focuses on constant force and force jump DNA hairpin single-molecule assays using magnetic tweezers to investigate SSB-ssDNA binding modes, as well as the interactions between SSB and E. coli RecQ helicase. RecQ helicase in association with SSB plays a key role in cellular genome maintenance, such as regulation of recombination and replication fork maintenance [24]. The presence of SSB has been found to stimulate RecQ unwinding activity [16, 24–27]. Affinity purification studies had established that the winged helix domain of RecQ interacts with the C-terminal tail of SSB [25]. However, our recent single-molecule studies, using different RecQ and SSB variants, have shown that the interaction of RecQ with SSB extends beyond the winged helix domain [16]. Moreover, these studies show that SSB enhances the overall RecQ helicase activity, increasing its binding affinity, unwinding rate, and processivity [16]. The assays developed involve constant force and force modulation experiments with a DNA hairpin substrate. The SSB binding and helicase activity are detected through changes in DNA extension measured by tracking a micron-sized magnetic bead attached to the DNA hairpin. The detailed protocols for these assays are the main subject of this chapter.

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Materials 1. DNA substrate: A 1239 bp hairpin with a 4 nt loop, a 76 nt 50 -biotinylated ssDNA tail, and a 146 bp 30 -digoxigeninlabeled dsDNA tail [28]. The dsDNA tail specifically binds to an anti-digoxigenin-coated coverslip. The other extremity of the hairpin with a biotin label is bound to a micron-sized magnetic bead coated with streptavidin. 2. Purified E. coli SSB and variants with mutations in the C-terminal [29] (see Note 1). 3. Purified SSB-Ct peptide [29]. 4. Purified gp32-SSB-Ct fusion protein [29]. 5. Purified E. coli RecQ-wt and RecQ-ΔC (the RecQ variant with HRDC domain removed) [30]. 6. Magnetic trap. 7. Passivated and surface-treated microchannel: Protocols for passivation and surface treatment of the microchannel and troubleshooting tips are given in a previously published article [11, see Notes 3 and 4]. 8. Micron-sized magnetic beads.

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9. Working buffer for SSB and helicase assays: 20 mM Tris–HCl pH 7.6, 25 mM NaCl, 3 mM MgCl2, 2% BSA, 0.5 mM DTT, and 0.5 mM ATP. 10. 24-mer Oligonucleotide complementary to the loop region of the hairpin.

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Methods A schematic diagram of the magnetic tweezers setup is shown in Fig. 1. The magnetic bead is trapped in a magnetic tweezers generated by a pair of permanent magnets, which produce a force due to the vertical magnetic field gradient. These permanent magnets are fixed to a holder whose distance from the coverslip surface can be changed by the precise movement of a PI DC motor, thereby changing the force exerted on the magnetic bead. This geometry allows one to exert a mechanical force on the bead (and on the tethered DNA hairpin), ranging from 102 to 30 pN, with sub-pN accuracy (see Note 5). The force is calculated from the Brownian fluctuations of the tethered bead. A microscope objective images the bead onto a CCD/CMOS camera for real-time position 3D tracking with an acquisition frequency of 30 Hz. Thus tracking the z-fluctuations of the tethered bead enables us to monitor the change in extension of the DNA hairpin in real time, with about 2 nm accuracy. The extension of the hairpin changes abruptly at forces greater than 15 pN, due to the mechanically induced unzipping. The hairpin can also be unzipped at lower forces in the presence of a helicase and ATP. Typically around 50 beads are tracked in real time simultaneously, leading to large statistics of single-molecule events (see Note 2). The image of the bead displays diffraction rings that are used to estimate its 3D position as explained elsewhere [9]. From the fluctuating positions of the bead both the mean elongation of the molecule and the force applied to it can be deduced [9]. The z-fluctuations are acquired at 30 Hz and recorded by a CMOS camera. The binding of a ligand on DNA (such as SSB) induces a change in the DNA elasticity. On the other hand, a helicase that unwinds a DNA hairpin under tension adds two bases of stretched ssDNA for each unwound base pair (corresponding to about 1 nm increase in the measured extension at 10 pN). Therefore, both binding and helicase activity induce a change in the DNA extension that is observed as a change in the z-coordinate of the bead.

3.1 DNA Force-Extension Measurements to Study SSB Binding

The binding of ligands, such as SSB proteins, changes the elasticity of ssDNA. Measuring the extension as a function of the force of a ssDNA molecule in the presence of ligands is a way to characterize the ligand binding affinity [31–35]. By performing titration

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Fig. 1 Sketch of the magnetic tweezers setup. The biomolecule of interest, a ssDNA in the figure, is attached to a paramagnetic bead, kept in the magnetic field generated by a pair of magnets. The other end of the ssDNA molecule has a digoxigenin, which binds to the anti-digoxigenin-coated cover glass surface in a microchannel. The microfluidic chamber is equipped with the inlet and outlet, through which appropriate buffers can be flown in and ejected by a syringe pump. The force applied on the bead is tuned by changing the height of the magnets with respect to the microchannel. By rotating the magnets, torsion can also be applied on torsionally constrained molecules (like double-stranded DNA). Since ssDNA is not sensitive to torsion it is possible to distinguish a single ssDNA from a situation where multiple biomolecules are tethered between the bead and the surface [43]. The microchannel is kept on an inverted bright-field microscope and video imaging of many beads is done with a CCD or CMOS camera

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experiments, one can measure ssDNA force-extension curves at different ligand concentrations (c0 ) and compute the extension change in the molecule length ΔL (ΔL ¼ (XssDNA ([c] ¼ 0)  XssDNA([c] ¼ c0 ))) at a given force, which is related to the ligand occupancy. If the ligand has a unique mode of binding, the affinity constant and the binding cooperativity can be characterized by measuring the dependence of ΔL versus the enzyme concentration and fitting the results to a ligand-binding model, such as the Hill model [32, 36] or the McGhee Von–Hippel model [33, 34, 37]. For complex ligand binding presenting several binding modes, as it is the case of E. coli SSB [38–41], more complex models need to be applied and the elasticity measurements alone might not be enough to characterize binding. However, as shown below, these assays can throw some light on the dominating binding mode. Since E. coli SSB exists as a homo-tetramer, it has four oligonucleotide-binding domains that bind to ssDNA. Depending on the concentration of SSB with respect to ssDNA and salt concentration in the buffer, SSB can bind ssDNA with multiple conformations that wrap the ssDNA substrates to different degrees, involving different number of nucleotides (from 17 to 65) [38– 41]. The two main binding modes are the (SSB)35 and (SSB)65 modes [39, 40]. In the (SSB)35 mode, SSB homo-tetramer wraps around itself 35 nucleotides, whereas in the latter, 65 nucleotides are wrapped around the SSB homo-tetramer. Of the two modes, the (SSB)35 mode is highly cooperative, and is predominant in low NaCl or MgCl concentration (below 200 mM NaCl or below 2 mM MgCl), and at high SSB-to-ssDNA ratio [38, 39]. The (SSB)56 mode is observed at intermediate-salt conditions [38]. The force applied on the ssDNA molecule might also affect the SSB binding preferences since the (SSB)65 and (SSB)56 modes compact ssDNA more than the (SSB)35 and (SSB)17 modes. Indeed, recent studies with optical tweezers proved that the equilibrium between different SSB binding modes is affected by the presence of the stretching force [41]. Moreover, single-molecule studies combining optical tweezers and fluorescence have shown that force can unwrap the ssDNA around the SSB and induce the SSB unbinding [42]. One difficulty in ssDNA micromanipulation is to avoid that a part of the molecule adsorbs nonspecifically to surfaces. When manipulating a ssDNA molecule with magnetic tweezers, part of the molecule might be adsorbed on the surface of the magnetic bead and/or the microfluidic chamber, which makes it difficult to control the number of nucleotides that are stretched by the tweezers. In order to avoid this problem, we use a hairpin and the oligonucleotide-blocking method [13] to measure the ssDNA elasticity in the presence or absence of SSB. Figure 2a shows a scheme of the protocol used below.

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Fig. 2 (a) Sketch of the protocol to study the changes in ssDNA elasticity upon SSB binding. A DNA hairpin is first unzipped mechanically, in the presence of an oligonucleotide, which has a sequence complementary to the loop of the DNA hairpin. Hybridization of this oligo blocks hairpin reformation when the force is decreased back to 6 pN. Since the oligonucleotide is only 24 nt, effectively a large ssDNA substrate is produced from the 1239 bp hairpin. SSB is then introduced in the microchannel in the working buffer, and force ramp cycle is initiated after few seconds from nearly zero force. (b) Average force-extension curves of the ssDNA molecule in the working buffer (3 mM Mg2+), in the presence of SSB at the indicated concentrations, obtained using the protocol described in panel A. The protocol is done by gradually increasing and decreasing the force. At SSB concentrations less than or equal to 260 pM, the force increasing curve and the force decreasing curve do not superimpose. The observed hysteresis is due to SSB binding/unbinding, dynamic rearrangements of ssDNA loops, and diffusion of bound SSBs that occur in timescales that are larger than the typical rate at which the force is changed. The inset shows a typical force-extension curve for 65 pM SSB, which presents the largest hysteresis between increasing and decreasing force trajectories, with the mean force increasing and force decreasing curves shown in purple dots. The average between the force increasing and the force decreasing curves is shown as a yellow dashed line with the corresponding error bars. Due to the hysteresis, the errors in the average extension are very large for [SSB]  260 pM. In this concentration range, we only show in the main plot the averaged curves as dashed curves. Errors are lowered for [SSB]  520 pM, because the hysteric behavior disappears. For comparison, the force-extension curve of the ssDNA in the absence of SSB is also shown. The ssDNA curve is fitted with the improved Marko-Siggia worm-like chain (WLC) model [49] (gray curve). (c) The compaction due to SSB binding as a function of SSB concentration is shown as C ¼ (XssDNAXssDNA + SSB)/XssDNA, where XssDNA and XssDNA + SSB are, respectively, the extension of ssDNA in the absence and presence of SSB. The compaction at 2 pN is calculated by extracting the ssDNA extension at 2 pN from the Marko-Siggia WLC fit to the experimental data

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1. Attach the hairpins with magnetic beads to the lower surface of the microfluidic chamber (see [10, 43] for details). 2. Introduce the 24-mer oligonucleotide complementary to the loop region of the hairpin in the microfluidic chamber (typically at ~1 μM concentration) in the working buffer. 3. Apply high force (above 15 pN) and mechanically unzip the hairpins. This hybridizes the oligonucleotide in the hairpin loop. 4. Decrease the force to a low value (3–6 pN). At this force the hairpin formation is kinetically arrested by the oligonucleotide hybridized at the loop region. Below ~3 pN the hairpin rezips in the working conditions. 5. Measure the force-extension curve of the ssDNA molecule by performing gradual force ramps from 3.5 pN to 17 pN and then back to 3.5 pN. 6. Measure next the ssDNA elasticity in the presence of SSB. For this, inject the SSB at a given concentration in the working buffer into the microfluidic chamber at an intermediate force (Fhold ¼ 6 pN); at this force the hybridized oligonucleotide still prevents hairpin reannealing (Fig. 2a). 7. After waiting some time to allow for SSB binding (typically a few seconds, as SSB binding is fast in the working conditions), set the force to a low value (~0.1 pN). Record a force cycle, where the force is increased to 17 pN and next decreased again at a given average loading rate of ~0.1 pN/s. In order to prevent hairpin reformation below ~4 pN the ssDNA needs to be significantly covered with SSB. So the full force-extension curve, along the full range of forces 0–17 pN, could only be measured above a certain SSB concentration limit of 65 pM in our working conditions. At concentration below 65 pM the hairpin rezips at low forces, showing that, in this low concentration range, the bound SSBs do not cover the ssDNA molecule. In contrast, at 65 pM and above, the hairpins do not rezip even at very low forces, revealing that ssDNA is already mostly covered with SSB. As shown in Fig. 2b, by adding different concentrations of SSB, the ssDNA elasticity changes significantly. A clear result is that, for all SSB concentrations, the SSB shortens the ssDNA molecule. This result is expected since ssDNA is known to wrap along a SSB tetramer (in loops of either 17 nt, 35 nt, 56 nt, or 65 nt) [38–41]. Moreover, at low SSB concentration (below 0.5 nM), a hysteresis is observed at high forces (above 2–3 pN) and the force-increasing curve and the force-decreasing curve do not overlap (see inset in Fig. 2b). These results have two implications: (1) the SSB binding and unbinding kinetics depend on the force applied, e.g., kon(F) and koff(F), and

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(2) at low SSB concentration (below 0.5 nM) the system is not able to equilibrate and the kinetics of binding/unbinding is slower than the rate at which the force is changed. As a result, for SSB concentrations below 0.5 nM, the force-extension data have large errors (the average curves are shown as dashed lines in Fig. 2b and errors are shown in the inset). In contrast, at high concentrations (above 0.5 nM) the hysteresis disappears meaning that the binding/ unbinding kinetics becomes faster and the system equilibrates. The relative compaction factor defined as C ¼ (XssDNAXssDNA + SSB)/XssDNA (where XssDNA and XssDNA + SSB are, respectively, the extension of ssDNA in the absence and presence of SSB) can be used to characterize the extent of compaction and gives some idea about which SSB mode is predominant. As shown in Fig. 2c, the compaction C is maximum at low SSB concentrations (65–130 pM) and minimum at high SSB concentrations. These results are consistent with significant ssDNA covering of SSB at subnanomolar concentrations with the (SSB)65 or (SSB)56 modes, which are the more compact states. As the SSB concentration increases, C decreases indicating the conversion from the (SSB)65/(SSB)56 modes to the less compacted (SSB)35/ (SSB)17 modes. At the lower concentration of 65 pM the average C at forces above 3 pN is lower than that at 130 pM (Fig. 2c). This is probably reflecting the slightly lower occupancy of SSB at 65 pM at these forces. However, at lower forces the highest compaction occurs at the lowest concentration of 65 pM and C presents a monotonic behavior (see estimated C at 2 pN in Fig. 2c). 3.2 Rezipping Assays to Study SSB Binding

The unzipping and rezipping of DNA hairpins in the presence of a constant force is a very cooperative process [13]. Hairpin unzipping and rezipping can be studied by performing force cycles: from a low force Flow to high force Fhigh and next reversing the process (from Fhigh to Flow) at a given loading rate. Starting at Flow with the hairpin formed and increasing progressively the force, the hairpin unzips in a single step when reaching a given force Fun. In the relaxing process the hairpin also rezips in a single step at Frez. Typically Fun is larger than Frez, which leads to some hysteresis. This hysteresis is due to the fact that the kinetic barrier for unzipping/rezipping is large at the midpoint force, especially for long hairpins, and the unzipping/rezipping kinetics becomes much slower than the rate at which the force is changed. Here we study how the hairpin rezipping reaction is affected by the presence of SSB. When ssDNA is covered with SSB, SSB needs to be expelled before the hairpin can rezip. Therefore SSB is expected to stall the rezipping process, reducing the kinetics of hairpin rezipping. Figure 3a, b shows the scheme of the protocol described below.

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Fig. 3 (a) Sketch of the force modulation protocol for studying SSB binding to ssDNA. A DNA hairpin is mechanically unzipped with a force Fopen ~ 22 pN (larger than the threshold value for unzipping). SSB that has been injected into the chamber binds to the fully stretched ssDNA. The force is lowered to Ftest and the ssDNA is pulled with this force for a certain interval of time ttest. The transition from Fopen to Ftest can be done in two ways: (i) performing a force jump or (ii) decreasing the force gradually with a well-defined rate. After ttest, the force is lowered to Freset ~ 0 pN, forcing the hairpin to rezip, expelling out the bound SSBs. This cycle can be repeated many times (around 100 times) to get statistically relevant single-molecule data. (b) Extension of the DNA hairpin (magenta trace) during force modulation (green trace) leading to hairpin unzipping and rezipping in the presence of SSB, [SSB] ¼ 2.6 nM

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1. Attach the hairpins with the magnetic beads to the lower surface of the microfluidic chamber (see [10, 43] for details). 2. Inject the SSB at 2.6 nM concentration in working buffer into the microfluidic chamber at a low force (below 15 pN) with the hairpin formed. 3. Unzip the hairpin by applying a constant force Fopen higher than the threshold for mechanical unzipping (Fun ~ 15 pN). Hairpin unzipping results in the formation of the ssDNA substrate, leading to SSB binding. This binding is almost instantaneous, and is independent of the duration of Fopen (Fig. 3). 4. Then decrease the force to Ftest (Ftest < 15 pN), but the hairpin formation is kinetically blocked by the bound SSBs. This appears as a plateau in the measured molecular extension as the force is held constant at Ftest. The bound SSB in the loop region prevents hairpin rezipping for a certain interval of time, tB (Fig. 4a). When the SSB at the loop unbinds the hairpin gradually rezips, ejecting out the bound SSBs along the ssDNA. The rezipping process takes a time tR (Fig. 4a). 5. Decrease the force to Freset, close to 0 pN, to eject out all SSBs and completely fold the hairpin, so that the initial configuration is recovered. 6. Repeat the force cycle N times (on the order of 100) for different Ftest forces to take good statistics of the mean blocking and rezipping rates (Fig. 4b, c) as a function of the applied force, shown as kB (F) and kR (F), respectively, in Fig. 4d, e. The whole process of blockage and rezipping depends upon the magnitude of Ftest. At low forces, the hairpin rezipping is favored over SSB binding, and bound SSBs are ejected out fast. At high forces, the initial blockage is persistent and the rezipping is hindered by pauses (Fig. 4a). Tracking about 50 hairpins in parallel a large number of blocking and rezipping events can be obtained, so statistically relevant data is obtained even though the experiments are performed at the single-molecule level. The distribution of both tB and tR at a given Ftest follows an exponential, from which the mean rates of blocking and rezipping, kB and kR, can be obtained (Fig. 4b, c). The mean kB and kR decrease exponentially with Ftest (Fig. 4d, e), which illustrates that the blocking and the rezipping are activated processes with a kinetic barrier B that increases with applied force, kB(R) ¼ 1/tB(R) ¼ exp[B] ¼ exp[(B0 + Fδ)/ kBT] ¼ k0exp[Fδ/kBT], where B0 is the kinetic barrier at zero force, k0 is the mean rate at zero force, and δ is related to the position of the transition state along the reaction coordinate. This formulation assumes that the distance δ is independent of the applied force, which in general is not true. However, the distance δ can be assumed constant if the assays are performed in a narrow range of forces where the ssDNA elasticity does not change

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Fig. 4 (a) A magnified view of one force modulation cycle (of Fig. 3b) showing blocking of hairpin reformation by bound SSBs for a time interval tB, and hairpin rezipping slowed down by bound SSBs during a time interval tR. (b, c) Distribution of the blocking time (tB) and the rezipping time (tR) obtained by repeating the force modulation cycles ~100 times for about 30 DNA hairpins. The distributions are well described by an

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significantly, as is presented here (Fig. 4d, e, forces span from ~7–7.5 pN to ~9.5–10 pN). The initial blocking is associated to the unwrapping of the SSB that is bound to the loop region, so that the measured δ can be interpreted as the distance from the SSB-bound state to the transition state for unbinding. From converting the estimated δ in nm (3.4  1.3 nm) to nucleotides (using a factor of ~0.41 nm per nucleotide t at the intermediate force of 8 pN), we find that this transition state implies the unwrapping of 8  3 nucleotides. 3.3 Constant Force Assays to Explore the Different SSB Binding Modes

Since SSB binds wrapping ssDNA in loops of 17 nt, 35 nt, 56 nt, or 65 nt [38–41], the binding and unbinding of SSB in different modes can be directly detected as a change in the extension of a tethered ssDNA molecule. Therefore by studying the changes in extension of a ssDNA substrate at the constant force Ftest, the different binding modes of SSB and their dynamic fluctuations can be studied. Figure 5a shows a scheme of the protocol described below. 1. Attach the hairpins with the magnetic beads to the lower surface of the microfluidic chamber (see [10, 43] for details). 2. Inject 24-mer oligonucleotide complementary to the loop region of the hairpin into the microfluidic chamber (typically at ~1 μM concentration). 3. Unzip the hairpin mechanically by applying a high force (above ~15 pN) and the oligonucleotide is hybridized. 4. Decrease the force to a value Ftest above 4 pN to avoid hairpin reannealing. 5. Inject 2.6 nM SSB in working buffer to microfluidic chamber to study SSB binding. Record the molecule extension. Comparison of extension traces in the presence and absence of SSB shows that in the presence of SSB, the molecular extension fluctuates much more (Fig. 5b), revealing the presence of dynamics associated to SSB binding (unbinding) and/or interchange between different SSB binding modes. The binding (unbinding) of an SSB is associated with the wrapping (unwrapping) of N nucleotides (17 nt, 35 nt, 56 nt, or 65 nt for the different binding modes). For a ssDNA molecule stretched by an ~8 pN force, this ssDNA looping (unlooping) would lead to a change in extension of

ä Fig. 4 (continued) exponential function and the results from the exponential fit are shown in the inset. (D, E) The force dependence of the blocking and rezipping rates, kB and kR, is fitted to an exponential function, with the fitted parameters shown in the inset. Taking the value of kBT ¼ 4.1 pNnm, the distance from the SSB-bound state to the transition state for unbinding, δ, is estimated to be ~3.4 nm (d) and ~5.9 nm (e)

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ΔX17 ~4 nm for the (SSB)17, ΔX35 ~10 nm for the (SSB)35 mode, ΔX56 ~18 nm for the (SSB)56 mode, and ΔX65 ~25 nm for the (SSB)65 mode (considering ~0.41 nm per nucleotide and an effective diameter of the SSB/ssDNA complex for each binding mode as described in [41]). Interestingly, the distribution of the measured ssDNA extension in the presence of SSB reveals peaks at distances compatible with the changes expected for the SSB binding in the 35 and 56 modes, ΔX35 and ΔX56 (Fig. 5c). The ssDNA molecule used in these assays is 2478 nt long, so it has many potential binding sites (about 38, 45, 70, and 145 for (SSB)65, (SSB)56, (SSB)35, and (SSB)17 modes, respectively), so many SSB binding/ unbinding reaction could occur in parallel. This would lead to hopping traces showing multiple levels consistent with multiple sites being occupied/unoccupied simultaneously. However, the extension traces typically show fast hopping between two welldefined extension values, which are separated by a fixed amount of nts until the extension is stabilized in one extension level (Fig. 5b). So, our interpretation is that we are observing the transient wrapping/unwrapping of a single SSB until it totally binds or unbinds. Interestingly, recent assays have shown the presence of an unknown SSB binding mode, where only 8 nt is bound to one of the SSB monomers without significant ssDNA wrapping [44]. Therefore, the fast hopping traces observed might be between the 8 nt unwrapped mode and one of the wrapped SSB modes. The kinetics of wrapping/unwrapping can be measured from the hopping traces from the residence time in each extension level: the residence time spent in the upper (lower) level, corresponding to the unwrapped (wrapped) state, gives the time for wrapping (unwrapping) twr (tun). Both residence times follow an exponential distribution from which the mean wrapping and unwrapping time can be measured (Fig. 5d, g). These experiments can be done at different forces to study how ssDNA wrapping and unwrapping kinetics is affected by force, as has been done recently using optical tweezers [41]. In those experiments, Suksombat et al. show how the different SSB binding modes are populated at different forces, with the more compacted (SSB)65 and (SSB)56 observed only at low forces and the less compacted (SSB)35 and (SSB)17 modes observed at intermediate forces (~8 pN) [41]. The fact that we have observed the presence of (SSB)56 at 8 pN might be a consequence that we are working at high-salt conditions (3 mM MgCl2), as compared to [41], in which the highly compacted modes (SSB)65 and (SSB)56 are favored. Finally, by performing these constant force assays at different SSB concentrations the relative binding in each mode as a function of the protein concentration can also be investigated.

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Fig. 5 (a) Sketch of the constant force assay to study SSB-ssDNA looping dynamics. (b) Changes in ssDNA due to dynamic wrapping and unwrapping of N nucleotides by SSB are observed as a decrease/increase in the measured extension, at [SSB] ¼ 2.6 nM. Wrapping of nucleotides by SSB decreases the extension, whereas unwrapping and the consequent release of nucleotides are observed as an increase in the extension. As a comparison, the blue curve shows the typical time series of the ssDNA in the absence of any bound SSBs. (c) Histograms of the extension traces showed in panel B using the same color code. The brown histogram shows peaks separated by ~19 nm, consistent with SSB binding and wrapping of 56 nt, (SSB)56 [41], whereas the gray histogram shows peaks separated by ~10 nm consistent with SSB binding and wrapping of 35 nt, (SSB)35 [41]. (d, e) The distribution of the wrapping (unwrapping) times, twr (tun) measured for the brown trace, corresponding to the wrapping (unwrapping) kinetics for the (SSB)56 mode. Results are fitted to an exponential function. (f, g) The distribution of the wrapping (unwrapping) times, twr (tun) measured for the gray trace, corresponding to the wrapping (unwrapping) kinetics for the (SSB)35 mode. Results are fitted to an exponential function

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3.4 Interaction Between SSB and RecQ Helicase

E. coli SSB is known to interact with a large number of proteins. In particular, it interacts with helicases involved in genome maintenance, such as E. coli RecQ helicase. E. coli RecQ contains three evolutionarily conserved domains: helicase, RecQ-C (comprising Zn2+-binding and winged helix (WH) domains), and “helicase and RNase D C-terminal” (HRDC) domain [30]. Previous studies [24–26] have shown that SSB stimulates the activity of RecQ via an interaction that involves the disordered C-terminal tail of SSB (SSB-Ct) and the WH domain from RecQ. However, it was not clear whether other interaction modes are present between the two proteins. In order to investigate the SSB-RecQ interaction we have used different RecQ and SSB variants.

3.4.1 Analyzing RecQ Variants

First, by manipulating a DNA hairpin using magnetic tweezers [9, 45–47] we study the DNA unwinding activity of wild-type RecQ (RecQ-wt) and two deletion variants: RecQ-ΔC where the HRDC domain is removed and RecQ-ΔΔC where both HRDC and WH domains are removed. Figure 6a shows a scheme of the protocol described below. 1. Attach the hairpins with the magnetic beads to the lower surface of the microfluidic chamber (see [10, 43] for details). 2. Keep the force constant to a value Ftest below the unfolding force (~15 pN), where the hairpin is mechanically stable and remains formed. Inject the RecQ protein (RecQ-wt or one of the RecQ variants) at a given concentration with the working buffer and 1 mM ATP. Above 4 pN, unwinding events catalyzed by the RecQ helicase are detected as an increase in the DNA molecular extension, since for each base pair (bp) unwound two nucleotides of ssDNA are released (typically ~1 nm increase for 1 bp unwound at 10 pN) (Fig. 6b). The lack of unwinding bursts below 4 pN is presumably due to the fact that the ssDNA loops around the enzyme. 3. For each RecQ variant, tune the RecQ concentration to achieve single-molecule conditions where DNA unwinding bursts are rare, with the time lag between events being approximately ten times larger than the typical duration of a single event. This concentration is typically 100–800 pM for RecQ-wt and RecQΔC and 1–10 nM for RecQ-ΔΔC. 4. Record the extension over time over long periods of time (1 h or more) to provide good data for the statistics of unwinding events. From the extension traces, conversion of extension to unwound bps can be done using the ssDNA elasticity ([16] and Fig. 2b). From these assays the unwinding rate and processivity can be directly computed at different forces (typically from 4 to 13 pN)

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(s) Fig. 6 (a) Sketch of the constant force DNA hairpin assay to study the RecQ helicase activity in the presence and absence of SSB. Unzipping of the DNA hairpin by a helicase at a constant force in the presence of ATP leads to two ssDNA nucleotides released for each base pair unwound (~1 nm extension change at 10 pN). The tracking of the magnetic bead allows observing changes in extension as a function of time, which allows following the helicase activity in real time. (b) A typical unwinding event, obtained for the RecQ-ΔC variant at 9.5 pN. The hairpin extension, converted to the number of base pairs unzipped, as a function of time is shown by the blue curve. From the slope one can infer the helicase rate (~68 bps/s). The helicase dissociates before it reaches the hairpin apex, which is observed as an abrupt decrease in extension at around T ¼ 20.5 s. The number of bases unwound (~1050 bp) before dissociating can be used to estimate the enzyme processivity

for each RecQ variant (see Fig. 6b and [16]). These assays reveal that RecQ-wt displays two distinct kinetic modes: a “slow,” backand-forth unwinding mode, similar to the persistent random walk, and a “normal,” uniform-speed unwinding mode (Fig. 7a and [16]), whereas RecQ-ΔC exhibits only the normal unwinding

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Fig. 7 (a) Unwinding trace obtained with the RecQ-wt. It shows the two unwinding modes. The slow mode is a random back-and-forth motion, shown as magenta and green trace, with speed around 15 bps/s. The activity bursts in the slow mode can have a long duration. In the normal mode, shown in blue, the helicase unwinds

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mode (Fig. 7c and [16]). For the RecQ-ΔΔC, binding affinity for ssDNA is severely compromised [48], but an unexpected unwinding activity with an irregular speed and low processivity is preserved (Fig. 7e and [16]), indicating that the WH domain is not absolutely required for DNA unwinding. 3.4.2 Stimulation of RecQ Helicase Activity by SSB

Influence of the presence of SSB on RecQ helicase activity is studied by performing helicase assays in the presence of SSB-wt and different SSB variants: SSB variants with mutations in the C-terminal, the SSB-Ct sequence and gp32-SSB-Ct fusion protein. Figure 6a shows a scheme of the protocol described below. 1. Attach the hairpins with the magnetic beads to the lower surface of the microfluidic chamber (see [10, 43] for details). 2. Keep the force constant to a value Ftest below the unfolding force. Inject SSB-wt or SSB variants at the saturating concentrations (typically 2–5 nM), together with RecQ-wt or one of the RecQ variants at a given concentration with the working buffer and 1 mM ATP. 3. Record the extension over long periods of time (1 h or more) to obtain good statistics of RecQ unwinding events in the presence of SSB-wt or SSB variants. The influence of SSB on RecQ activity can be characterized by the single-molecule effective affinity constant defined as KA ¼ Texp[c]/Tactive, where Texp and Tactive are, respectively, the total time of the assay and the time in which the enzyme is actively unwinding the DNA and [c] is the concentration of the RecQ enzyme.

ä Fig. 7 (continued) the hairpin with a uniform speed of around 65 bps/s. Conditions used are [ATP] ¼ 0.5 mM, [RecQ-wt] ¼ 1 nM, T ¼ 29  C, and F ¼ 8 pN. (b) SSB enhances the unwinding activity of RecQ-wt measured here by the ratio r of time duration during which the enzyme is active in a burst, Ton, to the total time separating the beginning of two consecutive bursts (Ton + Twait), where Twait is the waiting time separating bursts. The probability P(r) of observing a ratio r is well fitted by an exponential function with a characteristic decay constant r0, which increased from 0.12  0.01 to 0.283  0.03 in the presence of SSB. (c) Unwinding event of RecQ-ΔC, illustrating that the helicase only works in the normal mode. Once the hairpin is fully open, RecQ-ΔC maintains the hairpin open in an active process by repeated strand switching until the enzyme dissociates. Conditions used are [ATP] ¼ 0.5 mM, [RecQ-ΔC] ¼ 400 pM, and T ¼ 29  C. (d) Experimental unwinding traces for RecQ-ΔC in the presence of SSB ([RecQ-ΔC] ¼ 400 pM, [ATP] ¼ 0.5 mM). Fast mode with unwinding speed around ~250 bp/s in addition to the normal mode is observed. RecQ-ΔC changes from normal to fast mode (with speed enhanced by a factor of 4) both while unwinding and translocating. E. Unwinding events observed with RecQ-ΔΔC. The majority of events have a very low processivity, and irregular speed. Conditions used are [RecQ-ΔΔC] ¼ 7 nM, [ATP] ¼ 0.5 mM, T ¼ 29  C, and F ¼ 8 pN. (f) Experimental unwinding trace for RecQ-ΔΔC in the presence of SSB ([RecQ-ΔΔC] ¼ 2.5 nM, [ATP] ¼ 0.5 mM). In these conditions, the enzyme fully unwinds the 1.2 kb hairpin. The RecQ-ΔΔC processivity and activity get remarkably enhanced in the presence of SSB

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The single-molecule affinity constant represents the concentration at which the enzyme is active 100% of the time (e.g., Tactive ¼ Texp) and integrates the effect of both binding affinity and enzyme processivity. In order to compare the activity of RecQ alone and in the presence of the different SSB variants, we report the ratio of the affinity constants in the absence and presence of SSB: KARecQ (wt,Δ,ΔΔ) /KARecQ(wt,Δ,ΔΔ) + SSB, with a value 1 corresponding to an enhancement effect [16]. Inclusion of SSB-wt enhances the binding and unwinding activity of the three RecQ variants assayed (RecQ-wt, RecQ-ΔC, and RecQ-ΔΔC) (Fig. 7b, d, and f). The stimulation of RecQ-ΔΔC that lacks the previously identified SSB binding site (Fig. 7f) suggests an unidentified interaction between SSB and helicase domain. Moreover, the presence of SSB leads to a new “fast” mode of DNA unwinding in RecQ-wt and different RecQ variants with rates increased by a factor of approximately four, in addition to the “slow” and “normal” modes observed in the absence of SSB (Fig. 7d and [16]). With the three enzymes, SSB enhancement is robust; it is more moderate with gp32-SSB-Ct fusion protein and disappears when the SSB-Ct sequence is altered [16]. With SSB-Ct variants, we observe an inhibition of the helicase activity, which could reflect a competition for binding ssDNA. SSB thus has two effects: first, it competes with the helicase for loading onto ssDNA, and second, it enhances the overall unwinding activity through improvements in binding affinity, unwinding rates, and processivity. The stimulation of the unwinding rate leading to the “fast” mode of DNA unwinding is not observed in the presence of the SSB-Ct alone but requires the full-length SSB [16], consistent with both RecQ/SSB and SSB/ssDNA interactions being important for stimulation of RecQ activity. Previous biochemical assays have identified a binding site for SSB in the RecQ WH domain. Our findings with RecQ-ΔΔC support a model in which a second SSB binding site apart from the WH domain, which has not been identified, plays a role in SSB-RecQ interaction.

4

Notes 1. Experiments with RecQ helicase should use protein concentrations in the single-molecule conditions, which can be deduced from binding events and separation between two unwinding bursts. 2. Before injecting the helicase or SSB with working buffer, good hairpins are selected by repeated unzipping and rezipping, although magnetic bead concentration is optimized before so that only a single hairpin is bound to a bead.

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3. Care should be taken so that sterile conditions are maintained and no nucleases are present in the microchannel due to contamination. 4. Surface passivation of microchannel by passivation buffer is needed during hairpin binding and testing; the passivation buffer has bovine serum albumin and Pluronic F127 surfactant in a specific concentration [11]. This prevents nonspecific adhesion of beads to the surface of microchannel. 5. Prior to starting experiments with new magnets, calibration of force is necessary with dsDNA using a well-documented protocol [9]. References 1. Huang B, Babcock H, Zhuang X (2010) Breaking the diffraction barrier: superresolution imaging of cells. Cell 143 (7):1047–1058. https://doi.org/10.1016/j. cell.2010.12.002 2. Biteen JS, Moerner WE (2010) Singlemolecule and superresolution imaging in live bacteria cells. Cold Spring Harb Perspect Biol 2 (3):a000448–a000448. https://doi.org/10. 1101/cshperspect.a000448 3. Hwang LC, Hohlbein J, Holden SJ, Kapanidis AN (2009) Single-molecule FRET: methods and biological applications. In: Handbook of single-molecule biophysics. Springer, New York, pp 129–163 4. Lang MJ, Fordyce PM, Engh AM, Neuman KC, Block SM (2004) Simultaneous, coincident optical trapping and single-molecule fluorescence. Nat Methods 1(2):133–139. https:// doi.org/10.1038/nmeth714 5. Neuman KC, Nagy A (2008) Single-molecule force spectroscopy: optical tweezers, magnetic tweezers and atomic force microscopy. Nat Methods 5(6):491–505. https://doi.org/10. 1038/nmeth.1218 6. van Mameren J, Peterman EJ, Wuite GJ (2008) See me, feel me: methods to concurrently visualize and manipulate single DNA molecules and associated proteins. Nucleic Acids Res 36 (13):4381–4389. https://doi.org/10.1093/ nar/gkn412 7. van Oijen AM (2011) Single-molecule approaches to characterizing kinetics of biomolecular interactions. Curr Opin Biotechnol 22 (1):75–80. https://doi.org/10.1016/j. copbio.2010.10.002 8. Greenleaf WJ, Woodside MT, Block SM (2007) High-resolution, single-molecule measurements of biomolecular motion. Annu Rev Biophys Biomol Struct 36:171–190. https://

doi.org/10.1146/annurev.biophys.36. 101106.101451 9. Gosse C, Croquette V (2002) Magnetic tweezers: micromanipulation and force measurement at the molecular level. Biophys J 82 (6):3314–3329. https://doi.org/10.1016/ s0006-3495(02)75672-5 10. Manosas M, Meglio A, Spiering MM, Ding F, Benkovic SJ, Barre FX, Saleh OA, Allemand JF, Bensimon D, Croquette V (2010) Magnetic tweezers for the study of DNA tracking motors. Methods Enzymol 475:297–320. https://doi.org/10.1016/s0076-6879(10) 75013-8 11. Lionnet T, Allemand JF, Revyakin A, Strick TR, Saleh OA, Bensimon D, Croquette V (2012) Magnetic trap construction. Cold Spring Harb Protoc 2012(1):133–138. https://doi.org/10.1101/pdb.prot067496 12. Manosas M, Spiering MM, Ding F, et al (2012) Mechanism of strand displacement synthesis by DNA replicative polymerases. Nucleic Acids Res 40:6174–6186. https://doi.org/10. 1093/nar/gks253 13. Hodeib S, Raj S, Manosas M, et al (2017) A mechanistic study of helicases with magnetic traps: Helicases Studied with Magnetic Tweezers. Protein Science 26:1314–1336. https:// doi.org/10.1002/pro.3187 14. Bagchi D, Manosas M, Zhang W, Manthei KA, Hodeib S, Ducos B, Keck JL, Croquette V (2018) Single molecule kinetics uncover roles for E. coli RecQ DNA helicase domains and interaction with SSB. Nucleic Acids Res 46 (16):8500–8515. https://doi.org/10.1093/ nar/gky647 15. Dessinges MN, Lionnet T, Xi XG, Bensimon D, Croquette V (2004) Single-molecule assay reveals strand switching and enhanced processivity of UvrD. Proc Natl Acad Sci U S A 101

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(17):6439–6444. https://doi.org/10.1073/ pnas.0306713101 16. Fiorini F, Bagchi D, Le Hir H, Croquette V (2015) Human Upf1 is a highly processive RNA helicase and translocase with RNP remodelling activities. Nat Commun 6:7581. https://doi.org/10.1038/ncomms8581 17. Manosas M, Perumal SK, Croquette V, Benkovic SJ (2012) Direct observation of stalled fork restart via fork regression in the T4 replication system. Science 338(6111):1217–1220. https://doi.org/10.1126/science.1225437 18. Manosas M, Spiering MM, Ding F, Bensimon D, Allemand JF, Benkovic SJ, Croquette V (2012) Mechanism of strand displacement synthesis by DNA replicative polymerases. Nucleic Acids Res 40 (13):6174–6186. https://doi.org/10.1093/ nar/gks253 19. Raj S, Bagchi D, Orero JV, Banroques J, Tanner NK, Croquette V (2019) Mechanistic characterization of the DEAD-box RNA helicase Ded1 from yeast as revealed by a novel technique using single-molecule magnetic tweezers. Nucleic Acids Res 47(7):3699–3710. https://doi.org/10.1093/nar/gkz057 20. Tanner NA, van Oijen AM (2010) Visualizing DNA replication at the single-molecule level. Methods Enzymol 475:259–278. https://doi. org/10.1016/s0076-6879(10)75011-4 21. Shereda RD, Kozlov AG, Lohman TM, Cox MM, Keck JL (2008) SSB as an organizer/ mobilizer of genome maintenance complexes. Crit Rev Biochem Mol Biol 43(5):289–318. https://doi.org/10.1080/ 10409230802341296 22. Curth U, Genschel J, Urbanke C, Greipel J (1996) In vitro and in vivo function of the C-terminus of Escherichia coli single-stranded DNA binding protein. Nucleic Acids Res 24 (14):2706–2711. https://doi.org/10.1093/ nar/24.14.2706 23. Liu J, Choi M, Stanenas AG, Byrd AK, Raney KD, Cohan C, Bianco PR (2011) Novel, fluorescent, SSB protein chimeras with broad utility. Protein Sci 20(6):1005–1020. https://doi. org/10.1002/pro.633 24. Shereda RD, Bernstein DA, Keck JL (2007) A central role for SSB in Escherichia coli RecQ DNA helicase function. J Biol Chem 282 (26):19247–19258. https://doi.org/10. 1074/jbc.M608011200 25. Killoran MP, Keck JL (2006) Sit down, relax and unwind: structural insights into RecQ helicase mechanisms. Nucleic Acids Res 34 (15):4098–4105. https://doi.org/10.1093/ nar/gkl538

26. Mills M, Harami GM, Seol Y, Gyimesi M, Martina M, Kova´cs ZJ, Kova´cs M, Neuman KC (2017) RecQ helicase triggers a binding mode change in the SSB-DNA complex to efficiently initiate DNA unwinding. Nucleic Acids Res 45(20):11878–11890. https://doi.org/ 10.1093/nar/gkx939 27. Shereda RD, Reiter NJ, Butcher SE, Keck JL (2009) Identification of the SSB binding site on E. coli RecQ reveals a conserved surface for binding SSB’s C terminus. J Mol Biol 386 (3):612–625. https://doi.org/10.1016/j. jmb.2008.12.065 28. Manosas M, Spiering MM, Zhuang Z, Benkovic SJ, Croquette V (2009) Coupling DNA unwinding activity with primer synthesis in the bacteriophage T4 primosome. Nat Chem Biol 5(12):904–912. https://doi.org/10. 1038/nchembio.236 29. George NP, Ngo KV, Chitteni-Pattu S, Norais CA, Battista JR, Cox MM, Keck JL (2012) Structure and cellular dynamics of Deinococcus radiodurans single-stranded DNA (ssDNA)-binding protein (SSB)-DNA complexes. J Biol Chem 287(26):22123–22132. https://doi.org/10.1074/jbc.M112.367573 30. Bernstein DA, Keck JL (2003) Domain mapping of Escherichia coli RecQ defines the roles of conserved N- and C-terminal regions in the RecQ family. Nucleic Acids Res 31 (11):2778–2785. https://doi.org/10.1093/ nar/gkg376 31. Bell JC, Liu B, Kowalczykowski SC (2015) Imaging and energetics of single SSB-ssDNA molecules reveal intramolecular condensation and insight into RecOR function. elife 4: e08646. https://doi.org/10.7554/eLife. 08646 32. Camunas-Soler J, Manosas M, Frutos S, TullaPuche J, Albericio F, Ritort F (2015) Singlemolecule kinetics and footprinting of DNA bis-intercalation: the paradigmatic case of Thiocoraline. Nucleic Acids Res 43 (5):2767–2779. https://doi.org/10.1093/ nar/gkv087 33. Chaurasiya KR, Paramanathan T, McCauley MJ, Williams MC (2010) Biophysical characterization of DNA binding from single molecule force measurements. Phys Life Rev 7 (3):299–341. https://doi.org/10.1016/j. plrev.2010.06.001 34. Chen J, Le S, Basu A, Chazin WJ, Yan J (2015) Mechanochemical regulations of RPA’s binding to ssDNA. Sci Rep 5:9296. https://doi. org/10.1038/srep09296 35. Lipfert J, Klijnhout S, Dekker NH (2010) Torsional sensing of small-molecule binding using magnetic tweezers. Nucleic Acids Res 38

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Chapter 7 High-Throughput Screening to Identify Inhibitors of SSB-Protein Interactions Andrew F. Voter Abstract The bacterial single-stranded DNA-binding protein (SSB) uses an acidic C-terminal tail to interact with over a dozen proteins, acting as a genome maintenance hub. These SSB-protein interactions are essential, as mutations to the C-terminal tail that disrupt these interactions are lethal in Escherichia coli. While the roles of individual SSB-protein interactions have been dissected with mutational studies, small-molecule inhibitors of these interactions could serve as valuable research tools and have potential as novel antimicrobial agents. This chapter describes a high-throughput screening campaign used to identify inhibitors of SSB-protein interactions. A screen targeting the PriA-SSB interface from Klebsiella pneumoniae is presented as an example, but the methods may be adapted to target nearly any SSB interaction. Key words Protein-protein interactions, Antibacterial drugs, High-throughput screening, DNA repair, DNA replication, Single-stranded DNA-binding protein

1

Introduction Bacterial single-stranded DNA-binding protein (SSB) detects and protects single-stranded (ss) DNA. Exposed ssDNA is wrapped by the essential N-terminal DNA-binding domain of SSB. Once bound, the highly conserved and acidic C-terminus of SSB (SSBct) interacts with over a dozen SSB-interacting proteins, recruiting them to sites of DNA replication and repair, facilitating their activity [1]. Bacterial cells have evolved to exploit the localization of SSB to ssDNA, enabling SSB to act as a repair hub by coordinating essential functions [1–3]. All the structurally characterized SSB-interacting proteins have a conserved docking site comprising a basic rim that surrounds a hydrophobic pocket [4–9]. The complementary SSB C-terminus (Escherichia coli and Klebsiella pneumoniae sequence: N-Met-AspPhe-Asp-Asp-Asp-Ile-Pro-Phe-C) has two critical features. The first is a series of acidic residues that contact the basic rim and the second is a pair of hydrophobic Pro-Phe residues that dock into the

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_7, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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hydrophobic pocket. In vitro, these interactions can be disrupted by removing the C-terminal-most Phe of SSB or mutating the basic lip of the interacting protein [10]. Deletion of the final Phe from SSB is lethal in E. coli, highlighting the essential nature of the SSB-protein interactions [11–13]. These vital interfaces might also act as an Achilles’ heel, as chemical disruption of this set of protein-protein interactions (PPIs) is expected to be lethal and no equivalent interaction is known to exist in mammalian systems [14]. Thus, SSB PPIs might serve as effective antibacterial agents or tool compounds to aid the study of these interactions in vivo and in vitro. An additional benefit of this compact interaction site is the ability to use the SSBct peptide in screening assays rather than the full-length protein, greatly reducing the protein requirement needed for screening. Identification of small-molecule inhibitors nearly always requires a process known as high-throughput screening (HTS). In a HTS campaign, a large initial small-molecule collection, known as a library, is winnowed through a series of screening steps that eliminate inactive or promiscuous compounds. A wide range of screening methods have been developed both in vitro and in silico and the specific assays best suited for any particular HTS campaign are dependent on the nature of the interaction and the infrastructure available to the research team. This chapter details a HTS campaign to identify inhibitors of SSB PPIs using the SSB-PriA interaction from the human pathogen K. pneumoniae as an example (Fig. 1, left) [15]. In this screening campaign, a 75,000-compound library was screened at a concentration of 33 μM using a bead-based proximity assay known as AlphaScreen (AS) (Fig. 1, AS). In an AS, the two protein partners are tethered to donor and acceptor beads. Photoactivation of the donor bead generates a short-lived singlet oxygen species that stimulates light emission only from nearby acceptor beads (that is, if the two beads are connected by the PriA-SSBct interface). If the PriA-SSBct interaction is disrupted by the addition of a small molecule, the singlet oxygen decays before reaching the acceptor bead and no signal is generated. Small molecules that block this interaction are called “hits.” Next, a counterscreen was performed to identify promiscuously acting compound. Using the same AS beads as in the primary screen, the PriA and SSBct were replaced by a single biotinylated-6
-His peptide. As the peptide has no interface that can be blocked by a small molecule, hits active in both assays are likely to act by disrupting the AS chemistry rather than the protein interface. These nonspecific compounds are not pursued farther. To further refine the set of active compounds, a fluorescence anisotropy (FA) screen was performed. In this assay, a fluoresceinlabeled SSBct (F-SSBct) is incubated with PriA and excited with polarized light. Without an inhibitor, the F-SSBct-PriA complex

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Fig. 1 Overview of the PriA-SSB PPI HTS detailed in this chapter. Left, overview of the HTS and the expected number of compounds at each stage of the screen. Right, schematic depiction of each of the screening assays

tumbles slowly in solution and the emitted light remains polarized. If the addition of a small molecule displaces the F-SSBct from PriA, the free peptide tumbles rapidly, and the resulting fluorescence emission will be depolarized. The fraction of F-SSBct bound to PriA can be determined from the ratio of polarized to unpolarized emission (Fig. 1, FA). Compounds active in both the AS and FA assays were then tested for direct interaction with PriA with a variety of biophysical techniques (Fig. 1, biophysical confirmation). These methods are discussed elsewhere in this book and will not be described in detail in this chapter. Successful completion of the screening campaign described here will yield selective and potent inhibitors of the targeted SSB-protein interaction. These screening assays are not specific to the SSB-PriA interface and have been adapted to screen for inhibitors of the SSB-DnaG and SSB-χ interactions with almost no modification. However, exact replication of the experiments detailed in this chapter would be nearly impossible due to the availability of specific screening libraries, and specialized equipment and facilities. In light of this, specific instructions are provided when possible and when this was not possible, general principles or considerations are given to guide the reader.

2

Materials

2.1 Protein Expression

1. Rosetta 2 competent cells. 2. K. pneumoniae PriA gene in a bacterial overexpression plasmid with a T7 promoter, an N-terminal 6
-His-tag, and a kanamycin resistance marker (see Note 1).

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3. Lysogeny broth (LB). 4. LB + 100 μg/mL kanamycin and 25 μg/mL chloramphenicol. 5. LB agar plates + 100 μg/mL kanamycin. 6. LB agar plates + 100 μg/mL kanamycin and 25 μg/mL chloramphenicol. 7. Spectrophotometer, with compatible 1 mL disposable cuvettes. 8. 1 M Isopropyl β-D-1-thiogalactopyranoside (IPTG) in water, filtered. 9. Centrifuge and 1 L bottles. 10. Centrifuge and 50 mL tubes. 2.2 Protein Purification

1. Lysis buffer: 10 mM HEPES-HCl [pH 7.0], 500 mM NaCl, 100 mM glucose, 10% (v/v) glycerol, 20 mM imidazole, 1 mM β-mercaptoethanol, 1 mM phenylmethylsulfonyl fluoride, one protease inhibitor tab. 2. Dilution buffer: 10 mM HEPES-HCl [pH 7.0], 10 mM DTT, 10% (v/v) glycerol. 3. Buffer A: 10 mM HEPES-HCl [pH 7.0], 150 mM NaCl, 10 mM DTT, 10% (v/v) glycerol. 4. Buffer B: 10 mM HEPES-HCl [pH 7.0], 1 M NaCl, 10 mM DTT, 10% (v/v) glycerol. 5. Centrifuges and bottles (for pelleting cells, clarifying protein lysate) and 50 mL tubes. 6. Sonicator (cell disruptor heat systems—Ultrasonics W 200 R). 7. Gravity column. 8. Ni-NTA affinity resin. 9. Fast protein liquid chromatography system with SPFF and S-300 size-exclusion columns. 10. 10% SDS-PAGE gels, electrophoresis rig, and stain.

2.3

AS Pilot and AS

1. White 1536-well plates. 2. AS nickel chelate detection kit (see Notes 2 and 3). 3. SSBct peptide (WMDFDDDIPF), lyophilized (see Note 4). 4. SSBctΔF control peptide (WMDFDDDIP), lyophilized. 5. Biotinylated SSBct peptide (Biotin- WMDFDDDIPF ), lyophilized. 6. Labcyte Echo 550 liquid handler (see Note 5). 7. Formulatrix Mantis liquid handler. 8. Small-molecule library (a 75,000-compound library from Life Chemicals was used here; see Subheading 3.4 for guidance on library selection).

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9. Small-molecule pilot library: An ~1000-compound subset of the primary library may also be used (see Note 6). 10. Software for analysis of screening data (e.g., CDD vault) (see Note 7). 11. GraphPad Prism. 12. Plate reader capable of reading 1536-well AS plates. 13. Foil plate seals. 14. Orbital shaker. 15. Darkroom (see Note 8). 16. Centrifuge compatible with 1536-well plates. 17. AS master mix: 10 mM HEPES-HCl [pH 7.5], 150 mM NaCl, 1 mM MgCl2, 10 mM DTT, 1 mg/mL bovine serum albumin, 0.01% Triton X-100, 0.1 μM PriA, 0.1 μM Bio-SSB (see Note 9). 2.4 AS Counterscreen

1. Biotinylated 6
-His (see Note 10).

2.5

1. Black 384-well plates.

FA Screening

2. AS master mix for counterscreening: 10 mM HEPES-HCl [pH 7.5], 150 mM NaCl, 1 mM MgCl2, 10 mM DTT, 1 mg/mL bovine serum albumin, 0.01% Triton X-100, 1 nM biotinylated 6
-His.

2. F-SSBct (FAM-WMDFDDDIPF), lyophilized. 3. SSBct peptide (WMDFDDDIPF), lyophilized. 4. SSBctΔF (WMDFDDDIP), lyophilized. 5. Centrifuge compatible with 384-well plates. 6. FA-compatible plate reader. 7. FA master mix: 10 mM HEPES-HCl [pH 7.5], 150 mM NaCl, 1 mM MgCl2, 1 mM DTT, 4.85 μM PriA, 0.01 μM F-SSBct.

3

Methods

3.1 Protein Expression

1. Transform Rosetta 2 competent cells with the PriA overexpression plasmid, plate on kanamycin/chloramphenicol plates, and grow at 37  C overnight. 2. Inoculate 100 mL of LB + 50 μg/mL kanamycin and 25 μg/mL chloramphenicol with a single colony of the transformed Rosetta 2 cells. Grow this culture at 37  C overnight with shaking. 3. Add 21 mL of the dense overnight culture to each of the four 1 L flasks of LB + 50 μg/mL kanamycin and 25 μg/mL chloramphenicol. Grow at 37  C with shaking.

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4. Monitor the growth of the culture by measuring the absorbance at 600 nm (OD600) with a spectrophotometer. Once the OD600 reaches 0.4–0.6, add 1 mM IPTG, continue to shake at 37  C for 4 h, and then harvest the cells by centrifugation at 10,000
g for 12 min. Discard the supernatant and transfer the cell pellets to a 50 mL conical tube. Multiple liters of cell culture can usually be combined into a single tube (see Note 11). 3.2 Protein Purification

1. All steps should be carried out at 4  C, unless otherwise noted. 2. Add lysis buffer to the cell pellet to a final volume of 45 mL. Using a metal spatula, resuspend and homogenize the cell pellet. Submerge the conical fully in an ice bath, and then sonicate the cells at 70% amplitude, for 40 s (0.8 s on, 0.2 s off). Invert the sample three times and allow to cool on ice for 1 min. Repeat the sonication and cooling steps twice. 3. Clarify the lysate by centrifugation (35,000
g at 4  C for 30 min). 4. While centrifuging the lysate, transfer 5 mL of Ni-NTA resin to a 50 mL conical tube, and add 30 mL of lysis buffer. After mixing, centrifuge for 5 min at 2500
g to pellet the resin and discard the supernatant. Repeat this wash once with another 30 mL of lysis buffer. Add the clarified cell lysate (from step 3) to the washed Ni-NTA resin and incubate with rocking for 1 h. 5. Pour the Ni-NTA resin into a gravity column and allow the solution to run through. To remove unbound proteins, wash the resin with a further 100 mL of lysis buffer. Do not allow the column to dry out. Once thoroughly washed, elute the protein in 20 mL of lysis buffer plus 300 mM imidazole. 6. To reduce the ionic strength of the elution buffer, add the 20 mL of eluted PriA to 180 mL of dilution buffer. Load over an SPFF column that has been equilibrated in buffer A. Elute PriA from the column by a gradient from buffer A to buffer B over 12 column volumes. Identify PriA-containing fractions and assess their purity by SDS-PAGE analysis. If further purification is required, the PriA-containing fractions can be pooled, concentrated by centrifugation, and then further purified by an S-300 size-exclusion column equilibrated with buffer B. 7. Determine the concentration of PriA using the A280 (MW ¼ 108,860 g/mol, ε ¼ 83,350 M*cm1), and adjust the concentration to 150 μM by centrifugation or dilution. Divide the PriA into 50 μL aliquots in 0.6 mL microcentrifuge tubes, flash freeze by dropping into liquid nitrogen, and store at 80  C until needed (see Note 12).

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AS Pilot Screen

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1. Follow the methods detailed in Subheadings 3.5 and 3.6 to perform the pilot AS using the selected pilot library and assess the data quality. 2. Review the screen with respect to the data quality. Sources of poor data quality (low Z0 scores, persistent plate effect, or variation when the same plate is repeated multiple times) need to be identified and corrected. 3. It is also important to review the screen with respect to workflow. Issues that appear minor during a pilot screen may become intolerable when repeated across dozens of screening plates. Some issues to consider include shared equipment (see Note 13), matching the speed of liquid-handling and data collection steps, and timing of breaks.

3.4 Selection of a Screening Library

Once a high-throughput screening assay has been developed, the most important consideration is which small-molecule library to screen. The size, chemical diversity, and quality of the library to be screened can impact the decisions made during the actual screening process [16]. The first consideration is the size of the chemical library. While inhibitors of PPIs have been developed, as a rule, they are more difficult to disrupt than other drug targets [17, 18]. Thus, to have the best chance of identifying a bona fide PPI inhibitor, the library should be as large as possible. Beyond the number of compounds available, the chemical diversity of a library must be considered as well. Small-molecule libraries are often made by a combinatorial chemistry approach, where sets of functionalized building blocks are linked together with different synthetic techniques. While the number of unique compounds that can be synthesized by this approach is massive, the amount of chemical space actually sampled may be much smaller. This occurs when many chemically similar compounds are linked by only a few methods [19]. Fortunately, techniques have been developed to assess the diversity of a chemical library and this diversity should be considered as well [20]. Another consideration for a screening library is the amenability of the compounds to secondary screening and follow-up work. Very likely, the initial hits identified by the HTS will be weakly active and must be improved upon through structure-activity relationship (SAR) studies. In an SAR study, the functional groups of a hit are systematically varied to identify which groups are important for activity and which can be improved. If the potency and specificity of early hits cannot be improved, you can be trapped in the unenviable position of having a weak, albeit real, inhibitor with no easy path forward. This is common with especially hydrophobic or insoluble compounds, where chemical modification is nearly impossible.

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Sets of rules have been developed to assess how “drug-like” a compound is. As an example, Lipinski’s rule considers the number of potential hydrogen bonds, size, and hydrophobicity of a potential drug [21]. While rules can be applied overzealously to exclude compounds that might otherwise be valuable, they can be a reasonable metric for how optimizable a starting hit might be. Look for libraries with high number of compliant compounds. Additionally, if a sufficiently diverse library was used for the primary screen, there should be relatively few structural analogs of any of the observed hits within the library. Many of the compounds should be commercially available to allow one to conduct “SAR-by-catalog” rather than relying on expensive on-demand chemical synthesis. A huge number of screening libraries are commercially available, and the simplest way to obtain a library is to purchase one. However, the up-front cost of a large library (>105 compounds) is likely to be prohibitively expensive to an academic lab. While it may be tempting to purchase as large of a library as possible and hope for the best, it is often better to seek out the services of a highthroughput screening core facility. These facilities are found at many larger research institutions and should have a large chemical library that is available for users to screen at cost. Core facilities will also have the infrastructure (e.g., low-volume liquid-handling capabilities, high-density plate readers) and the expertise to assist in troubleshooting any issues that may arise. If no such facility or library is available, it can be beneficial to perform in silico screening on a larger commercial library to enrich the purchased compounds [22, 23]. 3.5

AS

1. Dissolve the SSBct, Bio-SSBct, and SSBctΔF peptides in DMSO. Determine the protein concentration using the A280 and dilute each to 7.5 mM in DMSO. (a) SSBct: MW ¼ 1300 g/mol, ε ¼ 5500 M*cm1. (b) Bio-SSBct: MW ¼ 1527, ε ¼ 5500 M*cm1. (c) SSBctΔF: MW ¼ 1153 g/mol, ε ¼ 5500 M*cm1. 2. Using the Echo, prepare the DMSO control and screening plates (Fig. 2). For the DMSO control plates, add 10 nL of DMSO to every well of three 1536-well plates. For the screening plates, add 10 nL of the SSBctΔF peptide in column 1 of a 1536-well plate (final concentration will be 25 μM), 10 nL of SSBct in column 2 (25 μM), a serial dilution of SSBct to column 25 (three each of 0.2, 0.4, 0.8, 1.6, 3.2, 6.4, 12.8 nL, corresponding to 0.5, 1, 2, 4, 8, 16, 32 μM), and 10 nL of DMSO to the remainder of column 25 and column 26. In the remaining wells, plate 10 nL of a 10 mM compound stock in DMSO to a final concentration of 33 μM (see Note 14). Plates may be covered with a foil seal and stored at 80  C until ready for use.

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Fig. 2 Suggested plate map for 1536-well AS plates

3. Prepare the AS master mix fresh. Once prepared, add the AS donor and acceptor beads to 5 μg/mL each (see Note 15). Mix well, and then dispense 3 μL per well using the Mantis. Cover with a new foil plate seal and incubate at room temperature for 1 h with orbital shaking. 4. Centrifuge the plate briefly, carefully remove the foil seal while avoiding light exposure, and measure the AS signal across the plate. 3.6

Data Analysis

Once collected, the data must be normalized to account for plate and edge effects (Fig. 3, see Notes 16 and 17). If normalization data was collected during the pilot screen, there is no need to repeat it here. In this section, screening data will be normalized to correct for variations across the plate and then row-by-row variation. These are caused by the plate reader scanning pattern and temperature gradients across the plate, respectively. 1. Open an Excel file and copy the data from the three DMSOonly plates. On a new Excel sheet, make a 1536-well template, where the value for each well is the mean of reading from the three DMSO-only plates. In a new cell, take the mean value of the three entire plates. 2. In a new Excel sheet, again make a 1536-well format, and for each well, divide the mean DMSO read of that well by the mean reading of the entire DMSO plates. This is the DMSO correction factor (D) that will be applied to this well in all screening plates.

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Fig. 3 Data normalization reduces the impact of plate and edge effects. An example of the readings from a 1536-well AS plate before (left) and after (right) normalization. Red wells indicate inhibition of the AS reaction, while blue wells have higher signal. Note the correction of artifactual lower signal of the top row, the vertical gradient across the plate, and marked improvement in the Z0 score

3. Open the data set corresponding to your screening results. 4. For a given row of the screening data, rowi, calculate the mean value of each well within that row (x row i ). 5. Calculate the mean value of all the rows (x all rows ). 6. Create a new sheet with a 1536-well format. This will contain the final, corrected readings (xc). For each well in rowi, use Eq. 1 to obtain xc, i from the initial reading (x):
 x row i ð1Þ x c,i ¼ x∗D∗ x all row 7. Upload the corrected values to CDD vault or your screening database of choice. Likely, the software will calculate Z0 scores and % inhibitions automatically. If so, skip steps 8 and 10 (Subheading 3.6). 8. Calculate a Z0 score for each plate with Eq. 2 (see Note 18), where σ is the standard deviation, μ is the mean, and pc and nc denote the positive and negative controls, respectively [24]: 1 0   B3 σ pc þ σ nc C ð2Þ Z 0 ¼ 1  @   A μpc  μnc  9. Rerun any plates with Z0 scores 50% inhibition at 33 μM are considered hits (see Note 19):

HTS for Inhibitors of SSB-Protein Interactions.

2



X  μpc

127

3

5 % Inhibition of well X ¼ 100∗41   μnc  μpc

ð3Þ

11. Compile a list of hits and upload the hit list to the FAF-Drugs4 server [http://fafdrugs4.mti.univ-paris-diderot.fr/] to flag pan-assay interference compounds (PAINS) [25, 26]. Likely PAINS should be eliminated from further screening (see Note 20). 3.7 AS: Counterscreen (See Note 21)

1. Using the Echo, replate the non-PAINS, hit compounds onto a 1536-well plate (you will need a new plate map). Use the same concentration and location of controls as before. Make three total copies of this plate. 2. Prepare the counterscreening master mix and perform step 3 (Subheading 3.5). 3. Calculate a Z0 score for each plate and a % inhibition for each compound as outlined in Subheading 3.6. Compounds with >50% inhibition at 33 μM are highly likely to be acting as inhibitors of the AS chemistry or reagents and should be deprioritized.

3.8

FA Screening

Despite sifting and winnowing through both the primary and counterscreens, the majority of hits at this stage will still not be worth pursuing. It is worthwhile to use an addition screening step, known as a secondary screen, that uses an orthogonal assay to eliminate another set of false-positive hits. FA assays have been developed to monitor the interaction between the SSBct and a variety of interacting partners and they are readily amendable to miniaturization and HTS formats. 1. Dissolve the F-SSBct in DMSO and adjust the concentration to 5 mM (MW ¼ 1659 g/mol, ε ¼ 5690 M*cm1). 2. Into a black 384-well plate, add 2.5, 5, 10, 15, 20, 25, 30, 40, and 50 nL of 10 mM DMSO stock of each of the small molecules that were found to inhibit the PriA-SSB AS, but not the Bio-6
His counterscreen AS (Fig. 4). Additionally, add 25 nL of the 5 mM SSBct and SSBctΔF stocks to four wells each to serve as the positive and negative controls, respectively. Cover plate with a foil seal until ready to screen. 3. Prepare the FA master mix in the darkroom. 4. Remove the foil seal from the plate. Using the Mantis, add 10 μL of the master mix to each well. Cover with a foil plate seal and incubate with orbital shaking for 1 h. 5. Centrifuge the plate briefly, carefully remove the plate seal, and read the anisotropy and total fluorescence intensity from each well of the plate.

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Andrew F. Voter

Fig. 4 Suggested plate map for 384-well FA plates

6. Calculate the Z0 score for each plate; repeat any plate with Z0 < 0.5. 7. Calculate the average fluorescence intensity of each of the control reactions. Compounds that produce intensities >2
this average are likely fluorescent. Fluorescent molecules confound the FA reading and these compounds should be deprioritized. 8. Calculate the average % inhibition for each concentration of each compound. Using Prism, plot the concentration-response curve for each compound and calculate an IC50. Hits are classified as compounds with an IC50 < 33 μM and should be advanced to the hit validation step. 3.9

Hit Validation

1. Using the techniques outlined above, prepare three copies of AS plates with each of the compounds active in both the AS and FA assays. The compounds should be present in a 2
serial dilution (0.2–50 μM). 2. Run the AS and perform the data analysis as before. Using Prism, plot the concentration-response curve and calculate the IC50. The IC50 should be used to prioritize hits for further biophysical evaluation, with more potent compounds first. 3. Before using any of the remaining hits as tool compounds or undertaking extensive in vivo experiments, it is vital to obtain further evidence of the specificity of these compounds. After

HTS for Inhibitors of SSB-Protein Interactions.

129

multiple rounds of screening, the number of remaining compounds is likely small enough that lower throughput biophysical methods are practical. These methods are beyond the scope of this chapter but are reviewed elsewhere [27]. The optimal method of hit validation for a specific target will depend on the number of hit compounds remaining and the availability of instrumentation. NMR, isothermal titration calorimetry, differential scanning fluorimetry, surface plasmon resonance, microscale thermophoresis, and bio-layer interferometry have all been used to examine inhibitors of SSB PPIs.

4

Notes 1. The PriA gene is readily cloned from K. pneumoniae and may be inserted into any convenient overexpression plasmid. Alternatively, it may be requested from the Keck lab at the University of Wisconsin-Madison, Department of Biomolecular Chemistry. 2. AS kits are available with a wide range of functionalities. The AS nickel chelate detection kits are convenient if you prepare proteins with a 6
-His-tag for immobilized metal affinity chromatography purification. However, if your protein of interest is intolerant of a 6
-His-tag, other AS chemistries may be used. 3. In my experience, AS kits remain stable until opened. However, once opened, the signal intensity degrades to unacceptable levels over 1–2 weeks. While it may appear to be significantly cheaper to buy larger kits, be sure that the entire kit will be used within 1 week or the remaining reagents may be wasted. 4. These relatively short peptides may be purchased from commercial sources. This can be expensive but is more convenient than attempting to synthesize and achieve a high degree of labeling in-house. 5. While compound dispensing can theoretically be performed with more affordable alternatives such as multichannel pipets, in practice this rarely will be cost effective. DMSO concentrations must be held relatively low in the final assay mix and multichannels tend to have poor precision and accuracy at low volumes ( 0.5 are acceptable and it is difficult to interpret results from plates with lower Z0 scores. 19. In addition to being a rich source of puns for manuscript titles, PAINS are a subset of compounds that are active across

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Andrew F. Voter

multiple, independent HTS [25, 29]. PAINS act by a variety of mechanisms but tend to interfere with the assay chemistry rather than inhibiting the targeted biological activity [30]. There has been some debate in the field about the utility of discarding PAINS prior to further testing and there are reports of PAINS being successfully developed. Nevertheless, prioritizing known PAINS is a risky strategy that should only be undertaken if more promising leads cannot be identified. In these cases, it is imperative to run appropriate controls and validate the putative mechanism of action biophysically. 20. A typical primary screen should have a hit rate of P1;1JMC structure:1JMC: :B: :B::Homo sapiens:2.4: -------KVVPIASLTPYQSKWTICARVTNKSQIRTWSNSRGEGKLFSLELVDESGEIRATAFNEQVDKFFPLIEVNK VYYFSKGTLKIANKQFTAVKNDYEMTFNNETSVMPCEDD--HHLPTVQFDFTGIDDLENKSKDSLVDIIGICKSYEDATKITVRSNNREVAKRNIYLMDTSGKVV TATLWGEDADKFDGSRQPVLAIKGARVSDFGGRSLSVLSSSTIIANPDIPEAYKLRGWFDAEGQ*

13. model_build.py file used for LaRPA-1/OBF1-OBF2 molecular modeling: from modeller import * from modeller.automodel import * #from modeller import soap_protein_od

184

Carlos A. H. Fernandes et al. env = environ() a = automodel(env, alnfile=’align.ali’, knowns=’1JMC’, sequence=’UKNP’, assess_methods=(assess.DOPE, #soap_protein_od.Scorer(), assess.GA341)) a.starting_model = 1 a.ending_model = 1000 a.make()

14. align.ali file is used for LaRPA-1-OBF3 molecular modeling. Notice here and in the model_build.py file (see Note 15) that 1L1O and UKNP refer to the template and LaRPA-1 sequences, respectively. These two names must be the same here and in the model_build.py file (see Note 15): >P1;UKNP sequence:UKNP:1

:A:199

:A::::

- RGRKYLDEIQSDGIGRGLKPEYVDVRCVPIYFKQDAQWYDACPT -- CNKKVTEE GAQGDRFRCEKCDKTV -T PTQRYLVSIQVTDNVSQAWLTLFNEAGIEFFGMEAAELKRRA QEDPLYIAKLAQGRMNRPVVMRLRVKEETSSNAMTGEESDRLRMSVVRISEFMPIAGT SEETRRRLAQNLRTECDEILRLIEAY* >P1;1L1O structure:1L1O: :C: :C::Homo sapiens:2.8: -TNWKTLYEVKSENLGQGDKPDYFSSVATVVYLRKENCMYQACPTQDCNKKVIDQQN--GL YRCEKCDTEFPNFKYRMILSVNIADFQENQWVTCFQESAEAILGQNAAYLGELKDKNEQA FEEVFQNANFRSFIFRVRVKVETY ---------– IKATVMDVKPVDYR ----------------EYGRRLVMSIRRS*

15. model_build.py file used for LaRPA-1-OBF3 molecular modeling: from modeller import * from modeller.automodel import * #from modeller import soap_protein_od env = environ() a = automodel(env, alnfile=’align.ali’, knowns=’1L1O’, sequence=’UKNP’, assess_methods=(assess.DOPE, #soap_protein_od.Scorer(), assess.GA341)) a.starting_model = 1 a.ending_model = 1000 a.make()

Recombinant Leishmania sp. RPA-1

185

16. ions.mdp file (parameters for adding ions): ; ions.mdp - used as input into grompp to generate ions.tpr ; Parameters describing what to do, when to stop and what to save Integrator emtol

= steep

emstep

= 0.01

nsteps

= -1

; Algorithm (steep = steepest descent minimization)

; Stop minimization when the maximum force < 10.0 kJ/mol

= 1000.0

; Energy step size ; Maximum number of (minimization) steps to perform

; Parameters describing how to find the neighbors of each atom and how to calculate the interactions nstlist

= 1

; Frequency to update the neighbor list and long range forces

cutoff-scheme

= Verlet

ns_type

= grid

rlist

; Method to determine neighbor list (simple, grid)

= 1.0

; Cut-off for making neighbor list (short range forces)

coulombtype

= cutoff

; Treatment of long range electrostatic interactions

rcoulomb

= 1.0

; long range electrostatic cut-off

rvdw

= 1.0

; long range Van der Waals cut-off

pbc

= xyz

; Periodic Boundary Conditions

17. em.mdp file (parameters for energy minimization): title

= Minimization

; Title of run

; Parameters describing what to do, when to stop and what to save integrator emtol

= steep

; Algorithm (steep = steepest descent minimization) ; Stop minimization when the maximum force < 10.0 kJ/mol

= 100.0

emstep

= 0.01

; Energy step size

nsteps

= -1

; Maximum number of (minimization) steps to perform

; Parameters describing how to find the neighbors of each atom and how to calculate the interactions nstlist

= 1

; Frequency to update the neighbor list and long range forces

cutoff-scheme

= Verlet

ns_type

= grid

rlist

; Method to determine neighbor list (simple, grid)

= 1.2

; Cut-off for making neighbor list (short range forces)

coulombtype

= PME

rcoulomb

= 1.4

vdwtype

= cutoff

vdw-modifier

= force-switch

rvdw-switch

; Treatment of long range electrostatic interactions ; long range electrostatic cut-off

= 1.0

rvdw

= 1.4

; long range Van der Waals cut-off

pbc

= xyz

; Periodic Boundary Conditions

DispCorr

= no

18. nvt.mdp file (parameters for NVT equilibration):

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Carlos A. H. Fernandes et al. title

= NVT equilibration

; Run parameters Integrator

= md

nsteps

= 50000

; leap-frog integrator ; 2 fs * 50000 = 100 ps

dt

= 0.002

; 2 fs

; Output control nstenergy

= 500

; save energies every 1.0 ps

nstlog

= 500

; update log file every 1.0 ps

nstxout-compressed

= 500

; save coordinates every 1.0 ps

nstvout

= 500

; save velocities every 1.0 ps

continuation

= no

; first dynamics run

constraint_algorithm

= lincs

; holonomic constraints

constraints

= h-bonds

; bonds to H are constrained

lincs_iter

= 1

; accuracy of LINCS

lincs_orde

= 4

; also related to accuracy

; Bond parameters

; Neighbor searching and vdW cutoff-scheme

= Verlet

ns_type

= grid

nstlist

= 20

rlist

= 1.2

vdwtype

= cutoff

; search neighboring grid cells ; largely irrelevant with Verlet

vdw-modifier

= force-switch

rvdw-switch

= 1.0

rvdw

= 1.2

; short-range van der Waals cutoff (in nm)

; Electrostatics coulombtype

= PME ; Particle Mesh Ewald for long-range electrostatics

rcoulomb

= 1.2

; short-range electrostatic cutoff (in nm)

pme_order

=4

; cubic interpolation

fourierspacing

= 0.16

; grid spacing for FFT

; Temperature coupling tcoupl tc-grps

= V-rescale ; modified Berendsen thermostat = Protein Water_and_ions ; two coupling groups - more accurate

tau_t ref_t

= 0.1 0.1 = 310 310

; time constant, in ps ; reference temperature, one for each group, in K

; Pressure coupling pcoupl

= no

; no pressure coupling in NVT

; Periodic boundary conditions pbc

= xyz

; 3-D PBC

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; Dispersion correction is not used for proteins with the C36 additive FF DispCorr

= no

; Velocity generation gen_vel

= yes

; assign velocities from Maxwell distribution

gen_temp

= 310

; temperature for Maxwell distribution

gen_seed

= -1

; generate a random seed

19. To analyze the temperature progression, use the command gmx energy -f nvt.edr -o temperature.xvg and type 16.0 at the prompt to select the system temperature and exit. In the resulting graph (-o temperature.xvg), the system temperature should quickly reach the target value and remain stable for the remainder of the equilibration. If the temperature is not yet stabilized, additional time is required. 20. npt.mdp file (parameters for NPT equilibration): title

= NPT equilibration

; Run parameters Integrator

= md

; leap-frog integrator

nsteps

= 50000

; 2 fs * 50000 = 100 ps

dt

= 0.002

; 2 fs

; Output control nstenergy

= 500

; save energies every 1.0 ps

nstlog

= 500

; update log file every 1.0 ps

nstxout-compressed

= 500

; save coordinates every 1.0 ps

nstvout

= 500

; save velocities every 1.0 ps

continuation

= yes

; continuing from NVT

constraint_algorithm

= lincs

; holonomic constraints

constraints

= h-bonds

; bonds to H are constrained

lincs_iter

= 1

; accuracy of LINCS

lincs_order

= 4

; also related to accuracy

; Bond parameters

; Neighbor searching and vdW cutoff-scheme

= Verlet

ns_type

= grid

; search neighboring grid cells

Nstlist

= 20

; largely irrelevant with Verlet

Rlist

= 1.2

vdwtype

= cutoff

vdw-modifier

= force-switch

188

Carlos A. H. Fernandes et al. rvdw-switch

= 1.0

rvdw

= 1.2

; short-range van der Waals cutoff (in nm)

coulombtype

= PME

; Particle Mesh Ewald for long-range electrostatics

rcoulomb

= 1.2

; Electrostatics

pme_order

=4

; cubic interpolation

fourierspacing

= 0.16

; grid spacing for FFT

; Temperature coupling tcoupl

= V-rescale ; modified Berendsen thermostat

tc-grps

= Protein Water_and_ions ; two coupling groups - more accurate

tau_t

= 0.1 0.1

; time constant, in ps

ref_t

= 310 310

; reference temperature, one for each group, in K

; Pressure coupling pcoupl

= Berendsen ; pressure coupling is on for NPT

pcoupltype

= isotropic ; uniform scaling of box vectors

tau_p

= 2.0

ref_p

= 1.0

; reference pressure, in bar

compressibility

= 4.4e-5

; isothermal compressibility of water, bar^-1

refcoord_scaling

= com

; time constant, in ps

; Periodic boundary conditions pbc

= xyz

; 3-D PBC

; Dispersion correction is not used for proteins with the C36 additive FF DispCorr

= no

; Velocity generation gen_vel

= no

; velocity generation off after NVT

21. To analyze the density progression, use the command gmx energy -f npt.edr -o density.xvg and type 24.0 at the prompt to select the density of the system and exit. In the resulting plot (-o density.xvg), the density of the system must be stable over time to indicate that the system is well equilibrated. If the density is not yet stabilized, additional time is required. 22. md.mdp file (parameters for molecular dynamics simulation): title

= Protein MD simulation

; Run parameters integrator

= md

nsteps

= 50000000

; leap-frog integrator ; 2 fs* 50000000 = 100000 ps (100 ns)

dt

= 0.002

; 2 fs

Recombinant Leishmania sp. RPA-1

189

; Output control nstenergy

= 5000

nstlog

= 5000

; save energies every 10.0 ps ; update log file every 10.0 ps

nstxout-compressed

= 25000

; save coordinates every 10.0 ps

nstvout

= 5000

; save velocities every 10.0 ps

compressed-x-grps

= System

; replaces xtc-grps

; Bond parameters continuation

= yes

; continuing from NPT

constraint_algorithm = lincs

; holonomic constraints

constraints

= h-bonds

; bonds to H are constrained

lincs_iter

= 1

; accuracy of LINCS

lincs_order

= 4

; also related to accuracy

; Neighbor searching and vdW cutoff-scheme

= Verlet

ns_type

= grid

; search neighboring grid cells

nstlist

= 20

; largely irrelevant with Verlet

rlist

= 1.2

vdwtype

= cutoff

vdw-modifier

= force-switch

rvdw-switch

= 1.0

rvdw

= 1.2

; short-range van der Waals cutoff (in nm)

; Electrostatics coulombtype

= PME

rcoulomb

= 1.2

; Particle Mesh Ewald for long-range electrostatics

pme_order

=4

; cubic interpolation

fourierspacing

= 0.16

; grid spacing for FFT

; Temperature coupling tcoupl

= V-rescale ; modified Berendsen thermostat

tc-grps

= Protein Water_and_ions ; two coupling groups - more accurate

tau_t

= 0.1 0.1

; time constant, in ps

ref_t

= 310 310

; reference temperature, one for each group, in K

; Pressure coupling pcoupl

= Parrinello-Rahman

pcoupltype

= isotropic ; uniform scaling of box vectors

; pressure coupling is on for NPT

tau_p

= 2.0

; time constant, in ps

ref_p

= 1.0

; reference pressure, in bar

Compressibility

= 4.4e-5

; isothermal compressibility of water, bar^-1

; Periodic boundary conditions pbc

= xyz

; 3-D PBC

; Dispersion correction is not used for proteins with the C36 additive FF DispCorr

= no

190

Carlos A. H. Fernandes et al. ; Velocity generation gen_vel

= no

; continuing from NPT equilibration

; COM motion removal nstcomm comm-mode

= =

1 Linear

; frequency for COM motion removal ; removal type (Linear removes COM translation;

Angular removes COM translation and rotation; None no removal) comm-grps

=

Protein

Water_and_ions; groups for COM removal

23. To evaluate the energy stabilization of the model after the molecular dynamics simulations, perform a root mean square deviation (RMSD) analysis. After the trajectory processing, type the command gmx rms -f md_2.xtc -s md.tpr -o rms.xvg -n index.ndx -tu ns and choose option 4 (backbone). The -tu flag will output the results in terms of ns, although the trajectory was written in ps. In the resulting plot (-o rms.xvg), it will be shown the RMSD relative over time. Low variation values of RMSD (~0.1 nm) in the last 30 ns of the molecular dynamics simulations may indicate that the structure is energetically minimized in a local minimum value. If the RMSD is not yet stabilized, additional time is required. References 1. Wold MS (1997) Replication protein A: a heterotrimeric, single-stranded DNA-binding protein required for eukaryotic DNA metabolism. Annu Rev Biochem 66:61–92 2. Smith J, Zou H, Rothstein R (2000) Characterization of genetic interactions with RFA1: The role of RPA in DNA replication and telomere maintenance. Biochimie 82:71–78 3. Fanning E, Klimovich V, Nager AR (2006) A dynamic model for replication protein A (RPA) function in DNA processing pathways. Nucleic Acids Res 34:4126–4137 4. Bochkarev A, Pfuetzner RA, Edwards AM, Frappier L (1997) Structure of the singlestranded-DNA-binding domain of replication protein A bound to DNA. Nature 485:176–181 5. Bochkareva E, Korolev S, Lees-Miller SP, Bochkarev A (2002) Structure of the RPA trimerization core and its role in the multistep DNA-binding mechanism of RPA. EMBO J 21:1855–1863 6. Fan J, Pavletich NP (2012) Structure and conformational change of a replication protein A heterotrimer bound to ssDNA. Genes Dev 26:2337–2347 7. Binz SK, Wold MS (2008) Regulatory functions of the N-terminal domain of the 70-kDa

subunit of replication protein A (RPA). J Biol Chem 283:21559–21570 8. Mitton-Fry RM, Anderson EM, Hughes TR, Lundblad V, Wuttke DS (2002) Conserved structure for single-stranded telomeric DNA recognition. Science 296:145–147 9. Chan SWRL, Blackburn EH (2004) Telomeres and telomerase. Philos Trans R Soc Lond B 359:109–121 10. Rice C, Skordalakes E (2016) Structure and function of the telomeric CST complex. Comput Struct Biotechnol J 14:161–167 11. Palm W, de Lange T (2008) How shelterin protects mammalian telomeres. Annu Rev Genet 42:301–334 12. Dewar JM, Lydall D (2012) Similarities and differences between “uncapped” telomeres and DNA double-strand breaks. Chromosoma 12:117–130 13. Miyake Y, Nakamura M, Nabetani A, Shimamura S, Tamura M, Yonehara S, Saito M, Ishikawa F (2009) RPA-like Mammalian Ctc1-Stn1-Ten1 complex binds to singlestranded DNA and protects telomeres independently of the Pot1 pathway. Mol Cell 36:193–206 14. Price CM, Boltz KA, Chaiken MF, Stewart JA, Beilstein MA, Shippen DE (2010) Evolution of

Recombinant Leishmania sp. RPA-1 CST function in telomere maintenance. Cell Cycle 9:3157–3165 15. Lewis KA, Wuttke DS (2012) Telomerase and telomere-associated proteins: Structural insights into mechanism and evolution. Structure 20:28–39 16. Lue NF (2018) Evolving linear chromosomes and telomeres: a C-strand-centric view. Trends Biochem Sci 43:314–326 17. Lopes AH, Souto-Padro´n T, Dias FA, Gomes MT, Rodrigues GC, Zimmermann LT, Alves e Silva TL, Vermelho AB (2010) Trypanosomatids: odd organisms, devastating diseases. Open Parasitol J 4:30–59 18. Neto JLS, Lira CBB, Giardini MA, Khater L, Perez AM, Peroni LA, dos Reis JRR, FreitasJunior LH, Ramos CHI, Cano MIN (2007) Leishmania replication protein A-1 binds in vivo single-stranded telomeric DNA. Biochem Biophys Res Commun 358:417–423 19. Pavani RS, Fernandes C, Perez AM, Vasconcelos EJR, Siqueira-Neto JL, Fontes MR, Cano MIN (2014) RPA-1 from Leishmania amazonensis (LaRPA-1) structurally differs from other eukaryote RPA-1 and interacts with telomeric DNA via its N-terminal OB-fold domain. FEBS Lett 588:4740–4748 20. Pavani RS, da Silva MS, Fernandes CAH, Morini FS, Araujo CB, Fontes MR d M, Sant’Anna OA, Machado CR, Cano MI, Fragoso SP, Elias MC (2016) Replication protein A presents canonical functions and is also involved in the differentiation capacity of Trypanosoma cruzi. PLoS Negl Trop Dis 10:e0005181 21. Da Silveira RDCV, Da Silva MS, Nunes VS, Perez AM, Cano MIN (2013) The natural absence of RPA1N domain did not impair Leishmania amazonensis RPA-1 participation in DNA damage response and telomere protection. Parasitology 140:547–559 22. Brown GW, Melendy TE, Ray DS (1992) Conservation of structure and function of DNA replication protein A in the trypanosomatid Crithidia fasciculata. Proc Natl Acad Sci U S A 89:10227–10231 23. Ferna´ndez MF, Castellari RR, Conte FF, Gozzo FC, Sabino AA, Pinheiro H, Novello JC, Eberlin MN, Cano MIN (2004) Identification of three proteins that associate in vitro with the Leishmania (Leishmania) amazonensis G-rich telomeric strand. Eur J Biochem 271:3050–3063 24. Pavani RS, Vitarelli MO, Fernandes CAH, Mattioli FF, Morone M, Menezes MC, Fontes

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MRM, Cano MIN, Elias MC (2018) Replication protein A-1 has a preference for the telomeric G-rich sequence in Trypanosoma cruzi. J Eukaryot Microbiol 65:345–356 25. Fernandes CAH, Morea EGO, dos Santos GA, da Silva VL, Vieira MR, Viviescas MA, Chatain J, Vadel A, Saintome´ C, Fontes MRM, Cano MIN (2020) A multi-approach analysis highlights the relevance of RPA-1 as a telomere end-binding protein (TEBP) in Leishmania amazonensis. Biochim Biophys Acta Gen Subj 1864:129607 26. Eswar N, John B, Mirkovic N, Fiser A, Ilyin VA, Pieper U, Stuart AC, Marti-Renom MA, Madhusudhan MS, Yerkovich B, Sali A (2003) Tools for comparative protein structure modeling and analysis. Nucleic Acids Res 31:3375–3380 27. Berendsen HJC, van der Spoel D, van Drunen R (1995) GROMACS: A message-passing parallel molecular dynamics implementation. Comput Phys Commun 91:43–56 28. The PyMol Molecular Graphics System, Version 2.0 29. Pettersen EF, Goddard TD, Huang CC, Couch GS, Greenblatt DM, Meng EC, Ferrin TE (2004) UCSF Chimera—a visualization system for exploratory research and analysis. J Comput Chem 25:1605–1612 30. Humphrey W, Dalke A, Schulten K (1996) VMD: Visual molecular dynamics. J Mol Graph 14:33–38 31. So¨ding J (2005) Protein homology detection by HMM-HMM comparison. Bioinformatics 27:951–960 32. Oostenbrink C, Soares TA, Van Der Vegt NFA, Van Gunsteren WF (2005) Validation of the 53A6 GROMOS force field. Eur Biophys J 34:273–284 33. Kelley LA, Mezulis S, Yates CM, Wass MN, Sternberg MJE (2015) The Phyre2 web portal for protein modeling, prediction and analysis. Nat Protoc 10:845–858 34. Yang J, Yan R, Roy A, Xu D, Poisson J, Zhang Y (2014) The I-TASSER suite: protein structure and function prediction. Nat Methods 12:7–8 35. Lira CBB, Gui KE, Perez AM, da Silveira RCV, Gava CH, Ramos I, Cano MIN (2009) DNA and heparin chaperone the refolding of purified recombinant replication protein A subunit 1 from Leishmania amazonensis. Biochim Biophys Acta 1790:119–125

Chapter 11 Single-Stranded DNA Curtains for Single-Molecule Visualization of Rad51-ssDNA Filament Dynamics Upasana Roy and Eric C. Greene Abstract Homologous recombination (HR) is a highly conserved DNA repair pathway required for the accurate repair of DNA double-stranded breaks. DNA recombination is catalyzed by the RecA/Rad51 family of proteins, which are conserved from bacteria to humans. The key intermediate catalyzing DNA recombination is the presynaptic complex (PSC), which is a helical filament comprised of Rad51-bound singlestranded DNA (ssDNA). Multiple cellular factors either promote or downregulate PSC activity, and a fine balance between such regulators is required for the proper regulation of HR and maintenance of genomic integrity. However, dissecting the complex mechanisms regulating PSC activity has been a challenge using traditional ensemble methods due to the transient and dynamic nature of recombination intermediates. We have developed a single-molecule assay called ssDNA curtains that allows us to visualize individual DNA intermediates in real-time, using total internal reflection microscopy (TIRFM). This assay has allowed us to study many aspects of HR regulation that involve complex and heterogenous reaction intermediates. Here we describe the procedure for a basic ssDNA curtain assay to study PSC filament dynamics, and explain how to process and analyze the resulting data. Key words DNA curtains, Single molecule, Homologous recombination, Single-stranded DNA, Rad51, RPA, Srs2

1

Introduction

1.1 Homologous Recombination

Homologous recombination (HR) is a critical DNA repair pathway required for the repair of double-stranded DNA breaks (DSBs) and recovery of stalled or collapsed replication forks [1, 2]. Defects in HR can lead to loss of genomic integrity, and have been associated with multiple types of cancer and cancer-prone syndromes [3, 4]. However, excessive or inappropriate HR also leads to chromosomal instability [5], highlighting the importance of precisely regulating HR in vivo. DNA recombination during HR is carried out by the highly conserved RecA/Rad51 family of recombinases, which bind ssDNA generated at the DSB site, to form a helical filament called the presynaptic complex (PSC). This is the key intermediate in HR, responsible for carrying out homology search

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_11, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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i. DSB

ii. 3’ ssDNA 3’

iii.

5’

5’ 3’

RPA–ssDNA 3’

5’

iv. 5’

3’

Rad51–ssDNA 5’

v.

5’

D-loop intermediate 5’

vi. 5’

Homologous DNA

Fig. 1 Early steps of homologous recombination (HR). A highly simplified version of the initial steps of the HR pathway is shown. Genomic DNA (i) can be damaged by a variety of sources to generate a double-stranded break (DSB) (ii). Nucleases resect DNA around the DSB, producing 30 ssDNA overhangs (iii) which are rapidly bound by the ssDNA-binding protein RPA (replication protein A) (iv). Rad51 recombinase displaces RPA to form a Rad51-ssDNA filament (v), which searches the genome for a homologous DNA template. Rad51-ssDNA filaments invade the donor duplex to form a D-loop intermediate (vi). The 30 end is then extended by DNA polymerases based on the homologous template, and downstream intermediates are processed by DNA resolution/dissolution factors

and DNA pairing with the correct DNA template in the genome, thereby ensuring accurate repair [1]. In S. cerevisiae, DNA around the DSB is resected by nucleases to expose 30 ssDNA overhangs, which are rapidly bound by the ssDNA-binding protein RPA (replication protein A) (Fig. 1). Rad51 displaces RPA with the help of additional HR factors to form the PSC, which searches the genome for homologous target sequences. Once the PSC locates the correct target DNA duplex, the broken 30 ssDNA end is paired with the

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complementary DNA template and then extended by DNA polymerases. The resulting strand invasion recombination intermediate can then be resolved/dissolved in a number of ways by various nucleases [2, 6, 7]. DNA recombination needs to be precisely regulated spatially and temporally to maintain the genomic integrity of a cell. As a result, there exist multiple regulators of HR including mediators that promote HR, and anti-recombinases that suppress aberrant HR. A bona fide anti-recombinase is the S. cerevisiae 30 –50 translocase Srs2 that acts at replication forks to disrupt HR intermediates, and instead promotes repair by translesion synthesis (TLS) [8]. Mechanistically, Srs2 suppresses HR by disrupting DNA recombination intermediates, as well as displacing bound Rad51 from ssDNA [9–12]. 1.2 Single-Molecule Studies of HR

Genetic studies have provided the framework for our current understanding of the role of HR regulators. However, obtaining a clear understanding of how these factors operate at the mechanistic level has been difficult. Using traditional bulk biochemical experiments to address such questions has proven to be challenging, as HR proceeds through highly dynamic and transient intermediates (Fig. 1). As a result, ensemble studies that rely on population averaging are unable to dissect how individual HR intermediates are regulated by regulatory factors. Single-molecule (SM) experiments are a powerful way to obtain highly refined information about complex reactions that involve heterogenous reaction intermediates. Importantly, they also allow detection of distinct subpopulations, or rare and transient intermediates that arise during the reaction. Recently, singlemolecule studies have revealed a wealth of information about how each step in HR is regulated. In particular, given the central role of PSC in mediating recombination, there have been multiple studies aimed at understanding the regulation of PSC. However, there are a few common challenges in using single-molecule assays. Firstly, nonspecific adsorption of DNA/proteins to the anchoring surface is a common problem. Secondly, many SM experiments are typically low throughput in nature and may not provide enough data to be statistically relevant.

1.3

Our lab has developed a SM technique called DNA curtains that allows real-time TIRF-based imaging of hundreds of individual ssDNA complexes anchored to a fluid lipid bilayer [13–17]. Our setup overcomes some of the common problems associated with SM methods while still providing highly detailed mechanistic insight into DNA-based processes. The instrumentation and experimental setup for DNA curtains has been described in detail previously [13–17]. Briefly, chrome is deposited onto quartz slides in nanoscale patterns of barriers and

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anchors using E-beam lithography. These patterned slides are used to assemble flow cells—a microfluidic device with nanoports, through which buffers can be flowed in and out (Fig. 2a, b). Biotinylated lipids are used to coat the slide surface in the microfluidic chamber, thereby providing a fluid medium for ssDNA molecules to diffuse, and preventing nonspecific adsorption of the DNA molecules onto the quartz surface (Fig. 2c). Streptavidin is then used to anchor biotinylated single-stranded DNA (ssDNA)

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Fig. 2 ssDNA curtain schematic. (a) Flow cell with nanoports and central sample chamber. (b) The patterned quartz slide forms the base of the central sample chamber, and contains nanoscale chrome barriers and anchors deposited by E-beam lithography. (c) Biotinylated lipids form a bilayer that passivates the quartz slide surface. (d) Biotinylated ssDNA is attached to the lipid layer via streptavidin and stretched to the downstream anchor points by buffer flow and RPA binding. ssDNA can be visualized using labeled RPA by TIRFM. (e) Rad51 and ATP are added to the sample chamber, leading to displacement of RPA and formation of Rad51-ssDNA, which is visualized by the loss of RPA signal

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onto the biotinylated lipid layer. Buffer flow is used to align the ssDNA molecules against the chrome barriers, which act as a barrier to lipid diffusion. The ssDNA is stretched out to the downstream attachment anchor by a combination of buffer flow, urea denaturation, and addition of RPA (which removes the secondary structures of ssDNA). Once attached to the downstream anchor, the ssDNA is tethered at both ends, and can be used for experiments even in the absence of flow (Fig. 2d). The stretched doubletethered ssDNA is visualized by binding of mCherry-RPA using TIRFM. A broad overview for ssDNA curtain setup is shown in Fig. 3, and all nanofabrication and instrumentation procedures have been described elsewhere [14, 16]. In this chapter we describe in detail

ssDNA Curtain Overview

a Nanofabrication Preparation of patterned slides by E-beam lithography [13,16]

b Stock Solutions Preparation of ssDNA and biotinylated lipid stocks [18]

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iv) Inject proteins v) Record data vi) Analyze data

Fig. 3 Overview of ssDNA curtain methodology. Shown here is a general overview of the different aspects of ssDNA curtain experiments [13–18]. (a) Nanofabrication. Chrome patterns are deposited onto quartz slides by E-beam lithography [13, 16]. (b) Preparation of ssDNA and lipid stock solutions required for DNA curtains [18]. (c) ssDNA curtains are visualized by prism-based TIRF microscopy [14, 16]. (d) Patterned slides from (a) are used to assemble flow cells, which are microfluidic devices with a central sample chamber and nanoports for flowing buffers in and out [16]. (e) Flow cells are then used for daily experiments (described in this chapter in detail). The workflow involves passivating the quartz surface with lipids, attaching ssDNA to the lipid bilayer, mounting the flow cell on a prism-based TIRF microscope, and connecting it to the fluidics system. Proteins of interest are injected into the sample chamber, and the experiment is recorded and analyzed to obtain singlemolecule, high-throughput information about the reaction

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the daily workflow for studying Rad51 filament dynamics and Srs2 activity using ssDNA curtains (Fig. 3e). We also discuss how to process and analyze the resulting data.

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Materials Prepare all solutions in ultrapure Milli-Q water and thoroughly clean tubing and connectors with ultrapure water before use. Prepare ssDNA and liposome stock solutions as described in [18] (Fig. 3b). 1. Lipid buffer: 10 mM Tris–HCl [pH 8], 100 mM NaCl. 2. Biotinylated lipid stock solution: 100 mg/mL DOPC, 10 mg/ mL PEG-2000 DOPE, 0.5 mg/mL biotinylated DOPE [18]. 3. BSA buffer: 40 mM Tris–HCl [pH 8], 2 mM MgCl2, 1 mM DTT, 0.2 mg/mL BSA. 4. HR buffer: 30 mM Tris–Ac [pH 7.5], 50 mM KCl, 20 mM MgAc2, 1 mM DTT, 0.2 mg/mL BSA. 5. 1 mg/mL Streptavidin solution. 6. ssDNA stock solution in 50 mM Tris–HCl [pH 7.5], 4 mM DTT, 10 mM ammonium sulfate, 10 mM MgCl2 [18]. 7. 2% Hellmanex solution. 8. 1 N NaOH. 9. 1 mL Syringes with slip tip. 10. 3 mL Syringes with Luer-lock tip. 11. 10 mL Syringes with Luer-lock tip. 12. 30 mL Syringes with Luer-lock tip. 13. Teflon PFA resin tubing 1/1600 OD  0.0200 ID. 14. PEEK Headless short 10–32 coned connector for 1/1600 OD. 15. Two-piece finger-tight fitting, 10–32 coned, for 1/1600 OD. 16. PEEK female-to-female Luer adapter, 10–32. 17. Kd Scientific Dual Syringe Pump (KDS-201). 18. High-pressure switch valve. 19. 1 μM Yeast RPA-mCherry stock in 30 mM NaHPO4 [pH 7.5], 250 mM KCl, 10% glycerol, 0.02% Tween-20, 1 mM TCEP, 0.5 mM EDTA, 0.25% myoinositol [18]. 20. yeast Rad51 stock in 25 mM Tris–Cl [pH 7.5], 10% glycerol, 200 mM NaCl, 1 mM DTT [19]. 21. yeast GFP–Srs2 stock in 40 mM NaHPO4 [pH 7.5], 300 mM KCl, 10% glycerol, 0.01% Tween-20, 1 mM TCEP, 0.5 mM EDTA, 0.125% myoinositol [18].

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Methods We have published detailed procedures on the different aspects of a DNA curtain setup (Fig. 3a–d) [13–18], and here we focus on the everyday workflow of ssDNA curtain experiments, as well as image and data analysis (Fig. 3e).

3.1 Lipid Passivation and ssDNA Attachment

1. The flow cell is first flushed with water, and air bubbles are removed before injection of lipid solutions. For this, cut two 10 cm pieces of tubing. On one end of each tube, attach a connector (to be used for attachment to nanoports on the flow cell). On the other end of each tube, attach a connector and then a female-to-female Luer peek adaptor. This end will be used to connect the syringes and microfluidics. 2. Take a 10 mL syringe and fill with Milli-Q water. Remove air bubbles from the syringe and attach it to one end of a prepared tubing. Push 2–3 mL of water through the tubing to remove any air. With the syringe attached, connect the other end of the tubing to port A on the flow cell (Fig. 2a). 3. Take the second piece of tubing, and attach it to the flow cell via port B (Fig. 2a). 4. Push 2–3 mL water from the syringe attached to port A, flushing the flow cell and tubing with water. 5. Take a second 10 mL syringe and fill it with Milli-Q water, being careful to remove all air bubbles. Attach this syringe to the port B via the connected tubing using drop-to-drop connections. 6. Now that both syringes are attached, gently push water back and forth between the two syringes to flush the sample chamber, and remove any trapped air bubbles. Make sure to remove all air from the flow cell and tubing (see Note 1). This takes about 5–10 min. 7. Take a 3 mL syringe and fill it with lipid buffer, being careful to exclude air bubbles. Remove the 10 mL syringe attached to port A of the flow cell. Using the 10 mL syringe still connected to port B, gently push a droplet of water and attach the 3 mL syringe by drop-to-drop connection (see Note 2). Disconnect the 10 mL syringe on port B. 8. Push 2.5 mL lipid buffer through the sample chamber, flushing out the water and replacing it with lipid buffer. 9. Make a lipid dilution by mixing 40 μL of biotinylated liposome stock solution [18] with 960 μL lipid buffer. Fill a 1 mL syringe with this lipid dilution being careful to exclude air bubbles, and attach it to port B. Disconnect the syringe attached to port A.

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10. Using the attached 1 mL syringe on port B, gently push ~200 μL of the lipid solution into the sample chamber. Incubate for 5–8 min. Repeat until all the lipid solution is used up. 11. Fill a 3 mL syringe with lipid buffer, and attach it to port A. Disconnect the 1 mL syringe on port B. 12. Using the attached 3 mL syringe on port A, gently push 2.5 mL lipid buffer through the sample chamber. Incubate for 20 min at room temperature to allow lipid bilayer to passivate the flow cell surface. 13. Attach a 3 mL syringe filled with BSA buffer at port B. Detach the syringe on port A, and gently push 2.5 mL BSA buffer through the sample chamber. 14. Dilute streptavidin solution by mixing 10 μL streptavidin stock solution (1 mg/mL) in 990 μL BSA buffer. Fill a 1 mL syringe with the streptavidin dilution and attach at port A. Gently flush sample chamber with ~400 μL streptavidin solution and incubate at room temperature for 5 min. Repeat with the leftover solution. 15. Fill a 3 mL syringe with BSA buffer and replace the syringe at port B. Detach syringe connected to port A, and gently flush 2.5 mL BSA buffer to remove unbound streptavidin. 16. Dilute 10 μL of the ssDNA stock [18] by adding 1 mL of BSA buffer. Fill a 1 mL syringe gently with ssDNA solution and attach to port A (see Note 3). Gently push ~200 μL ssDNA solution into the sample chamber (without disconnecting the second syringe; see Note 4). Incubate for 5–8 min at room temperature, and repeat until all the ssDNA has been injected. 17. Once all the ssDNA is injected, the flow cell can then be mounted on the microscope and connected to the sample injection system. It is critical to make all attachments to the fluidic system via drop-to-drop connections as well (see Note 5). 18. The flow cell chamber is maintained at a temperature of 30  C for all experiments with yeast recombinant proteins. 3.2 Labeling ssDNA Using Fluorescently Tagged RPA

ssDNA in the sample chamber can be visualized by injection of GFP- or mCherry-labeled RPA, as it binds ssDNA with a high affinity. Secondary structures in the ssDNA are disrupted by injection of urea and RPA, and ssDNA molecules are extended during this time by the application of buffer flow at a high rate of 0.8 mL/ min. Together, this strategy fully extends the ssDNA allowing extensive binding by RPA and attachment to the downstream anchor points (Fig. 2d).

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1. Dilute RPA-mCherry stock [18] with HR buffer to a final concentration of 0.1 nM. Fill a 30 mL syringe with the diluted RPA solution being careful to avoid bubbles. 2. Fill a second 30 mL syringe with HR buffer supplemented with 2 mM ATP. Attach both syringes to the sample injection syringe pump, such that HR buffer + RPA is connected to the flow cell, and HR buffer + ATP goes to discard. 3. Flush sample chamber with RPA solution at a rate of 0.8 mL/ min. Immediately, also inject 500 μL 7 M urea solution into the sample chamber. Allow RPA solution to flow for 8–10 min. The combination of urea, RPA, and buffer flow removes secondary structures and extends the ssDNA, allowing attachment to the downstream anchor points. 4. Once RPA-mCherry is bound, the ssDNA curtain can be visualized by TIRFM. RPA-ssDNA is the physiological substrate for Rad51 binding, and forms the starting point for our studies on Rad51 dynamics. 5. Using a low laser setting (~20 mW) visualize RPA-ssDNA in the presence of buffer flow, and select a field of view with wellresolved ssDNA strands (see Note 6). It is best to choose a set of barriers where the ssDNA is not too crowded, to avoid analysis of overlapping ssDNA molecules. This will be used as the field of view for recording purposes. 3.3 Measuring Rad51 Assembly and Disassembly Kinetics

1. Flush the sample chamber with HR buffer containing ATP at a flow rate of 0.8 mL/min for at least 2 min (see Note 7). 2. Begin recording data 1–2 min before you are ready to inject Rad51. We use a laser power of 50 mW and image integration time of 100 ms. Data shown in this chapter have been acquired by recording an image every 10 s. 3. Inject 4 μM yRad51 (or desired concentration) in HR buffer containing ATP, and stop flow to allow Rad51 to assemble onto RPA-coated ssDNA. The Rad51 filament assembly reaction can be visualized by the continuous loss of RPA-mCherry signal. We typically incubate Rad51 for 15–20 min at this step, but the duration of incubation will depend on the Rad51 concentration used (Fig. 4a). 4. Flush the sample chamber with buffer containing ATP at a flow rate of 0.8 mL/min for at least 2 min to wash off any unbound Rad51. The sample chamber now contains Rad51-ssDNA filaments which can be used as a starting point for studying the binding and activity of downstream HR factors. 5. We have also used ssDNA curtains to measure the rate of Rad51 disassembly after ATP depletion, as an indicator of Rad51-ssDNA filament stability. For disassembly experiments,

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Fig. 4 Visualization of Rad51 filament assembly and disassembly. (a) Kymographs showing loss of RPA-mCherry signal when 4 μM Rad51 is injected. Time of injection is indicated by a white dashed line. (b) Graph showing kinetics of RPA-mCherry after 4 μM Rad51 injection (n ¼ 33). Mean normalized intensities are plotted and shaded area represents 95% CI. (c) Kymograph showing rebinding of RPA-mCherry due to Rad51ssDNA disassembly. Time of ATP depletion is indicated with a white dashed line. (d) Graph showing kinetics of RPA-mCherry rebinding during Rad51-ssDNA disassembly (n ¼ 28)

flush sample chamber with HR buffer lacking ATP, but supplemented with 0.1 nM RPA-mCherry, at a flow rate of 0.8 mL/ min for 1 min. This step rapidly washes away ATP already in the sample chamber. 6. Reduce flow rate to 0.2 mL/min, and continuously flow in the buffer lacking ATP, supplemented with RPA-mCherry. 7. Disassembly of the Rad51 filament can be visualized by the rebinding of free RPA-mCherry to the ssDNA exposed by Rad51 dissociation (Fig. 4c). 3.4 Data Analysis for Rad51-ssDNA Assembly and Disassembly Kinetics

The loss of RPA-mCherry intensity is used to calculate the assembly rate of Rad51. Similarly, the rebinding of RPA-mCherry after ATP depletion is used to calculate the rate of Rad51 disassembly. Images are acquired through Nikon NIS Elements software, and each frame of TIRFM data is exported as a raw TIFF image. These image files are analyzed using Fiji software (ImageJ 1.48b, Wayne Rasband, National Institutes of Health, USA).

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1. Combine TIFF files from one channel into a single image stack. 2. The image stack is used to generate kymographs of individual ssDNA molecules for data analysis. A kymograph is the 2D image of a selected ssDNA molecule, where the Y-axis represents position along the ssDNA, and the X-axis represents time. To generate kymographs, draw a 1-pixel wide ROI (region of interest) on an individual ssDNA molecule and use the “Reslice” function in Fiji (see Note 8). 3. On the kymograph generated, draw a rectangular ROI selecting the whole image. Use the “Plot Profile” function to produce a list of the average intensity value at each time point in the kymograph (see Note 9). 4. Divide each value in the list by the maximum intensity value, to generate a list of the “normalized intensity” value for each time point. 5. Plot the mean and standard error of the resulting distribution as an X–Y scatterplot where the Y-axis represents mean normalized RPA intensity, and the X-axis represents time (we use GraphPad Prism software) (Fig. 4b). 6. The rate of Rad51 disassembly is measured by the rate of RPA-mCherry rebinding, and is calculated in the same way (Fig. 4d). 3.5 Rad51 Disruption by Srs2

We have also used ssDNA curtains to study the activity of the antirecombinase Srs2, and its regulation of Rad51 filament dynamics [12, 20]. In our assays, labeled or unlabeled Srs2 is co-injected with labeled RPA onto Rad51-ssDNA filaments. Srs2 translocates along the filaments displacing Rad51 from ssDNA, which is visualized by the rapid rebinding of free labeled RPA in the buffer. By imaging the trajectories of Srs2, we can calculate its translocation velocities. 1. Prepare Rad51-ssDNA filaments as described in Subheadings 3.1–3.3, step 4. 2. Using a 150 μL sample loop, inject 0.1 nM GFP-Srs2 [18] in HR buffer supplemented with 2 mM ATP and 0.1 nM RPA-mCherry, at the rate of 0.2 mL/min. Translocation of Srs2 can be observed by the GFP signal, and disruption of Rad51 from ssDNA can be visualized by rebinding of RPA-mCherry behind translocating GFP-Srs2 molecules (Fig. 5a) [12]. 3. Generate kymographs for individual ssDNA filaments as described in Subheading 3.4, to be used for calculating the velocity of GFP-Srs2 translocation. 4. Using the “multi-point” tool in Fiji, mark the start and end point of a single GFP-Srs2 trajectory in a kymograph (see Note 10).

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Fig. 5 Visualization of Rad51 disruption by translocating Srs2. (a) Kymograph showing GFP-Srs2 translocation on Rad51-ssDNA filaments. GFP-Srs2 (green) disrupts Rad51-ssDNA filaments as it translocates, exposing naked ssDNA which is rapidly bound by free RPA-mCherry (magenta) in the buffer. (b) Quantification of GFP-Srs2 velocities (panels reproduced with permission from Kaniecki, K., De Tullio, L., Gibb, B., Kwon, Y.- H., Sung, P., & Greene, E. C. (2017). Dissociation of Rad51 presynaptic complexes and heteroduplex DNA joints by tandem assemblies of Srs2. Cell Reports, 21(11), 3166–3177)

5. Using the “Analyze” tool, retrieve the X and Y coordinates of the two marked points: X1Y1 and X2Y2. 6. GFP-Srs2 translocation velocity is calculated from the number of pixels translocated per unit time ¼ (Y2 – Y1)/(X2 – X1) for each GFP-Srs2 molecule. In this experiment each position pixel in the Y-axis corresponds to ~725 nucleotides (nt) of Rad51ssDNA [18], and each time pixel in the X-axis corresponds to 10 s (since frame rate is 0.1 fps). Using this conversion factor, calculate the Srs2 velocity in units of nt/s. 7. The velocity data can be plotted as a frequency distribution, and fitted to a Gaussian curve to retrieve the mean velocity of GFP-Srs2 translocation (Fig. 5b) [12]. 3.6 Disassembly and Cleaning of Slides

Patterned slides can be reused multiple times after disassembling the flow cells and cleaning the slides. 1. Store the flow cells in ethanol for 48–72 h, until the glued nanoports fall off. 2. Carefully remove the coverslip and double-sided tape with a blade. Make sure to remove all residual adhesive leftover on the patterned slide (see Note 11). 3. Wash the patterned slides with fresh ethanol for 1 h using a stir bar. 4. Rinse the slides thoroughly with ultrapure distilled water to remove all traces of ethanol. 5. Wash the slides in 2% Hellmanex solution overnight using a stir bar (see Note 12).

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6. Wash slides in ultrapure distilled water for 30 min using a stir bar. 7. Wash slides in 1 N NaOH solution for 1 h using a stir bar. 8. Wash slides in ultrapure distilled water for 30 min using a stir bar. 9. Rinse slides first in ethanol, and then in isopropanol. 10. Dry the slides using a stream of nitrogen gas, making sure that no residue is left on the slides. 11. Store the slides carefully in a clean container away from dust. The clean patterned slide can now be used for assembly of flow cells.

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Notes 1. Extreme care should be taken to avoid introduction of air bubbles into the sample chamber, especially after injection of the lipids. Air bubbles will destroy the lipid bilayer and lead to loss of signal. 2. All syringe attachments should be done by drop-to-drop connections. As a result, be careful never to detach both syringes at the same time. 3. At this step, care should be taken to inject ssDNA on the inlet side (port A, Fig. 2a), i.e., the side closer to the barriers, not the anchors. 4. It is important to have both syringes attached to the flow cell when injecting ssDNA. The second syringe will be required for making a drop-to-drop connection to the fluidic system when mounted on the microscope. 5. Before connecting the flow cell to fluidics, make sure that the lines have no air bubbles. This can be done by running buffer through the lines for 3–4 min before connecting the flow cell. 6. Be careful not to expose the RPA-ssDNA filaments to high laser intensities at this step, as this will cause loss of signal before recording due to photobleaching. 7. It is important to ensure that the sample chamber has been flushed with ATP-containing buffer before Rad51 is injected, as Rad51 activity requires ATP. 8. If the ssDNA is floppy, a wider ROI can be specified, such as 2or 3-pixel wide line using the plug-in “KymoReslideWide.” However, in all cases the ROI should be selected carefully, to make sure that only one ssDNA molecule is included, and adjacent ssDNA molecules are excluded. It is also recommended to select the ROI length over the central region of

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the ssDNA molecule, avoiding a few pixels adjacent to the barriers and anchors, as they may accumulate proteins nonspecifically over the course of the experiment. 9. Each intensity value in the list is an average over the entire length of the ssDNA molecule for that time point. 10. Be careful to mark the start and end points of the trajectory in same orientation across all kymographs, i.e., 30 first and 50 second. This method can also be used with unlabeled Srs2. In this case, the Srs2 translocation trajectory can be visualized by the binding of labeled RPA. 11. Be careful not to touch or damage the central area containing chrome barriers and anchors. Carefully work around the edges to remove the tape. 12. If the slide needs to be cleaned more thoroughly, the Hellmanex wash can be increased to 24–48 h. The solution should be changed to a fresh Hellmanex solution midway. References 1. San Filippo J, Sung P, Klein H (2008) Mechanism of eukaryotic homologous recombination. Annu Rev Biochem 77:229–257. https://doi.org/10.1146/annurev.biochem. 77.061306.125255 2. Kowalczykowski SC (2015) An overview of the molecular mechanisms of recombinational DNA repair. Cold Spring Harb Perspect Biol 7(11):a016410. https://doi.org/10.1101/ cshperspect.a016410 3. Kass EM, Moynahan ME, Jasin M (2016) When genome maintenance goes badly awry. Mol Cell 62(5):777–787. https://doi.org/10. 1016/j.molcel.2016.05.021 4. Prakash R, Zhang Y, Feng W, Jasin M (2015) Homologous recombination and human health: the roles of BRCA1, BRCA2, and associated proteins. Cold Spring Harb Perspect Biol 7(4):a016600. https://doi.org/10. 1101/cshperspect.a016600 5. Klein HL (2008) The consequences of Rad51 overexpression for normal and tumor cells. DNA Repair (Amst) 7(5):686–693. https:// doi.org/10.1016/j.dnarep.2007.12.008 6. Symington LS, Rothstein R, Lisby M (2014) Mechanisms and regulation of mitotic recombination in Saccharomyces cerevisiae. Genetics 198(3):795–835. https://doi.org/10.1534/ genetics.114.166140 7. Wright WD, Shah SS, Heyer WD (2018) Homologous recombination and the repair of DNA double-strand breaks. J Biol Chem 293

(27):10524–10535. https://doi.org/10. 1074/jbc.TM118.000372 8. Niu H, Klein HL (2017) Multifunctional roles of Saccharomyces cerevisiae Srs2 protein in replication, recombination and repair. FEMS Yeast Res 17(2):fow111. https://doi.org/10. 1093/femsyr/fow111 9. Krejci L, Van Komen S, Li Y, Villemain J, Reddy MS, Klein H, Ellenberger T, Sung P (2003) DNA helicase Srs2 disrupts the Rad51 presynaptic filament. Nature 423 (6937):305–309. https://doi.org/10.1038/ nature01577 10. Veaute X, Jeusset J, Soustelle C, Kowalczykowski SC, Le Cam E, Fabre F (2003) The Srs2 helicase prevents recombination by disrupting Rad51 nucleoprotein filaments. Nature 423(6937):309–312. https://doi. org/10.1038/nature01585 11. Liu J, Ede C, Wright WD, Gore SK, Jenkins SS, Freudenthal BD, Todd Washington M, Veaute X, Heyer WD (2017) Srs2 promotes synthesis-dependent strand annealing by disrupting DNA polymerase delta-extending D-loops. elife 6:e22195. https://doi.org/10. 7554/eLife.22195 12. Kaniecki K, De Tullio L, Gibb B, Kwon Y, Sung P, Greene EC (2017) Dissociation of Rad51 presynaptic complexes and heteroduplex DNA joints by tandem assemblies of Srs2. Cell Rep 21(11):3166–3177. https:// doi.org/10.1016/j.celrep.2017.11.047

ssDNA Curtains for Studying Rad51-ndash;ssDNA Filament Dynamics 13. Gorman J, Fazio T, Wang F, Wind S, Greene EC (2010) Nanofabricated racks of aligned and anchored DNA substrates for single-molecule imaging. Langmuir 26(2):1372–1379. https://doi.org/10.1021/la902443e 14. Greene EC, Wind S, Fazio T, Gorman J, Visnapuu ML (2010) DNA curtains for highthroughput single-molecule optical imaging. Methods Enzymol 472:293–315. https://doi. org/10.1016/S0076-6879(10)72006-1 15. Fazio T, Visnapuu ML, Wind S, Greene EC (2008) DNA curtains and nanoscale curtain rods: high-throughput tools for single molecule imaging. Langmuir 24 (18):10524–10531. https://doi.org/10. 1021/la801762h 16. Ma CJ, Steinfeld JB, Greene EC (2017) Singlestranded DNA curtains for studying homologous recombination. Methods Enzymol 582:193–219. https://doi.org/10.1016/bs. mie.2016.08.005 17. Graneli A, Yeykal CC, Prasad TK, Greene EC (2006) Organized arrays of individual DNA

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molecules tethered to supported lipid bilayers. Langmuir 22(1):292–299. https://doi.org/ 10.1021/la051944a 18. De Tullio L, Kaniecki K, Greene EC (2018) Single-Stranded DNA Curtains for Studying the Srs2 Helicase Using Total Internal Reflection Fluorescence Microscopy. Methods Enzymol 600:407–437. https://doi.org/10.1016/ bs.mie.2017.12.004 19. Gibb B, Ye LF, Kwon Y, Niu H, Sung P, Greene EC (2014) Protein dynamics during presynaptic-complex assembly on individual single-stranded DNA molecules. Nat Struct Mol Biol 21(10):893–900. https://doi.org/ 10.1038/nsmb.2886 20. De Tullio L, Kaniecki K, Kwon Y, Crickard JB, Sung P, Greene EC (2017) Yeast Srs2 helicase promotes redistribution of single-stranded DNA-bound RPA and Rad52 in homologous recombination regulation. Cell Rep 21 (3):570–577. https://doi.org/10.1016/j.cel rep.2017.09.073

Chapter 12 Following Trypanosoma cruzi RPA-DNA Interaction Using Fluorescent In Situ Hybridization Coupled with Immunofluorescence (FISH/IF) Raphael S. Pavani and Maria Carolina Elias Abstract Fluorescent in situ hybridization coupled with immunofluorescence (FISH/IF) is an assay that has been widely used to study DNA-protein interactions. The technique is based on the use of a fluorescent nucleic acid probe and fluorescent antibodies to reveal the localization of a DNA sequence and a specific protein in the cell. The interaction can be inferred by the quantification of the co-localization between the protein and the DNA. Here, we describe a detailed FISH/IF methodology that our group used to study RPA-telomere interaction in the pathogenic protozoa parasite Trypanosoma cruzi. Key words Fluorescent in situ hybridization, FISH, Immunofluorescence, IF, Trypanosoma, RPA, ssDNA

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Introduction Immunofluorescence (IF) is a technique used to study the location of specific molecules in the cells. The principle of this method relies on the use of fluorescent-labeled antibodies that are able to recognize specific target antigens [1]. Single-stranded DNA-binding proteins have been studied in different organisms using this technique due to its ability to accumulate in the exposed singlestranded DNA (ssDNA), forming discrete nuclear foci, especially after DNA damage or replicative stress that can generate long tracts of this type of DNA molecule [2–8]. Different from immunofluorescence, the fluorescent in situ hybridization (FISH) assay consists of the hybridization of a fluorescent nucleic acid probe to its complementary sequence in order to locate specific DNA or RNA sequences within a cell [9, 10]. This technique has a wide range of applications, from basic biology research to diagnosis of chromosomal DNA aberrations/abnormalities or infectious diseases [11].

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_12, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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To study DNA-protein interactions using microscopy-based approaches, scientists coupled these two techniques in one assay, named FISH/IF. The co-localization of the protein and the DNA signals can be quantified and used to infer the DNA-protein interactions. Recently, we used FISH/IF to study the interaction of RPA (the major ssDNA-binding protein of eukaryotes) with the telomeric DNA in the protozoa parasite Trypanosoma cruzi [12]. In this chapter, we describe in detail how to perform the FISH/IF assay in this parasite, making notes and highlighting important and crucial steps to have a clean and successful FISH/IF experiment.

2

Materials 1. T. cruzi epimastigote forms in exponential growth cultivated in LIT medium [13]. 2. 0.1% Poly-L-lysine. 3. Microscope glass slides. 4. Coverslips. 5. 50 mL Falcon tubes. 6. Fixative buffer: 4% Paraformaldehyde in PBS. This buffer can be stored at
20  C for a few months. 7. PBS: 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, pH 7.4. 8. Permeabilization buffer: PBS/0.1% Triton X-100. Filter and store at 4  C. 9. Blocking buffer: PBS/1% BSA. 10. Primary antibody solution: Anti-RPA-1 antibody [3, 14] diluted 1:300 in blocking buffer. 11. Secondary antibody solution: Alexa fluor 555 anti-rabbit antibody diluted 1:500 in blocking buffer. 12. 70% Ethanol, 80% ethanol, and 90% ethanol, made with deionized water. Store at
20  C. 13. Telomere PNA Kit/FITC. 14. 1 Wash buffer—diluted 20. 15. Frame-Seal in situ PCR and hybridization slide chambers, 9  9 mm (see Fig. 1). 16. Hybridization buffer: 20 mM Tris–HCl pH 7.0, 70% formaldehyde, and 1% BSA. 17. Mounting medium with DAPI.

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Fig. 1 Mounting the slide with the Frame-Seal. (a) Microscope glass slide and the components of the FrameSeal. (b–d) Glue the blue square on the glass slide; the experiment will be performed in the blue square region. (e, f) Fluorescent probe being applied on the slide (the probe color can vary according to the fluorophore or can also be colorless; the red color here is just to facilitate the demonstration). (g) Seal the blue square with the provided adhesive

3

Methods All procedures are performed at room temperature unless otherwise specified.

3.1

Slide Preparation

1. Before starting the experiment, soak the slides in 70% ethanol for 1 h to remove impurities and allow it to dry at room temperature. Glue the blue square from the frame seal in the slides, to create the region where the sample will be applied later (Fig. 1a–d). Apply 5 μL of 0.1% poly-L-lysine and spread it with a pipette tip. Allow it to dry completely (see Note 1). 2. Harvest approximately 1  107 exponentially growing T. cruzi epimastigotes, by centrifuging culture at 1000  g for 5 min. Carefully wash with PBS once and resuspend cell pellet in 1 mL PBS.

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3. Apply 80 μL of the resuspended parasite cells inside the blue square region of the slide and incubate for 5 min. This step will allow the parasites to adhere to the slide. 3.2 Fixation and Permeabilization

1. Remove PBS and apply 80 μL of the fixative buffer. Incubate for 10 min. 2. Remove the fixative buffer and wash the slides three times with PBS. 3. Apply the permeabilization buffer, incubate for 10 min, and wash three times with PBS.

3.3 Immunofluorescence

1. Apply the blocking solution for at least 30 min. 2. Remove blocking solution and apply the primary antibody solution. Incubate for 1 h (see Note 2). 3. Wash three times with PBS and apply the secondary antibody solution for 1 h (see Notes 3 and 4). 4. Perform three washes with PBS and incubate with cold fixative solution for 20 min. Wash three times with PBS (see Note 5).

3.4 Fluorescence In Situ Hybridization

1. To start the FISH, the cells need to be dehydrated. Submerge the slide completely in cold 70% ethanol in a 50 mL Falcon for 2 min. 2. Remove the slide with a pair of tweezers and transfer to another 50 mL Falcon tube containing cold 80% ethanol for 2 min. 3. Remove the slide with a pair of tweezers and transfer to another 50 mL Falcon tube containing cold 90% ethanol for 2 min. 4. Remove the slide with a pair of tweezers and let it dry at room temperature. 5. Apply 8–10 μL of the telomeric DNA probe (Fig. 1e) and close the blue square with the leaky-proof adhesive (Fig. 1f–i) (see Note 6). 6. Incubate the slide at 95  C for 5 min, transfer it to 37  C, and incubate overnight (see Notes 7 and 8). 7. In the next day, remove the adhesive and incubate the slide in the washing solution for 5 min at 60  C. 8. Dehydrate the slide with cold ethanol by repeating steps 1–4. Let it dry completely. 9. Carefully remove the blue square from the slide and apply 8–10 μL of the antifade mounting media containing DAPI (see Notes 9 and 10).

3.5

Analyze the Slide

1. Analyze the slide in the microscope and take images corresponding to your probe and antibody signals (Fig. 2a) (see Note 11).

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Fig. 2 FISH/IF showing co-localization of RPA-1 and telomeres in T. cruzi. (a) FISH/IF assays were done in T. cruzi epimastigote cells. RPA-1 is labeled in red, telomeres are labeled in green (telomere PNA probe/FITC), and DAPI is blue. The flagella (F) are labeled in green and were identified to determine the phase of the cell cycle (G1/S is one nucleus/one flagellum; G2/M is one nucleus/two flagella; and cytokinesis is two nuclei/two flagella). k kinetoplast DNA, N nucleus. (b) Quantification of cells containing RPA-1-telomere co-localization (we count positive more than one yellow dot). (c) Graph showing the amount of telomere co-localizing with RPA per cell quantified using the Image J JaCoP plug-in. The percentage was obtained by Mander’s overlap coefficient (M1). This figure is reproduced from ref. 12 with permission from John Wiley and Sons—license number 4771261319636

2. To analyze the pictures, we installed the JACoP (Just Another Co-localization Plugin) plug-in on Image J (see Note 12) [15]. 3. Open the images on Image J, and change them to 16-bit (Image ! Type ! 16-bit). 4. Open the JaCoP plug-in and select your images as Image A and Image B. Select M1 and M2 coefficients (Mander’s coefficient) and press analyze. The result will show you the fraction of A overlapping with B and vice versa, ranging from 0 to 1 (see Note 13).

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Notes 1. To dry the slides in a quicker way, you can allow them to dry inside the laminar flow hood, which should take about 5 min. 2. The best concentration and time of incubation for your primary antibody can vary between IF alone and FISH/IF and need to be empirically tested. 3. After the secondary antibody incubation, avoid direct light exposure on the slides until the end of the experiment (fluorescent antibodies are light sensitive). You can use foil paper or dark boxes to protect it from light. 4. Examine the excitation and emission spectra of the selected fluorophores for your secondary antibody and your FISH probe to avoid overlap between the signals. 5. The post-IF fixative step is fundamental to maintain the antibody-antigen interaction since the next steps involve dehydration with ethanol. 6. We used telomere PNA probe/FITC in hybridization solution from the telomere PNA Kit/FITC from Dako. However, there are available protocols to build your own probe [16]; make note that changes in the protocol above may be needed. 7. The best way to incubate the slides at 95  C is using slidecompatible thermoblocks. In case you do not have this equipment, use a Falcon tube filled with water at 95  C. The Falcon tube with your slide can be immersed in a Becker containing 95  C water in order to maintain the high temperature. 8. After 95  C incubation, transfer the slides quickly to the 37  C incubator. It is important to avoid fast decrease in the temperature or allow temperatures below 37  C. 9. This protocol can be used to perform IF without doing FISH. Just stop in step 3 (Subheading 3.3) and mount the slide as described in step 9 (Subheading 3.4). In this case, the second incubation with the fixative solution is not necessary. 10. We use the antifade reagent Vecta-Shield containing DAPI. If your antifade reagent does not contain DAPI, prepare a solution of 1 μg/mL of DAPI in PBS and incubate for 5 min before mounting the slides. 11. Analyzing the result using a confocal microscope or a microscope able to create Z-stacks is the best way to verify and quantify co-localization. 12. We quantified our experiments using the software ImageJ, which can be downloaded for free. 13. The JaCoP plug-in allows you to also choose other methods to quantify co-localization, such as Pearson’s correlation

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coefficient. More information about the methods and analysis can be found at https://imagej.nih.gov/ij/plugins/track/ jacop2.html.

Acknowledgments The authors want to thank Simone Calderano for helping with the photos presented in Fig. 1. M.C.E. is a fellow from the National Council for Scientific and Technological Development (CNPq: 306199/2018-1). RSP received a Fundac¸˜ao de Amparo a` Pesquisa do Estado de Sa˜o Paulo fellowship (FAPESP 2014/02978-0). References 1. Odell ID, Cook D (2013) Immunofluorescence techniques. J Invest Dermatol 133(1):e4 2. Kobayashi T, Tada S, Tsuyama T, Murofushi H, Seki M, Enomoto T (2002) Focus-formation of replication protein A, activation of checkpoint system and DNA repair synthesis induced by DNA double-strand breaks in Xenopus egg extract. J Cell Sci 115:3159–3169 3. Pavani RS, da Silva MS, Fernandes CAH, Morini FS, Araujo CB, Fontes MR de M, Sant’Anna OA, Machado CR, Cano MI, Fragoso SP, Elias MC (2016) Replication protein A presents canonical functions and is also involved in the differentiation capacity of Trypanosoma cruzi. PLoS Negl Trop Dis 10: e0005181 4. Croft LV, Ashton NW, Paquet N, Bolderson E, O’Byrne KJ, Richard DJ (2017) hSSB1 associates with and promotes stability of the BLM helicase. BMC Mol Biol 18:1–10 5. Liu J-S, Kuo S-R, Melendy T (2006) DNA damage-induced RPA focalization is independent of γ-H2AX and RPA hyperphosphorylation. J Cell Biochem 99:1452–1462 6. Chen H, Lisby M, Symington LS (2013) RPA coordinates DNA end resection and prevents formation of DNA hairpins. Mol Cell 50:589–600 7. Lisby M, Rothstein R (2009) Choreography of recombination proteins during the DNA damage response. DNA Repair (Amst) 8:1068–1076 8. Pavani RS, Lima LP, Lima AA, Fernandes CAH, Fragoso SP, Calderano SG, Elias MC (2020) Nuclear export of replication protein A in the nonreplicative infective forms of Trypanosoma cruzi. FEBS Lett 594:1596–1607

9. Huber D, Voith von Voithenberg L, Kaigala GV (2018) Fluorescence in situ hybridization (FISH): history, limitations and what to expect from micro-scale FISH? Micro Nano Eng 1:15–24 10. Volpi EV, Bridger JM (2008) FISH glossary: an overview of the fluorescence in situ hybridization technique. BioTechniques 45:385–409 11. Cui C, Shu W, Li P (2016) Fluorescence in situ hybridization: cell-based genetic diagnostic and research applications. Front Cell Dev Biol 4:89 12. Pavani RS, Vitarelli MO, Fernandes CAH, Mattioli FF, Morone M, Menezes MC, Fontes MRM, Cano MIN, Elias MC (2018) Replication protein A-1 has a preference for the telomeric G-rich sequence in Trypanosoma cruzi. J Eukaryot Microbiol 65:345–356 13. Contreras VT, Araujo-Jorge TC, Bonaldo MC, Thomaz N, Barbosa HS, Meirelles M de N, Goldenberg S (1988) Biological aspects of the Dm 28c clone of Trypanosoma cruzi after metacyclogenesis in chemically defined media. Memo´rias do Inst. Oswaldo Cruz 83:123–133 14. Damasceno JD, Nunes VS, Tosi LRO (2013) LmHus1 is required for the DNA damage response in Leishmania major and forms a complex with an unusual Rad9 homologue. Mol Microbiol 90:1074–1087 15. Bolte S, Cordelie`res FP (2006) A guided tour into subcellular colocalization analysis in light microscopy. J Microsc 224:213–232 16. Elias MCQB, Faria M, Mortara RA, Motta MCM, de Souza W, Thiry M, Schenkman S (2002) Chromosome localization changes in the Trypanosoma cruzi nucleus. Eukaryot Cell 1:944–953

Chapter 13 Quantifying the Affinity of Trypanosoma cruzi RPA-1 to the Single-Stranded DNA Overhang of the Telomere Using Surface Plasmon Resonance Marcela de Oliveira Vitarelli and Maria Carolina Elias Abstract Surface plasmon resonance (SPR) biosensors provide real-time binding affinity measurements between a pair of biomolecules, characterizing its interaction dynamics. An example of Trypanosoma cruzi’s RPA-1 and a single-stranded DNA telomere sequence is presented with detailed guidelines and fundamentals for SPR technology. Key words Biosensor, Surface plasmon resonance, Protein-DNA interactions, RPA, Single-stranded DNA, Kinetics, Trypanosoma cruzi

1

Introduction Biosensor surface plasmon resonance (SPR) is a method employed to evaluate the interactions between labeled and free biomolecules in real time. Although it was first designed to analyze proteinantibody and protein-protein interactions, it is now greatly employed in more complex and challenging interactions using different biomolecule combinations such as protein-small drug targets and protein-nucleic acids [1–4]. SPR is a phenomenon based on the production of plasmons in a thin conducting metal film, located on the interface of media with different refractive indexes. The physical basis of this phenomenon started to be studied in the early 1900s, but it was in 1983 that Liedberg presented the first application of SPR for biosensing, using IgG antibodies [5, 6]. The majority of SPR sensor chips are composed of a gold layer deposited on a glass slide. To avoid the direct interaction of the ligand to the metal, the gold layer of the sensor chips can be covered with alkanethiol. This hydrophobic monolayer can be directly used in the sensor chip or can be covered with different

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matrices, such as carboxymethyl dextran, which favors the covalent interaction of ligands to the surface. Another example is the streptavidin sensor chip, covered with carboxymethyl dextran and streptavidin, favoring the immobilization of biotinylated ligands [7, 8]. In a given wavelength and angle (SPR angle), incident light in the glass slide excites the delocalized electrons on the metal surface in contact with the solution (Fig. 1). The resonantly excited electrons absorb some of the energy, thus diminishing the intensity of the reflected light. The smallest change in the external media, such as the binding of an analyte in a ligand immobilized in the sensor chip, can entail a change in its refractive index which leads to a change in the SPR angle. This change can be detected by a diode detector and depicted in the form of a sensorgram with the shift in the incident light angle measured in resonance units (RU) versus time (1 RU ¼ 0.0001 shift) (Fig. 2) [8–11]. SPR has great advantages when compared to other quantitative assays. Since there is no need for the incident light to pass through the analyte sample and due to the high sensitivity of the method, it requires very low concentrations of analyte bound to the surface. This also eliminates the need of additional probes to the sample in order to improve the response, excluding the labeling interference in the system [10]. The real-time response of SPR to a change in the refractive index allows for a quantitative analysis of complex formation [6, 12–15]. The interaction of a given analyte (A) with a ligand immobilized on the sensor chip (L) is given by ka L þ A Ð LA kd where ka is the association rate constant and kd the dissociation rate constant. The association and dissociation constants can be given by applying a global fit routine into a sensorgram of a kinetic experiment in one’s system of choice, allowing for the calculation of the equilibrium binding affinity (KA): KA ¼

ka 1 ¼ kd K D

where KD is the equilibrium dissociation constant. However, to obtain accurate measurements of the constants, a proper experimental design is needed. First, the ligand must be properly immobilized in the metal surface of the sensor chip. Then, a regeneration buffer must be set: it is necessary to rupture ligandanalyte interactions without removing the bound ligand in the chip surface; the regeneration scouting tests the best buffer for this

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Fig. 1 Schematic representation of a SPR biosensor. A LED plane polarized light focused in the glass slide in a condition of total internal reflection, forming a SPR angle. In these conditions, the delocalized electrons in the gold layer in contact with a flowing solution are excited and the intensity of the reflected light is reduced. The ligation of an analyte with a ligand immobilized in the sensor chip changes the refractive index of the flowing solution, leading to a shift in SPR angle. This shift is detected by a diode detector and reported in response units (RU). (Image created with BioRender.com) Figure adapted from [6, 12]

Fig. 2 Schematic representation of a sensorgram. A sensorgram is a graphic response generated by a SPR biosensor, given in RU versus time. It reports binding, dissociation, and regeneration events that are happening in real time in a given flow cell. In a kinetic assay, those measurements allow for the calculation of association (KA) and dissociation constants (KD) of a binding event. (Image created with BioRender.com) Figure adapted from [12, 16]

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intent. The next step is a binding event, to test binding conditions and minimize mass transport effects [16, 17] to finally proceed to the kinetics and affinity analysis. Here, we present a detailed protocol for protein-singlestranded DNA (ssDNA) interaction using the Trypanosoma cruzi RPA-1 protein and a telomeric DNA sequence. Replication protein A (RPA) is the major ssDNA-binding protein in eukaryotes and is composed of three subunits: RPA-1, RPA-2, and RPA-3, which contain specific oligonucleotide-binding folds (OBFs), responsible for wrapping around the ssDNA through its β-sheet structure. This conserved heterotrimeric protein is involved in different cell responses, such as DNA replication, recombination, repair, checkpoint, and telomere maintenance [18, 19]. T. cruzi’s RPA (TcRPA) was recently characterized by Pavani et al. (2016), presenting the three subunits of the complex: TcRPA1, TcRPA-2, and an annotated protein which is believed to be TcRPA-3. TcRPA also presents OBFs; however, it seems to be larger and possesses more flexible linkers than the ones already described for model eukaryotes. TcRPA-1 is the subunit of the complex responsible for interacting with the ssDNA and it was shown to participate in DNA replication and repair. Even though TcRPA-2 does not interact directly with the DNA, its depletion impacted the progression of cell division. Together with the change in localization of TcRPA during the different life stages of T. cruzi, this indicates that RPA also plays a role in the parasite’s metacyclogenesis; once the exportation of TcRPA from the nucleus has only been described in this organism’s non-replicative form [19, 20]. The role of TcRPA in the telomeres was not yet fully characterized, but it was shown that the heterotrimeric complex colocalizes with telomeric regions and it has an affinity for a 24 bp G-rich telomere sequence, suggesting its involvement at telomeric sites [21].

2

Materials

2.1 General Materials and Buffers

1. SPR instrument and software (see Note 1). 2. Thermoblock. 3. Plastic vials and caps for use with SPR systems. 4. Buffer 1: 0.01 M HEPES pH 7.4, 0.15 M NaCl. 5. Buffer 2: 0.01 M HEPES pH 7.4, 0.15 M NaCl, 3 mM EDTA, 0.005% v/v Surfactant P20. 6. Buffer 3: 0.1 M HEPES pH 7.4, 1.5 M NaCl, 0.5% v/v Surfactant P20.

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2.2 Streptavidin Immobilization

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1. Carboxymethyl dextran sensor chip. 2. Normalizing solution: 70% (w/w) Glycerol. 3. 75 mg/mL 1-Ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC). 4. 11.5 mg/mL N-Hydroxysuccinimide (NHS). 5. 1 M Ethanolamine hydrochloride-NaOH pH 8.5. 6. 200 μg/mL Streptavidin in 10 mM acetate buffer, pH 4.5. 7. 50 mM NaOH in Buffer 2.

2.3 DNA Binding to Streptavidin

1. 50 Biotinylated unrelated oligonucleotide: CACTCCCGTGAAAACTC (see Note 2).

GCTAGCT

2. 50 Biotinylated ssDNATel24: TTAGGGTTAGGGTTAGGGT TAGGG (see Note 2). 3. 1 M NaCl in 50 mM NaOH. 4. Isopropanol. 2.4 Regeneration Scouting, Protein-DNA Binding, and Kinetics

1. 100 μg/mL BSA, 5 mM MgCl2 in Buffer 3 (see Note 3). 2. Purified recombinant TcRPA-1 (rTcRPA-1) [19]. 3. 10% SDS. 4. 1 M NaCl in 100 μg/mL BSA, 5 mM MgCl2 in Buffer 3.

3

Methods

3.1 Streptavidin Immobilization

When using a carboxymethyl dextran sensor chip, it is necessary to first activate the carboxymethyl dextran surface and then immobilize the streptavidin using an amino coupling method (see Note 4). 1. Remove the carboxymethyl dextran sensor chip from the refrigerator and leave it at room temperature for at least 30 min. 2. Wash system three times with Buffer 1. 3. Dock the carboxymethyl dextran sensor chip. 4. Normalize sensor chip signal (see Notes 5 and 6). 5. Wash the system three times with Buffer 1. 6. Prepare streptavidin, EDC, NHS, and ethanolamine solutions and place them separately into capped vials. 7. Wash system one time with Buffer 2. 8. Immobilize streptavidin (200 μg/mL in 10 mM acetate buffer, pH 4.5) in all flow cells from the carboxymethyl dextran sensor chip, at 25  C, to reach a target level of 2700 RU (see Note 7).

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9. To remove excess of non-bound streptavidin, wash all flow cells with 50 mM NaOH in Buffer 2 at a flow rate of 2 μL/min for at least five times, or until there is a significant drop in RU. 10. Wash system one time with Buffer 2 and leave chip docked for 30 min or more until stabilization of RU levels (see Note 8). 3.2 DNA Binding to Streptavidin

In this step, the remaining unbound streptavidin is washed away and the biotinylated DNA is bound to the immobilized streptavidin. 1. Wash system one time with Buffer 2. 2. To remove the remaining unbound streptavidin, wash all flow cells with 1 M NaCl in 50 mM NaOH at a flow rate of 10 μL/ min, at 25  C, with 60 s of contact time (see Note 9). 3. If necessary, perform an extra wash with 50% isopropanol diluted in 1 M NaCl in 50 mM NaOH at a flow rate of 10 μL/min, at 25  C, with 60 s of contact time. 4. Wash system one time with Buffer 2. 5. Dilute the 5’ biotinylated unrelated oligonucleotide to 0.1 μg/ mL in Buffer 2, heat the samples at 95  C for 5 min immediately prior to binding, and place them separately into capped vials (see Note 10). 6. Select the flow cell for binding of the 5’ biotinylated unrelated oligonucleotide (see Note 11). 7. Wash selected flow cell one time with Buffer 2. 8. Run DNA binding cycle with the following parameters: 90 s of contact time, 600 s of dissociation time, and flow rate of 10 μL/min. 9. To remove the excess unbound DNA, wash selected flow cell with Buffer 2 with the following parameters: 45 s of contact time, 60 s of dissociation time, and flow rate of 10 μL/min, at 25  C (see Note 12). 10. Dilute the 50 biotinylated ssDNATel24 oligonucleotide to 0.1 μg/mL in Buffer 2, heat the samples at 95  C for 5 min immediately prior to binding, and place them separately into capped vials (see Note 10). 11. Select a different flow cell for binding of the 5’ biotinylated ssDNATel24 oligonucleotide (see Note 11). 12. Wash selected flow cell one time with Buffer 2. 13. Run DNA binding cycle with the following parameters: 90 s of contact time, 600 s of dissociation time, and flow rate of 10 μL/min.

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14. To remove the excess unbound DNA, wash selected flow cell with Buffer 2 with the following parameters: 45 s of contact time, 60 s of dissociation time, and flow rate of 10 μL/min, at 25  C (see Note 12). 15. Wash all flow cells one time with Buffer 2. 3.3 Regeneration Scouting

In this step, different regeneration buffers are tested in order to find one that removes protein-DNA interactions without removing streptavidin-DNA interactions from the surface of the sensor chip. 1. Prepare three solutions to be used as regeneration buffers: 0.05% SDS, 0.1% SDS, and 1 M NaCl. Dilute rTcRPA-1 to 10 nM in 100 μg/mL BSA and 5 mM MgCl2 in Buffer 3 and place all solutions separately into capped vials (see Notes 3 and 13). 2. Select the flow path used for one of the DNA-bound flow cells minus the control one (2-1 for example) (see Note 14). 3. Run five cycles of the regeneration scouting step for each of the three regeneration buffers with the following parameters: 30 s of contact time, 300 s of stabilization period, and flow rate of 100 μL/min. 4. Evaluate the best regeneration buffer (see Note 15). 5. Repeat for different protein concentrations if needed (see Note 16).

3.4 Protein-DNA Binding

After setting the regeneration buffer, a binding assay is performed prior to kinetics to set all binding conditions (see Note 17). 1. Prepare regeneration buffer of choice, dilute rTcRPA-1 to 50 nM in 100 μg/mL BSA and 5 mM MgCl2 in Buffer 3, and place into capped vials (see Notes 18–20). 2. Select the flow paths used for each of the DNA-bound flow cells minus the respective controls (2-1 and 4-3 for example). 3. Wash all flow cells one time with 100 μg/mL BSA and 5 mM MgCl2 in Buffer 3 (see Note 21). 4. Run one binding cycle with the following parameters: 45 s of contact time, 60 s of dissociation time, and flow rate of 50 μL/ min. 5. The binding cycle must be followed by a regeneration cycle with the best regeneration buffer, in this case 0.05% SDS, with the following parameters: 30 s of contact time, 300 s of stabilization period, and flow rate of 100 μL/min.

3.5

Kinetics

1. Prepare 0.05% SDS regeneration buffer; dilute rTcRPA-1 to 0, 12.5, 25, 50, 100, 100, and 200 nM in 100 μg/mL BSA and 5 mM MgCl2 in Buffer 3; and place into capped vials (see Notes 19 and 22).

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2. Select the flow paths used for each of the DNA-bound flow cells minus the respective controls (2-1 and 4-3, for example). 3. Wash all flow cells one time with 100 μg/mL BSA and 5 mM MgCl2 in Buffer 3. 4. Run two kinetic cycles with the parameters determined in the protein-DNA binding step. For binding: 45 s of contact time, 60 s of dissociation time, and flow rate of 50 μL/min. For regeneration: 30 s of contact time, 300 s of stabilization period, and flow rate of 100 μL/min. 5. With the kinetics sensorgram, adjust the Y-axis to the baseline, perform a 1:1 fitness curve, and evaluate the results (Fig. 3) (see Notes 23–27).

a

RU 150 100

Response

50 0 -50 -100 -150 -60

-40

-20

0

40

20

60

80

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Time

120 s

b RU 140 120 100

Response

80 60 40 20 0 -20 -60

-40

-20

0

40

20 Time

60

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120 s

Fig. 3 Kinetics software screenshots, showing two kinetic assays fitted in a 1:1 binding event with rTcRPA-1 and the oligonucleotides for (a) unrelated sequence (ka ¼ 7.354  104 (1/Ms), kd ¼ 0.4487 (1/s), KD ¼ 6.101  106 (M)) and (b) ssTelDNA24 (ka ¼ 3.931  105 (1/Ms), kd ¼ 0.005182 (1/s), KD ¼ 1.318108 (M)) (see Note 26)

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Notes 1. This protocol was designed using the Biacore T100 instrument, Biacore T100 Control Software, and Biacore T100 Evaluation Software, available from General Electric Healthcare company. Some adjustments might have to be made when using other SPR systems and software. 2. If desired, it is possible to ligate the biotin to an oligonucleotide. However, it is more practical to order the biotinylated oligos. 3. This buffer was used according to Vaidyanathan et al. [22]. 4. Streptavidin-coated sensor chips are also available and can be used. In this case, start at Subheading 3.2. 5. If sensor chip normalization returns with an error message, repeat the procedure. 6. Normalize the signal every time when using a new sensor chip. 7. If the RU of streptavidin immobilization almost reaches the targeted level but the software does not recognize it as a ligation event, lower the target level to the one observed in the reference line and try again. 8. Consider only the binding RU after stabilization and not the binding peaks. 9. The wash with 1 M NaCl in 50 mM NaOH is performed to remove streptavidin excess. This step is recommended for the streptavidin sensor chip. However, we also recommend it when using the carboxymethyl dextran sensor chip immobilized with streptavidin. 10. If the DNA forms secondary structures (e.g., G-quadruplex), heat samples at 95  C for 5 min immediately prior to binding. If not, the heating step is not necessary. 11. It is very important to always use one flow cell as control, without immobilized DNA, for each binding event. If you have, as in this case, two different DNAs, you will need two flow cells as controls. 12. For the regeneration step we used Buffer 2 to remove only the excess of non-bound DNA. If regeneration of the flow cell is needed, use 1 M NaCl in Buffer 2 with the same settings. 13. Always use the running buffer as the one in the diluted protein. 14. For testing purposes, there is no need to perform regeneration scouting in all the flow cells. 15. The best buffer will be the one with a regeneration response closer to the baseline.

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16. The regeneration scouting is also a good way to test binding concentrations and make improvements in the protein-DNA binding step if needed. You should be able to see the binding event and the dissociation in the regeneration scouting sensorgram. 17. If the sensor chip has already been through extensive tests, it is advisable to get a fresh one to perform the protein-DNA binding step. Be sure to repeat all the protocol for DNA ligation prior to protein-DNA interaction when using a fresh sensor chip. 18. Protein concentrations might change from the ones in the example depending on its interaction with the DNA sequence. 19. It is imperative that the protein is highly pure and its concentration is accurate. Quantification by more than one method is strongly recommended (e.g., BSA and nanodrop). 20. If necessary, different protein concentrations can be tested in the same run. 21. It is possible that Buffer 2 itself gives an RU response. It is strongly recommended to use the 100 μg/mL BSA and 5 mM MgCl2 in Buffer 3 to block empty streptavidin sites. 22. It is important to reproduce at least one point of the sensorgram in every kinetic assay in order to verify the reproducibility of the data. 23. To compare more than one flow cell, it is better to adjust the Y-axis by dividing it to the baseline outcome. 24. Check the quality control tab to confirm if all the results passed the quality assessment. If any has failed, make the necessary adjustments to repeat kinetics and fitting assays. 25. Be sure to check statistical data provided in the Reports tab: The U-value indicates if the calculated constant rates (ka and kd) and maximum response (Rmax) are correlated or not. This means that if the parameters are correlated (U-value >25) the absolute values for two or more parameters cannot be precisely measured, but their magnitude can be estimated in a combined function of the parameters. If U-value is 15, parameters are not correlated and can be individually measured. If U-value falls in the interval between 15 and 25, no statistical correlation can be established among the parameters. The Chi2 measures the variation among experimental data and the fitted curve by calculating the average squared residual per data point: Chi2 ¼

n X ðr f  r x Þ2 d 1

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where rf is the fitted value at a given point, rx is the experimental value at the same point, and d is the number of data points. For a Chi2 to be statistically significant, its value should be approximate to the mean square of the short-term noise level. The short-term noise level is given, in the sensorgram, by the difference among fitted and experimental data in a given point. Since it depends on the signal level, it can vary between samples and a cutoff value cannot be established. In the Parameters tab, the T-value should also be considered. It measures the significance of the parameters based on the standard error (SE): T  value ¼

Parameter value SE

A high T-value indicates a low SE. A T-value 10 is normally considered to be significant [23]. It is also advisable to investigate if the data has any biological meaning. 26. We recommend exporting the data to a different plotting software in order to eliminate the spiked curve underneath the fitted model. 27. Although it is advisable to have the kinetic curve in a proper “wave” format, this can be influenced by the interaction among ligand and analyte, and the formation of secondary structures by the immobilized DNA.

Acknowledgments This work was supported by Fundac¸˜ao de Amparo a` Pesquisa do Estado de Sa˜o Paulo (FAPESP 2013/07467-1; 2016/50050-2; 2017/07693-2) and Conselho Nacional de Pesquisa e Desenvolvimento (CNPq 306199/2018-1; 870219/1997-9). References 1. Quinn JG, O’Neill S, Doyle A, McAtamney C, Diamond D, MacCraith BD, O’Kennedy R (2000) Development and application of surface plasmon resonance-based biosensors for the detection of cell-ligand interactions. Anal Biochem 281(2):135–143. https://doi.org/10. 1006/abio.2000.4564 2. Davis TM, Wilson WD (2000) Determination of the refractive index increments of small molecules for correction of surface plasmon resonance data. Anal Biochem 284 (2):348–353. https://doi.org/10.1006/abio. 2000.4726 3. Papalia GA, Giannetti AM, Arora N, Myszka DG (2008) Thermodynamic characterization

of pyrazole and azaindole derivatives binding to p38 mitogen-activated protein kinase using Biacore T100 technology and van’t Hoff analysis. Anal Biochem 383(2):255–264. https:// doi.org/10.1016/j.ab.2008.08.010 4. Capelli D, Parravicini C, Pochetti G, Montanari R, Temporini C, Rabuffetti M, Maria Trincavelli L, Daniele S, Fumagalli M, Saporiti S, Bonfanti E, Abbracchio MP, Eberini I, Ceruti S, Calleri E, Capaldi S (2020) Surface plasmon resonance as a tool for ligand binding investigation of engineered GPR17 receptor, a G protein coupled receptor involved in myelination. Front Chem 7:910. https://doi.org/10.3389/fchem.2019.00910

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5. Liedberg B, Nylander C, Lunstro¨m I (1983) Surface plasmon resonance for gas detection and biosensing. Sensors Actuators 4:299–304. https://doi.org/10.1016/0250-6874(83) 85036-7 6. Liedberg B, Nylander C, Lunstro¨m I (1995) Biosensing with surface plasmon resonance— how it all started. Biosens Bioelectron 10(8): i–ix. https://doi.org/10.1016/0956-5663( 95)96965-2 7. Jason-Moller L, Murphy M, Bruno J (2006) Overview of Biacore systems and their applications. Curr Protoc Protein Sci 19:19.13.1–19.13.14. https://doi.org/10. 1002/0471140864.ps1913s45 8. Biacore® Sensor Surface Handbook (2003) Version AA. Biacore AB 9. Cullen DC, Brown RG, Lowe CR (1987) Detection of immuno-complex formation via surface plasmon resonance on gold-coated diffraction gratings. Bios 3(4):211–225. https:// doi.org/10.1016/0265-928x(87)85002-2 10. Liu Y, Wilson WD (2010) Quantitative analysis of small molecule-nucleic acid interactions with a biosensor surface and surface plasmon resonance detection. Methods Mol Biol 613:1–23. https://doi.org/10.1007/978-1-60327-4180_1 11. Wang S, Poon GMK, Wilson WD (2015) Quantitative investigation of protein-nucleic acid interactions by biosensor surface plasmon resonance. Methods Mol Biol 1334:313–332. https://doi.org/10.1007/978-1-4939-28774_20 12. Nguyen B, Tanious FA, Wilson WD (2007) Biosensor-surface plasmon resonance: quantitative analysis of small molecule–nucleic acid interactions. Methods 42(2):150–161. https://doi.org/10.1016/j.ymeth.2006.09. 009 13. Myszka DG (2000) Kinetic, equilibrium, and thermodynamic analysis of macromolecular interactions with BIACORE. Methods Enzymol 323:325–340. https://doi.org/10.1016/ s0076-6879(00)23372-7 14. Karlsson R (1999) Affinity analysis of nonsteady-state data obtained under mass transport limited conditions using BIAcore technology. J Molecul Recogn 12(5):285–292. https://doi.org/10.1002/(sici)1099-1352( 199909/10)12:53.0. co;2-y 15. Morton TA, Myszka DG (1998) Kinetic analysis of macromolecular interactions using

surface plasmon resonance biosensors. Methods Enzymol 295:268–294. https://doi.org/ 10.1016/s0076-6879(98)95044-3 16. Lin LP, Huang LS, Lin CW, Lee CK, Chen JL, Hsu SM, Lin S (2005) Determination of binding constant of DNA-binding drug to target DNA by surface plasmon resonance biosensor technology. Curr Drug Targets 5(1):61–72. https://doi.org/10.2174/ 1568008053174697 17. Schuck P, Huaying Z (2010) The role of mass transport limitation and surface heterogeneity in the biophysical characterization of macromolecular binding processes by SPR biosensing. Methods Mol Biol 627:15–54. https:// doi.org/10.1007/978-1-60761-670-2_2 18. Wold MS (1997) Replication protein A: a heterotrimeric, single-stranded DNA-binding protein required for eukaryotic DNA metabolism. Annu Rev Biochem 66(1):61–92. https://doi.org/10.1146/annurev.biochem. 66.1.61 19. Pavani RS, da Silva MS, Fernandes CAH, Morini FS, Araujo CB, Fontes MRM, Sant’Anna OA, Machado CR, Cano MI, Fragoso SP, Elias MC (2016) Replication protein A presents canonical functions and is also involved in the differentiation capacity of Trypanosoma cruzi. PLoS Negl Trop Dis 10(12):e0005181. https://doi.org/10.1371/journal.pntd. 0005181 20. Pavani RS, de Lima LP, Lima AA, Fernandes CAH, Fragoso SP, Calderano SG, Elias MC (2020) Nuclear export of replication protein A in the nonreplicative infective forms of Trypanosoma cruzi. FEBS Lett 594 (10):1596–1607. https://doi.org/10.1002/ 1873-3468.13755 21. Pavani RS, Vitarelli MO, Fernandes CAH, Mattioli FF, Morone M, Menezes MC, Fontes MRM, Maria Cano MIN, Elias MC (2017) Replication protein A-1 has a preference for the telomeric G-rich sequence in Trypanosoma cruzi. J Eukaryot Microbiol 65(3):345–356. https://doi.org/10.1111/jeu.12478 22. Vaidyanathan VG, Xu L, Cho BP (2013) Binding kinetics of DNA-protein interaction using surface plasmon resonance. Protocol Exchange. https://doi.org/10.1038/protex. 2013.054 23. Biacore® Sensor Surface Handbook T200 (2010) 28-9768-78 Version AA. General Electric Company

Chapter 14 Expression, Purification, and Solution-State NMR Analysis of the Two Human Single-Stranded DNA-Binding Proteins hSSB1 (NABP2/OBFC2B) and hSSB2 (NAPB1/OBFC2A) Serene El-Kamand, Mar-Dean Du Plessis, Teegan Lawson, Liza Cubeddu, and Roland Gamsjaeger Abstract Single-stranded DNA-binding proteins (SSBs) are essential to all living organisms as protectors and guardians of the genome. Apart from the well-characterized RPA, humans have also evolved two further SSBs, termed hSSB1 and hSSB2. Over the last few years, we have used NMR spectroscopy to determine the molecular and structural details of both hSSBs and their interactions with DNA and RNA. Here we provide a detailed overview of how to express and purify recombinant versions of these important human proteins for the purpose of detailed structural analysis by high-resolution solution-state NMR. Key words DNA repair, hSSB1, hSSB2, NMR, Cancer

1

Introduction Cellular transactions involving nucleic acids such as DNA replication, repair, recombination, and transcription result in the unwinding of the DNA double helix exposing single-stranded DNA (ssDNA). In order to protect exposed ssDNA from damage, all living organisms have evolved single-stranded DNA-binding (SSB) proteins. This family of SSBs is characterized by a conserved oligonucleotide/oligosaccharide-binding (OB) domain followed by a flexible extension that interacts with specific protein partners [1]. In humans, three SSB family members have evolved: replication protein A (RPA), human SSB1 (hSSB1), and human SSB2 (hSSB2). While RPA is exceptionally well characterized, the crucial roles of hSSB1 and hSSB2 in the maintenance of genomic stability have only emerged in the last 10 years [2, 3].

Serene El-Kamand, Mar-Dean Du Plessis, and Teegan Lawson contributed equally to this work. Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_14, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Nuclear magnetic resonance (NMR) spectroscopy has been successfully utilized to characterize protein-nucleic acid interactions in solution over many decades [4–7]. The advantages of NMR compared to other structural techniques lie in the facts that NMR [1] can be used in solution rather than in the solid phase (such as crystallography or cryo-electron microscopy), [2] is able to provide information about protein dynamics, and [3] can be applied to measure weak or ultra-weak (down to nM range) interactions between biomolecules such as proteins and nucleic acids. In addition, NMR spectroscopy can be used to delineate the structural details of these interactions by mapping and identifying important protein residues or DNA/RNA bases that are essential for complex formation. We, and others, have used NMR extensively to study the OB domain family of SSB proteins and their interaction with both DNA and RNA [8–13]. In this chapter, we describe in detail how to produce large-scale recombinant isotopically labeled hSSB proteins for the purpose of structurally characterizing protein-DNA and -RNA interactions by solution NMR spectroscopy.

2

Materials

2.1 Preparation, Cloning, and Transformation of hSSB Constructs

1. DNA constructs of OB domains of hSSB1 (1–123) and hSSB2 (1–125), respectively, or full-length hSSBs optimized for expression in Escherichia coli (with restriction recognition sites for BamH1 and EcoR1) in a generic plasmid. 2. pGEX-6P expression vector. 3. Elution buffer: 10 mM TRIS, pH 8. 4. KCM buffer: 500 mM KCl, 150 mM CaCl2, 250 mM MgCl2. 5. E. coli DH5α chemically competent cells (for DNA production) stored at 80  C. 6. E. coli BL21 (DE3) chemically competent cells (for protein expression) stored at 80  C. 7. Luria-Bertani (LB) medium: 1% (w/v) Casein peptone, 0.5% (w/v) yeast extract + 1% (w/v) NaCl. 8. LB agar: 1% (w/v) Casein peptone, 0.5% (w/v) yeast extract, 1% (w/v) NaCl, 100 μg/mL ampicillin + 1.5% (w/v) bacteriological agar. 9. 1 M Ampicillin (AMP). 10. Plasmid mini preparation kit. 11. BamH1 restriction endonuclease. 12. EcoR1 restriction endonuclease. 13. Alkaline phosphatase.

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14. 10 CutSmart buffer. 15. 6 DNA loading dye. 16. GeneRuler DNA Ladder mix. 17. TAE buffer, 1: 40 mM Tris–acetate (pH 8.2), 1 mM EDTA. 18. 1% (w/v) TAE-agarose gel supplemented with 0.0001% (v/v) GelRed nucleic acid stain. 19. PCR and gel kit. 20. Quick-Stick DNA ligase. 21. 4 Quick-Stick ligase buffer. 2.2 15Nand 13C-Labeled Recombinant hSSB Protein Expression

1. Benchtop biofermenter (see Fig. 1a). 2. LB medium. 3. 1 L Trace metal solution (see Note 1): 6 g FeSO4  7 H2O, 6 g CaCl2  2 H2O, 0.8 g MnCl2, 0.45 g CoCl2, 0.7 g ZnSO4  7 H2O, 0.3 g CuSO4, 0.02 g H3BO3, 0.25 g (NH4)6Mo7O24  4 H2O, 5 g EDTA. 4. 3 L Minimal media: 39 g KH2PO4, 30 g K2HPO4, 27 g Na2HPO4, 7.2 g K2SO4 (see Note 2). 5. Overnight (14–16 h) culture solution (see Note 3): 12 g Glucose, 0.025 g thiamine, 20 g yeast extract, 1 mL ampicillin, 10 mL trace metal solution, 2 g unlabeled NH4Cl (for singlelabeled 15N preparations) or 8 g labeled 15NH4Cl (for doublelabeled 13C–15N preparations).

Fig. 1 Expression of 15N-labeled hSSB1 in a biofermenter. (a) Image of used biofermenter. (b) Trace of temperature and oxygen content of the expression medium as a function of time. Note the two spikes in the oxygen content (labeled as 1 and 2 in the figure) indicating that the culture has exhausted its supply of ammonium chloride. Also note the subsequent increase in oxygen content as a consequence of the lower temperature (followed by addition of IPTG to induce protein expression; see Subheading 3 for details)

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6. Fermentation solution: 200 μL Antifoam solution, 25 mL trace metal solution, 2 mL ampicillin, 5 g MgCl2, 0.075 g thiamine, 20 g yeast extract, 30 g glucose, and 4 g of NH4Cl for single 15 N-labeled preparations, and 5 g glucose and 10 g of 15 NH4Cl for double-labeled preparations. 7. pH regulator: 5 M NaOH. 8. Protein expression inducer: 0.5 M Isopropyl β-D-1-thiogalactopyranoside (IPTG). 2.3 hSSB Protein Purification

1. Lysis buffer: 10 mM MES, pH 6.0 (see Note 4), 500 mM NaCl, 0.5 mM PMSF, 10 μg/mL DNase I, 0.1% Triton X-100 (3 mM Tris(2-carboxyethyl)phosphine hydrochloride (TCEP) if reduced, see Note 5). 2. Wash buffer: 10 mM MES pH 6.0, 500 mM NaCl (3 mM TCEP if reduced). 3. Ion-exchange buffer A/SEC buffer: 10 mM MES, pH 6.0, 50 mM NaCl (3 mM TCEP if reduced). 4. Ion-exchange buffer B: 10 mM MES, pH 6.0, 1 M NaCl (3 mM TCEP if reduced). 5. 1 LDS loading buffer: 0.3125 M Tris pH 6.8, 10% (w/v) LDS, 50% (w/v) glycerol, 0.5% (w/v) bromophenol blue. 6. 1 MES acrylamide gel running buffer: 50 mM MES (pH 7.3), 50 mM Tris, 0.01% (w/v) SDS, 1 mM EDTA. 7. Coomassie Brilliant Blue (BBR) acrylamide gel stain: 0.125% (w/v) Coomassie BBR, 40% (v/v) methanol, 70% (w/v) acetic acid. 8. Destain: 10% (w/v) Acetic acid, 30% (v/v) methanol. 9. Protein molecular weight standard. 10. Bis–Tris mini gradient gels (4–20%). 11. Glass gravity columns. 12. Glutathione sepharose (GSH) beads. 13. Human rhinovirus 3C (HRV-3C) protease. 14. Fast-performance liquid chromatography (FPLC) system.

2.4 NMR Spectroscopy

1. NMR buffer: 10 mM MES, pH 6.0, 50 mM NaCl (3 mM TCEP if reduced, see Note 5). 2. D2O and 4,4-dimethyl-4-silapentanesulfonic acid (DSS) for preparation of NMR samples. 3. NMR microtubes: 5 and 3 mm tubes. 4. NMR spectrometers: Bruker 600 or 800 MHz spectrometers run with TopSpin software.

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5. Software for analysis (other appropriate software can be used): Sparky (T. D. Goddard and D. G. Kneller, University of California at San Francisco).

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Methods

3.1 Preparation, Cloning, and Transformation of hSSB Constructs 3.1.1 Transformation and Preparation of hSSB Constructs

1. Transfer 1 μL of commercially synthesized hSSB construct in the generic plasmid (10% w/v in elution buffer) to labeled tubes containing 50 μL KCM and place on ice. 2. Thaw E. coli DH5α competent cells on ice. 3. Transfer 50 μL of E. coli cells into plasmid-containing tube and ice for 15–20 min. 4. Heat-shock the mixtures at 42  C for 90 s. 5. Add 200 μL LB medium to the tubes and incubate at 37  C for 60 min. 6. Transfer 100 μL of each mixture onto separate LB-AMP agar plates and incubate overnight at 37  C. 7. Add approximately 5 mL LB medium and 5 μL of AMP to a 15 mL conical tube (see Note 6). 8. Using a sterile pipette tip, collect a single bacterial colony from the transformation plate and inoculate the tube. 9. Incubate cultures at 37  C in a shaking incubator overnight. 10. Extract/purify the DNA from the transformed E. coli overnight culture using the plasmid mini kit following the manufacturer’s protocol.

3.1.2 Cloning of Prepared hSSB Constructs into Expression Vector (pGEX-6P)

1. To 30 μL of freshly prepared plasmid DNA, add 4 μL 10 CutSmart buffer, 4 μL filtered Milli-Q water, 1 μL BamH1 restriction endonuclease, and 1 μL EcoR1 restriction endonuclease (see Note 7). 2. Incubate at 37  C for 60 min. 3. Add 2 μL of alkaline phosphatase and incubate at 37  C for another 30 min. 4. Add 2 μL of 6 loading dye to each DNA sample and load onto 1% TAE-agarose gel. 5. Electrophorese the samples at 120 V for 30 min in 1 TAE buffer. 6. Image the gel using a UV transilluminator to identify the DNA bands correlating to the BamH1-EcoR1-digested hSSB construct (from generic plasmid) and the pGEX-6P vector. 7. Extract the DNA fragments from the excised gel using the PCR and gel kit, following the manufacturer’s protocol.

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8. Combine the appropriate amounts of BamH1/EcoR1 restriction-digested pGEX-6P vector and insert (molar ratio 1:3) to appropriate volumes of Quick-Stick DNA ligase, 4 Quick-Stick ligase buffer (1 final concentration), and filtered Milli-Q water, according to the manufacturer’s protocol. 9. Incubate at room temperature for 15 min. 10. Follow the steps for transformation, overnight cultures, and DNA extraction/preparation as described in Subheading 3.1.1 to obtain purified miniprep DNA of hSSB constructs cloned into pGEX-6P. 11. Perform a diagnostic digest (using the same restriction enzymes as described above) and TAE-agarose gel analysis to determine the size of the hSSB constructs and vector (pGEX6P), respectively, to confirm their identity (see Note 8). 12. Transform the hSSB construct in pGEX-6P into E. coli BL21 (DE3) as described in the preparation for protein expression (see Subheading 3.2). 3.2 15Nand 13C-Labeled Recombinant hSSB Protein Expression 3.2.1 Expression Day 1

1. Inoculate 5 mL of LB medium with a single colony of transformed E. coli, add 5 μL AMP, and incubate at 37  C for 8 h. 2. Prepare 3 L of minimal media (see Note 2) and autoclave. 3. Place foil over any open hole on the benchtop biofermenter (see Fig. 1a for an image), including the condenser, and autoclave. 4. Add 5 mL day culture and overnight culture solution to 1 L minimal medium in the biofermenter vessel, and incubate at 37  C for 16–18 h.

3.2.2 Expression Day 2

1. Connect pH and oxygen probes (see Note 9), temperature sensor, and air and cooling rod, and attach heating jacket, condenser, and stirrer to the fermenter. Ensure that all holes are covered, and seals are tight. 2. Turn on controller and cooler unit, and open fermenter software. 3. Pour 2 L of minimal media (see Note 2) into the fermenter, and set temperature to 37  C, stirrer to 100 rpm, and pH to 7. 4. Once 37  C is reached, add fermentation solution to the fermenter and wait for pH to equilibrate (hold stable at pH 7 during the entire growth by controlled addition of 5 M NaOH). 5. Add overnight culture to the fermenter. Take absorbance at starting time (absorbance should be around 1–1.5 OD600nm). 6. Set oxygen level (pO2) using the slope within the calibration software to a value of between 80 and 100% and agitation to 1000 rpm. The oxygen level must decrease as the culture grows.

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7. Measure absorbance at regular intervals. 8. Follow instructions in Subheading 3.2.3 or 3.2.4 depending on the type of isotopic labeling required. 3.2.3 Single-Labeled 15N Expression

1. The culture should ideally grow to an OD600nm value of between 3.5 and 5 (see Note 10). Once the first spike in the pO2 value is reached (culture has exhausted its supply of NH4Cl) add 0.5 g 15NH4Cl and wait for a second spike (see Note 11). 2. When the second spike is reached, remove heating jacket and set temperature to 25  C. 3. Once the culture reaches 25  C, add 3.75 g 15NH4Cl and 0.5 mL 1 M IPTG to induce recombinant protein expression. Incubate for 18 h and then harvest cells. 4. For a representative example of diagram depicting temperature, pH, and oxygen content as a function of time (15N-labeled hSSB1) please refer to Fig. 1b.

3.2.4 Double-Labeled 15 N–13C Expression

1. The culture should ideally grow to an OD600nm value of between 3.5 and 5 (see Note 10). Once the first spike in the pO2 value is reached (culture has exhausted its supply of glucose), add 0.5 g 13C-glucose and wait for a second spike (see Note 11). 2. When the second spike is reached, remove heating jacket and set temperature to 25  C. 3. Once the culture reaches 25  C, add 8 g 13C-glucose, 2 g 15 NH4Cl, and 0.5 mL 1 M IPTG to induce recombinant protein expression. Incubate for 18 h and then harvest cells.

3.3 hSSB Protein Purification

1. Resuspend harvested E. coli cell pellet in lysis buffer (20 mL for 5 g cells).

3.3.1 Cell Lysis

2. Mechanically lyse resuspended cells by sonication (3  1 min in 20 mL volume). 3. Centrifuge lysed cells at around 30,000  g for 30 min at 4  C to separate insoluble cell debris from the soluble cell lysate. Take a sample of the lysate for gel analysis.

3.3.2 Glutathione Sepharose (GSH) Affinity Chromatography

1. Prepare glass gravity column by adding GSH beads. 1 mL of beads is used per 1 L of culture. 2. Wash beads with MQW (3  10 CV). 3. Wash beads with wash buffer (3  10 CV). 4. Incubate cell lysate on GSH beads for 1 h at 4  C on a rocker.

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5. Allow the cell lysate to run through the column. Take a sample of the unbound fraction for gel analysis. 6. Wash beads with wash buffer (5  10 CV). 7. Wash beads with ion exchange buffer A (5  10 CV). Take a pre-cleavage bead sample for gel analysis. 8. Add HRV-3C protease directly to beads, incubate for 15 h at 4  C, and then elute the protein (see Note 12). Take a sample of the beads post-cleavage and the eluted protein for gel analysis. 3.3.3 Heparin Affinity Chromatography

1. Equilibrate a HiTrap HP heparin column (5 mL) with ion exchange buffer A (see Note 13). 2. Bind the eluate from GSH affinity chromatography onto the heparin column in ion exchange buffer A. 3. Elute the protein from the column by increasing salt concentration gradient from 50 mM NaCl (ion exchange buffer A) to 1000 mM NaCl (ion exchange buffer B) (see Note 14). 4. Collect fractions correlating to a distinct peak in UV absorbance (280 nm) and concentrate using a 3000 kDa molecular weight cutoff concentrator in preparation for NMR analysis. Figure 2a shows a representative chromatograph of the hSSB1 OB domain (1–123) purification. 5. During concentrating, top up the solution using NMR buffer at least five times at 1:10 to buffer exchange the protein in preparation for NMR analysis (see Subheading 3.4).

3.3.4 Protein Gels (SDS-PAGE)

1. Add 1 LDS loading dye to protein samples and load onto precast Bis–Tris gels with molecular weight standard. 2. Run the gel for 20 min at 180 V in 1 MES running buffer. 3. Stain with Coomassie Brilliant Blue gel stain for 30 min, rocking at room temperature. 4. Incubate with destain solution for 2 h in order to visualize the gel. Please see Fig. 2b for an example of a SDS gel of hSSB1 (1–123) purification.

3.4 NMR Spectroscopy

1. Ensure that the final concentration of the SSB protein from the concentrating step (see Subheading 3.3.3) is between 300 μM and 1 mM. 2. Prepare the NMR sample by using the appropriate volume (depending on the NMR tube used) of protein and add 5–10% D2O and 1 μL of DSS (for shimming and referencing, see Note 15).

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Fig. 2 Purification of hSSB1 by GSH and heparin affinity chromatography. (a) Heparin chromatograph depicting conductivity (mS/cm; blue line) and UV absorbance (AU, orange line) as a function of the elution volume (mL). Note the fraction of purified protein shaded in blue. (b) Image of SDS protein gel showing pre- and postinduction; soluble, unbound bead pre- and post-cleavage; and protein elution fractions

Fig. 3 Portion of representative HSQC spectrum of under reducing conditions

15

N-labeled hSSB1 (1–123)

3. Run the chosen NMR experiment on the NMR spectrometer at a temperature of 298 K (see Note 16). A representative 15N HSQC spectrum of hSSB1 (1–123) under reducing conditions is shown in Fig. 3. For experiments where DNA or RNA is titrated into a NMR sample (15N HSQC NMR titration experiment) containing either hSSB1 or hSSB2 the oligonucleotides must be dissolved in the exact same buffer as the sample. 4. Analyze the NMR data using TopSpin and Sparky software.

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Notes 1. Make up the volume to 1 L with Milli-Q water and stir constantly for >2 h; keep refrigerated at 4  C for storage. 2. Make up 3 L of minimal media in a 5 L flask, and split into 1 L (for overnight culture) and 2 L (for fermenter). 3. Dissolve glucose, thiamine, and yeast extract in minimal MilliQ water and heat until clear. Add ampicillin, NH4Cl or 15 NH4Cl, and trace metal solution after heating. 4. A pH of 6.0 is used for the purpose of NMR analysis. If the analysis technique allows, a higher pH can be used for purification. The use of a higher pH does increase protein yield. 5. TCEP has previously been established to prevent oligomer formation of hSSB1 [14]. Reduced conditions require an addition of 3 mM TCEP to the buffers detailed in Subheading 2. Purifying hSSB1 under reducing conditions (monomeric form) results in higher protein yields compared to nonreducing conditions. 6. For overnight cultures, at least two cultures per construct should be prepared, in the case of failed growth (correct labeling of all samples is essential). 7. The amounts of each component for the restriction digest will vary depending on the volume of plasmid DNA used. 8. For the diagnostic digest, only 5 μL of the ligated plasmid, 0.5 μL of each restriction enzyme, 2 μL of 10 CutSmart buffer, and 12 μL filtered Milli-Q water should be used. 9. The pH probe and the oxygen probe should be kept in 3 M KCl and Milli-Q water, respectively, when not in use. 10. A pre-recorded growth curve should be used as a guide to determine the best OD600nm value to induce protein expression (center point of exponential growth phase). Adjustments to the amounts of NH4Cl and glucose have to be made if the OD600nm is outside of the recommended range of 3.5–5 immediately prior to protein induction. 11. The first spike in pO2 indicates that the bacterial cells have exhausted their supply of NH4Cl or glucose; a small amount of either 15NH4Cl (single-labeled preparation) or 13C-glucose (double-labeled preparation) is added to ensure that the culture adjusts to the labeled material. 12. Cleaving and eluting multiple times at 1–2-h intervals before a final 15-h cleavage does increase protein yield. The HRV-3C protease leaves a 5-residue linker between the GST moiety and the start of the hSSB amino acid sequence.

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13. Heparin affinity chromatography is used for protein expressed in fermenters to remove any bound DNA from the protein. If bound DNA is not an issue, size-exclusion chromatography (SEC) could alternatively be used: (a) Equilibrate a Superdex 75 chromatography column with SEC buffer. (b) Inject eluate from GSH affinity chromatography and run through the column in SEC buffer. (c) Monitor absorbance at 280 nm and collect fractions that correlate to a distinct peak in absorbance. 14. The column can be operated on a fast-performance liquid chromatography (FPLC) system. 15. For transferring small amounts of protein solution into NMR tubes PTFE syringe tubing (gauge 12 and 16, 12 in.) can be used. 16. The type of experiments determines the pulse program chosen from the Bruker library. Initial experiments recorded with hSSBs were 1D and 2D 15N HSQC as well as 2D 13C HSQC (aliphatic and aromatic) followed by 3D CBCA(CO)NH, 3D HNCACB, 3D HNCO, 3D HN(CA)CO, 3D CC(CO)NH TOCSY, 3D HCC(CO)NH TOCSY, 3D (H)CCH TOCSY, and 3D 15N NOESY experiments to assign all backbone and side-chain resonances. Note that at a temperature of 298 K and below some of the signals experience intermediate exchange, preventing the observation of any intermolecular NOEs, whereas higher temperatures result in protein degradation which prevents a full 3D structure calculation of both hSSB1 and hSSB2 (for more details see [2, 3, 8–10, 12, 13]).

Acknowledgment We would like to thank Dr. Ann Kwan from the University of Sydney for expert advice and maintenance of NMR spectrometers. References 1. Dickey TH, Altschuler SE, Wuttke DS (2013) Single-stranded DNA-binding proteins: multiple domains for multiple functions. Structure 21:1074–1084. https://doi.org/10.1016/j. str.2013.05.013 2. Croft LV et al (2018) Human single-stranded DNA binding protein 1 (hSSB1, OBFC2B), a critical component of the DNA damage response. Semin Cell Dev Biol 86:121–128. https://doi.org/10.1016/j.semcdb.2018.03. 014

3. Lawson T, El-Kamand S, Kariawasam R, Richard DJ, Cubeddu L, Gamsjaeger R (2019) A structural perspective on the regulation of human single-stranded DNA binding protein 1 (hSSB1, OBFC2B) function in DNA repair. Comput Struct Biotechnol J 17:441–446. https://doi.org/10.1016/j.csbj. 2019.03.014 4. Bieri M, Kwan AH, Mobli M, King GF, Mackay JP, Gooley PR (2011) Macromolecular NMR spectroscopy for the non-spectroscopist:

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beyond macromolecular solution structure determination. FEBS J 278:704–715. https://doi.org/10.1111/j.1742-4658.2011. 08005.x 5. Kaptein R (2013) NMR studies on proteinnucleic acid interaction. J Biomol NMR 56:1–2. https://doi.org/10.1007/s10858013-9736-8 6. Kwan AH, Mobli M, Gooley PR, King GF, Mackay JP (2011) Macromolecular NMR spectroscopy for the non-spectroscopist. FEBS J 278:687–703. https://doi.org/10. 1111/j.1742-4658.2011.08004.x 7. O’Connell MR, Gamsjaeger R, Mackay JP (2009) The structural analysis of proteinprotein interactions by NMR spectroscopy. Proteomics 9:5224–5232. https://doi.org/ 10.1002/pmic.200900303 8. Kariawasam R, Knight M, Gamsjaeger R, Cubeddu L (2018) Backbone 1H, 13C and 15N resonance assignments of the OB domain of the single stranded DNA-binding protein hSSB2 (NABP1/OBFC2A) and chemical shift mapping of the DNA-binding interface. Biomol NMR Assign 12:107–111. https:// doi.org/10.1007/s12104-017-9789-9 9. Kariawasam R, Touma C, Cubeddu L, Gamsjaeger R (2016) Backbone (1)H, (13)C and (15)N resonance assignments of the OB domain of the

single stranded DNA-binding protein hSSB1 (NABP2/OBFC2B) and chemical shift mapping of the DNA-binding interface. Biomol NMR Assign 10:297–300. https://doi.org/10.1007/ s12104-016-9687-6 10. Lawson T et al (2020) The structural details of the interaction of single-stranded DNA binding protein hSSB2 (NABP1/OBFC2A) with UV-damaged DNA. Proteins 88:319–326. https://doi.org/10.1002/prot.25806 11. Morten MJ, Gamsjaeger R, Cubeddu L, Kariawasam R, Peregrina J, Penedo JC, White MF (2017) High-affinity RNA binding by a hyperthermophilic single-stranded DNA-binding protein. Extremophiles 21:369–379. https://doi.org/10.1007/s00792-016-0910-2 12. Touma C et al (2017) A data-driven structural model of hSSB1 (NABP2/OBFC2B) selfoligomerization. Nucleic Acids Res 45:8609–8620. https://doi.org/10.1093/ nar/gkx526 13. Touma C et al (2016) A structural analysis of DNA binding by hSSB1 (NABP2/OBFC2B) in solution. Nucleic Acids Res 44:7963–7973. https://doi.org/10.1093/nar/gkw617 14. Paquet N et al (2016) hSSB1 (NABP2/ OBFC2B) is regulated by oxidative stress. Sci Rep 6:27446. https://doi.org/10.1038/ srep27446

Chapter 15 Atomic Force Microscopy Reveals that the Drosophila Telomere-Capping Protein Verrocchio Is a Single-Stranded DNA-Binding Protein Alessandro Cicconi, Emanuela Micheli, Grazia Daniela Raffa, and Stefano Cacchione Abstract Atomic force microscopy (AFM) is a scanning probe technique that allows visualization of biological samples with a nanometric resolution. Determination of the physical properties of biological molecules at a single-molecule level is achieved through topographic analysis of the sample adsorbed on a flat and smooth surface. AFM has been widely used for the structural analysis of nucleic acid-protein interactions, providing insights on binding specificity and stoichiometry of proteins forming complexes with DNA substrates. Analysis of single-stranded DNA-binding proteins by AFM requires specific single-stranded/ double-stranded hybrid DNA molecules as substrates for protein binding. In this chapter we describe the protocol for AFM characterization of binding properties of Drosophila telomeric protein Ver using DNA constructs that mimic the structure of chromosome ends. We provide details on the methodology used, including the procedures for the generation of DNA substrates, the preparation of samples for AFM visualization, and the data analysis of AFM images. The presented procedure can be adapted for the structural studies of any single-stranded DNA-binding protein. Key words Atomic force microscopy, DNA-protein complex, Single-stranded DNA, Binding position analysis, Volume analysis, Ver, Telomeric DNA

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Introduction Atomic force microscopy (AFM) is a microscopy technique belonging to the scanning probe microscopy (SPM) family [1]. Compared with its predecessor scanning tunneling microscope (STM) that uses the tunneling of electrons between two conductors in close proximity [2], the AFM proved to be more suitable for biological samples both in air and under solution, because it does not require a conducting surface [3–6]. Rather, AFM imaging exploits the attractive and repulsive forces (i.e., van der Waals, electrostatic interactions, and Pauli repulsion) interacting between a probe and the sample surface to generate a tridimensional representation of

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_15, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Atomic force microscopy. (a) Picture of the MultiMode NanoScope used for sample imaging. (b) Schematic representation of the AFM setup. The probe scans the sample surface on the x- and y-axes (dotted path). A laser beam points to the edge of the cantilever, gets reflected on a mirror, and finally hits the photodetector, providing information on the z position of the cantilever. The feedback loop acts on the piezoelectric to modulate the distance between the sample and the tip, maintaining the oscillation amplitude of the cantilever constant. The information on the x, y, and z positions are translated into a topographic image

the sample surface topography. A sharp tip mounted on an elastic cantilever scans the sample surface on the xy plane, while the movements of the cantilever on the z axis are detected through the use of a laser beam and a photodetector. A feedback loop keeps constant the forces between the tip and the sample by acting on a piezoelectric element to modulate the tip-sample distance (Fig. 1a, b). For biological sample imaging, the cantilever vibrates above the sample surface, making contact with it only at its lower oscillation position. This approach is known as tapping mode and allows minimizing the contacts between the tip and the surface to preserve the integrity of soft biological samples. In tapping mode, the oscillation amplitude of the cantilever, that is dependent on the distance between the tip and the sample, is maintained constant by the feedback loop [7]. The main issue for obtaining high-resolution AFM imaging is the shape of the tip interacting with the sample surface. Due to the radius of curvature of the tip, that is ~10 nm for a standard silicon tip, the sample profile is broadened, an effect called tip convolution (Fig. 2a). This effect can be reduced using ultrasharp tips that end with a single-walled carbon nanotube (Fig. 2b) [8, 9]. Another limit imposed by the shape of the tip is represented by its inability to image structures with undercuts, such as a sphere. These tip-induced distortions must be considered when determining the volume of molecules by AFM imaging. Finally, additional distortions can be caused by a fracture of the tip or by the presence of debris that remain attached to the tip, causing the presence of artifacts in the topography of the image (Fig. 2c).

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Fig. 2 Tip convolution effect. (a) The radius of curvature of the tip determines the shape of the sample profile (dotted line). (b) Tips ending with a carbon nanotube dramatically reduce the tip convolution. (c) A fracture of the tip causes artifacts in the sample profile

Besides its high resolution, another advantage of AFM is the simple and quick sample preparation, since it does not require fixation or coating of the samples, preserving the structural features of biological molecules. Moreover, it allows imaging biological samples in physiological conditions in a liquid environment. For this reason, AFM imaging in liquid has been widely used to visualize in real time at a single-molecule level biological processes and protein conformational changes induced by the interaction with a specific ligand or substrate [10–15]. On the contrary, AFM in air requires the sample to be dried before imaging. This precludes its use to visualize dynamic processes in real time. Nevertheless, it allows the visualization of biological molecules at a single-molecule resolution, providing a tool to analyze individual molecules and rare events that would be otherwise undetectable with bulk biochemical assays. Thus, it is not surprising that AFM has been extensively used to analyze protein-nucleic acid interactions [16– 19]. AFM imaging allows measuring three main parameters of the features of DNA-protein complexes. The first one is the analysis of the binding position of a protein on a DNA substrate that can be dependent on the presence of a specific binding site on the substrate. The binding position of a protein on a DNA substrate can be determined by measuring the contour length (CL) of the DNA molecules [20–25]. This approach has also been used to analyze nucleosomal spacing on in vitro chromatin assembly samples [26– 30]. For single-stranded DNA (ssDNA)-binding proteins, the generation of a DNA construct containing single-stranded regions is required to analyze the binding of the protein to its position [31– 33]. Another parameter that can be analyzed through AFM imaging is the volume of globular proteins. This measurement can provide information on the stoichiometry of a protein and its multimerization status when it binds the DNA [23, 32, 34– 37]. Finally, AFM can be used to measure protein-induced bending of the DNA substrate [38–40]. Here we describe the details of the procedure used to analyze ssDNA-protein complexes formed by the Drosophila melanogaster protein Verrocchio (Ver) [41]. This protein is a component of the telomeric complex terminin, which localizes at Drosophila

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telomeres in a sequence-independent manner [42–48]. Several DNA constructs that mimic the hypothetical structure of Drosophila telomeres were generated in order to assess Ver binding to the terminal ssDNA on both 30 and 50 orientations. The described methodology, from the generation of the DNA substrates to the image analysis, can be applied to the study of any ssDNA-binding proteins with only minor modifications.

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Materials

2.1 Oligonucleotide Sequences

1. 1246 fw: 50 -GGTGATGACGGTGAAAACCT-30 . 2. 1246 rv: 50 -GCCTACATACCTCGCTCTGC-30 . 3. ss60: 50 - GCGGTGATCGTTCACGGATTGCTCAATGCC TATTGACCAGAGGAAAGCCGCAGATCCCTCT-30 . 4. ss68: 50 - TGCATGCAGCGGTGATCGTTCACGGATTGCT CAATGCCTATTGACCAAGAGGAAAGCCGCA GATCCTCT-30 . 5. Ada-30 : 50 -TCCGTGAACGATCACCGCTGCA-30 . 6. Ada-50 : 50 -TGCAAGAGGATCTGCGGCTTTC-30 .

2.2 Preparation of the DNA Substrates and Binding with the Protein

1. 0.5 and 1.5 mL Eppendorf tubes. 2. Micropipettes and disposable pipet tips. 3. Glassware. 4. Ultrapure water (18.2 MΩ∙cm). 5. 50 mL disposable plastic syringes. 6. Whatman syringe filter (0.02 μm). 7. 5 M NaCl solution. 8. 1 M Tris–HCl pH 7.5 solution. 9. 0.5 M EDTA pH 8 solution. 10. 1 M HEPES-KOH pH 7.5 solution. 11. 1 M HEPES-KOH pH 7.8 solution. 12. 1 M KCl solution. 13. 1 M DTT solution. 14. pUC18 plasmid. 15. Restriction enzymes: DraI, PstI, Alw44I. 16. Water bath. 17. Phenol:chloroform:isoamyl alcohol (25:24:1) mixture. 18. Chloroform solution. 19. Low-melting-point agarose gel.

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20. 1 TAE buffer: 40 mM Tris–acetate, 1 mM EDTA. 21. 10 mg/mL Ethidium bromide (EtBr). 22. UV transilluminator or UV lamp. 23. Scalpel for cutting agarose gels. 24. Gel extraction kit. 25. DreamTaq DNA Polymerase. 26. 10 mM each dNTP mix. 27. PCR thermocycler. 28. T4 polynucleotide kinase. 29. 100 mM ATP. 30. T4 DNA ligase. 31. Purified 6 His-Ver (or other protein of interest). 32. 2 Binding buffer: 100 mM HEPES-KOH pH 7.8, 100 mM KCl, 0.2 mM EDTA, 2 mM DTT. 33. 25% Glutaraldehyde solution. 34. PCR purification kit. 2.3 Substrate Preparation and Deposition on the Mica

1. Steel sample disk. 2. Cyanoacrylate glue (superglue). 3. Grade V1 mica sheets (75  25 mm). 4. Metal tweezers. 5. Adhesive tape. 6. 10 AFM buffer: 100 mM NaCl.

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7. 10 mM MgCl2 solution. 8. Nitrogen gas. 2.4 AFM Imaging and Analysis

1. Sharp bent stainless steel tweezers. 2. Standard silicon probe for tapping mode. 3. MultiMode SPM Nanoscope Digital III A, equipped with E-scanner or upgraded versions. 4. Heavy marble plate and elastic bungee cords for vibration isolation. 5. NanoScope software. 6. ImageJ software. 7. ImageSXM software.

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Methods

3.1 Preparation of the DNA Substrates

In order to analyze the DNA-binding properties of a specific protein it is essential to use an appropriate DNA substrate that can be specifically recognized by the protein of interest. In our study we analyzed Ver capability to bind the ssDNA portion of constructs resembling chromosome ends. Since the electrophoretic mobility shift assay (EMSA) analysis showed that Ver binds ssDNA in a sequence-independent manner, similarly to other terminin proteins [41], we adapted the protocol developed by Shlyakhtenko et al. [33] to generate three double-stranded DNA (dsDNA) constructs containing single-stranded overhangs of random sequences: a 30 single-tail construct (30 -STC) containing a 42-nucleotide (nt) 30 ssDNA overhang at one end, while the opposite end was blunt; a 30 double-tail construct (30 -DTC) presenting the same 42nt 30 -overhang at both ends; and a 50 double-tail construct (50 -DTC) with a 42-nt 50 single-stranded overhang at both ends. As a negative control, we generated a dsDNA construct with blunt ends (blunt-end construct, BEC). The length of these molecules was chosen so that they would not be too short (less than 100 base pairs), therefore appearing as a globular shape in AFM images, or too long (more than 10 kilobases) resulting in supercoiled shapes difficult to distinguish. These constructs were prepared according to the following protocol: 1. The BEC DNA fragment can be generated by endonuclease digestion of the pUC18 plasmid using a restriction enzyme that generates blunt DNA ends (Fig. 3a). Digest 10 μg of pUC18 with 50 U of DraI overnight at 37  C. 2. Purify the digested DNA with phenol/chloroform extraction. Purification of the digested product is required to obtain a high-quality DNA for AFM imaging. 3. Perform a standard ethanol precipitation to obtain a DNA pellet and let it air-dry. 4. Dissolve the DNA pellet in 45 μL of water. Add 5 μL of 50% glycerol (5% final concentration). 5. Run the DNA solution on a 1% w/v low-melt agarose gel in 1 TAE at 60 V. 6. Stain the gel in an EtBr bath (0.5 μg/mL in 1 TAE) for 30 min. 7. Use UV light to detect the 694 bp digestion product and cut the corresponding band with a clean scalpel. 8. Purify the DNA using a gel extraction kit and following the manufacturer’s instructions.

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Fig. 3 Schematics of the strategy used to generate the DNA constructs. (a) Generation of the BEC DNA fragment. The pUC18 plasmid was digested with DraI and the 694 bp digestion product was purified. (b) The 30 -STC DNA fragment was generated amplifying a 1246 bp region of the pUC18. Digestion of the PCR product with PstI and ligation with the 30 -tail adapter 1 generated the final construct. (c) The 30 -DTC DNA fragment was obtained digesting the pUC18 with Alw44I and purifying the 1216 bp digestion product. Ligation of this fragment with the 30 -tail adapter 2 resulted in the final construct. (d) The 943 bp DNA fragment obtained digesting the pUC18 with Alw44I was purified and ligated with the 50 -tail adapter to obtain the 50 -DTC DNA fragment

9. Elute the DNA with ultrapure water and determine its concentration (see Note 1). Store at 20  C. 10. To obtain the 30 -STC fragment, a dsDNA PCR product can be digested and purified. The ligation of one of the digestion fragments with an adapter generated by the annealing of two oligonucleotides can be performed to obtain the final DNA construct (Fig. 3b). Amplify by PCR a 1246 bp DNA fragment using as template 1 ng of the pUC18 plasmid and 0.2 μM of the primers 1246-fw/1246-rv. After denaturation at 94  C for 30 s, the primers were annealed for 30 s at 55  C before elongation at 72  C for 30 s. The PCR cycle was repeated for a total of 30 times. 11. Precipitate the PCR product with ethanol and air-dry as described above. 12. Resuspend the DNA pellet in 100 μL of H2O and quantify it.

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13. Digest 10 μg of the PCR product with 50 U of the restriction enzyme PstI overnight at 37  C. Digestion of the 1246 bp PCR product with PstI results in two DNA fragments 842 bp and 400 bp long, respectively, both with one blunt end and a 4 nt 50 -TGCA-30 overhang at the other end (see Note 2). 14. Perform an ethanol precipitation step as described in step 11, air-dry the DNA pellet, dissolve it in 45 μL of H2O, and add 5 μL of 50% glycerol. 15. Purify the 842 bp DNA fragment by agarose gel electrophoresis and DNA gel extraction, as described in steps 5–8. Elute with 20 μL of H2O. 16. Phosphorylate the 50 end of the single-stranded 60-nucleotide oligo (ss60). This step is required for the ligation step. Carry out the phosphorylation reaction with 450 pmol of ss60 and 20 U of T4 polynucleotide kinase in the enzyme reaction buffer supplemented with 500 μM ATP, for 30 min at 37  C. 17. To generate the 30 -tail adapter 1, anneal the ss60 with the oligonucleotide adapter 30 (Ada-30 ). Incubate 450 pmol of the phosphorylated ss60 with 450 pmol of Ada-30 in 100 mM NaCl, 10 mM Tris–HCl pH 7.5, and 1 mM EDTA at 95  C for 10 min. After that, slowly cool down the solution (~1  C/min) to 25  C in order to promote the annealing of the two oligos. To stabilize the annealed product, store the solution at 4  C for 2 h (see Note 3). 18. To obtain the 30 -STC, a ligation reaction must be performed with 5 pmol of the purified 842 bp DNA fragment and an excess (20 pmol) of the 30 -tail adapter 1. Carry out the reaction using 10 U of T4 DNA ligase in the reaction buffer supplied with the enzyme at 16  C overnight (see Note 4). 19. The next day, perform a phenol/chloroform purification followed by ethanol precipitation, air-dry the pellet, dissolve it in 45 μL of water, and add 5 μL of 50% glycerol for gel purification. 20. Purify the 30 -STC using agarose gel extraction as described in steps 5–8, loading the non-ligated 842 bp fragment on a separate lane as a control. A 1.6% agarose gel should be used to visualize the small shift in electrophoretic mobility in the ligated sample. 21. Elute the 30 -STC with ultrapure water and quantify. Store at 20  C. 22. The two double-tailed constructs are generated with the same procedure used to obtain the 30 -STC, but with minor adjustments. The pUC18 vector is digested with the restriction enzyme Alw44I to generate three DNA fragments of 1216, 943, and 320 bp in length, all of them with 30 -ACGT-50

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overhangs at both ends. The 1216 bp DNA fragment is ligated with the 30 -tail adapter 2, resulting from the annealing of a 68-nt oligo (ss68) with the Ada-30 , to generate the 30 -DTC (Fig. 3c). On the other end, 50 -DTC is obtained through the ligation of the 943 bp fragment with a 50 -tail adapter generated by the annealing of the ss60 oligo with an adapter 50 (Ada-50 ) (Fig. 3d). Here the 50 phosphorylation step must be performed on the Ada-50 , instead of the ss60. All the procedure steps are performed as described for the 30 -STC (steps 10–21), with the exception of the ligation step, where 5 pmol of the 1216 bp and the 943 bp fragments is incubated with 40 pmol of the 30 -tail adapter 2 and the 50 -tail adapter, respectively. 3.2 Preparation of the Deposition Substrate for AFM Imaging

While the AFM can scan a relatively large area in the x- and y-axes (up to 200  200 μm), the range on the z-axes is much more limited, typically 1–10 μm. Therefore, AFM samples should be deposited on flat, clean, and smooth surfaces. For biological samples, substrates like mica, silicon, and highly oriented pyrolytic graphite (HOPG) are commonly used. Mica muscovite is one of the most used materials, because it is inexpensive and it allows to readily and quickly obtain a new clean surface. This substrate is composed of overlying aluminum silicate layers held together by weak interactions mediated by potassium ions. This renders easy separation of consecutive layers and quick obtaining of a new clean substrate for sample deposition. A small square of mica can be placed on top of a metal specimen disk for AFM and peeled off with adhesive tape with the following procedure. 1. Apply a small amount of glue on one side of the metal disk. 2. Use scissors to cut a small square of mica (~5  5 mm). 3. Position the square of mica at the center of the metal disk with glue and firmly apply pressure on it to obtain an even adhesion of the mica to the metal disk. Keep the disk at room temperature for at least 1 day before use, to let the glue dry. Once ready, the disks with the mica substrate can be stored at room temperature until needed. 4. To peel off the mica, slightly press some adhesive tape against the mica surface. Use tweezers to hold the border of the metal disk and with a quick movement peel off the tape to expose a new mica layer. To determine if the new mica surface is smooth and flat without split or splintered layers carefully check the complementary surface of peeled-off layers on the tape. 5. If necessary, repeat step 4 until the mica surface is homogeneous. Freshly peeled mica must be promptly used for sample deposition to avoid environmental contaminations on the mica surface.

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3.3 DNA Substrate Deposition on the Mica

Before proceeding with the binding of the protein of interest to the DNA substrates, the quality of the DNA constructs must be assessed through visualization with the AFM. Analysis of the DNA molecules will also allow determining the total length of the constructs as they appear with AFM imaging. After addition of the DNA solution on the mica, the potassium ions dissociate from the mica outer layer resulting in a negatively charged surface. Since the DNA phosphate-deoxyribose backbone is negatively charged, bivalent cations must be added to the deposition solution to allow DNA adsorption on the mica by bridging the mica surface and the DNA molecules [16]. A 1.5 mM concentration of MgCl2 is usually added to the DNA solution with AFM buffer for this purpose. For each DNA construct perform the deposition as follows. 1. Prepare a 0.5–2 nM DNA solution in 1 AFM buffer (4 mM HEPES-KOH pH 7.5, 10 mM NaCl) containing 1.5 mM MgCl2 (see Note 5). 2. Add 10 μL of the DNA solution to the center of a freshly peeled mica surface. Be careful not to touch the surface with the pipet tip. 3. Let the DNA adsorb on the mica for 1 min. 4. Use tweezers to hold the metal disk on top of a waste container, tilt the disk at ~45 , and wash the mica surface adding 2 mL of ultrapure water dropwise. The water drops should reach the mica surface, quickly leave the surface, and get into the waste container, in order to wash away the excess of DNA solution. 5. Use a gentle flux of nitrogen to remove the remaining water from the mica surface. This step should be performed with a single quick gesture, moving the nitrogen flux from one corner of the mica to the opposite one and letting the water drop fall down from the metal disk. After that, use the nitrogen flux to completely dry the mica surface. 6. Keep the metal disk in a closed container until visualization, which should be performed within few hours.

3.4 DNA-Protein Complex Preparation and Deposition on the Mica Surface

Before AFM imaging of DNA-protein complexes, the binding conditions need to be optimized. Particularly important variables are the DNA:protein ratio and the buffer composition. Binding conditions should be empirically determined with other assays (e.g., electrophoretic mobility shift assay). After the reaction, the complexes must be cross-linked in order to stabilize the binding of the protein to the DNA. Before proceeding with the sample deposition on the mica, the DNA-protein complexes should be purified using a PCR purification kit. This step is needed to remove both the binding buffer (that could negatively affect the image quality) and the unbound protein molecules. The latter purpose is of utmost

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importance for the DNA-protein complex analysis, because the presence of several unbound protein molecules would result in the presence of DNA molecules only apparently bound by the protein, biasing the analysis with false-positive results. The crosslinking step is therefore required to stabilize the protein bound to the DNA during DNA purification. Once purified, the complexes must be diluted in AFM buffer, making them suitable for deposition on the mica surface as described before. The following steps describe the protocol to perform DNA-protein complex analysis using purified 6 His-Ver and one of the four DNA constructs described above, but can be performed for any ssDNA-binding protein, with the appropriate adjustments to the binding step (see Note 6). 1. Incubate 10 nM of the DNA substrate with 200 nM of 6 HisVer in 20 μL of 1 binding buffer (50 mM HEPES-KOH pH 7.8, 50 mM KCl, 0.1 mM EDTA, 1 mM DTT) for 30 min at 4  C. 2. Cross-link the DNA-protein complexes by adding 0.1% glutaraldehyde and incubating the sample on ice for 30 min (see Note 7). 3. Purify the DNA-protein complexes using a PCR purification kit following the manufacturer’s protocol. 4. Elute the purified DNA-protein complexes with 30 μL of ultrapure water and determine the concentration using a spectrophotometer. 5. Prepare a 0.5–2 nM solution of the DNA-protein complexes in 1 AFM buffer containing 1.5 mM MgCl2. 6. Add 10 μL of the solution from step 5 to the center of the mica and let it stand for 1 min. 7. Wash the excess of DNA-protein complexes with 2 mL of ultrapure water. 8. Dry the mica surface with a mild flux of nitrogen. 3.5 AFM Imaging in Tapping Mode

In tapping mode the cantilever oscillates on the sample surface in the absence of frictional forces, therefore preventing damaging of biological samples. The distance between the vibrating cantilever and the sample is kept constant by the feedback loop that acts on the piezoelectric, allowing the tip to touch the sample only at its lowest point of oscillation. The interaction between the tip and the surface causes a change in the resonance frequency of the cantilever. This change is compensated through the amplitude modulation operated by the feedback loop system that keeps the forces between the sample and the tip at a user-specified amplitude set point. Decreasing the set point results in a decrease in the oscillation amplitude of the cantilever due to dampening effects, while the

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forces between the tip and the sample increase. The feedback loop adjusts the amplitude of the oscillation to the set point value by modulating the z-position of the sample [49]. This amplitude modulation is recorded and converted into a height value for each point of the scanned surface, generating a topographic image of the sample (see Note 8). 1. Remove the microscope head by unfastening the retaining springs on both sides. Lift the head off and set aside. 2. Place the metal disk with the sample on top of the AFM scanner. An internal magnet will hold the disk in place. 3. Remove the tip holder from the AFM head and load the probe in the tip holder using sharp bent metal tweezers (see Note 9). 4. Reassemble the AFM head and secure it with retaining springs. 5. Before placing back the tip holder into the AFM head, increase the distance between the sample and the tip holder by using the Up switch on the microscope base. This step is necessary to avoid the cantilever crashing on the sample surface while putting back in place the tip holder. 6. Insert the loaded tip holder into the AFM head by lifting it carefully over the sample. 7. Align the laser at the edge of the cantilever using the knobs on the AFM head. When the laser is not on the cantilever it appears as a bright red spot on the surface below, while a shadow appears on the sample surface when the laser is aligned on the cantilever. The alignment can be verified by placing a piece of paper in front of the photodetector: the laser spot will appear on the paper. 8. Adjust the mirror level on the back of the microscope head to maximize the signal. 9. Adjust both vertical and lateral deflection to zero in order to center the laser on the photodetector. 10. Manually lower the cantilever on the sample until the probe is in proximity to the surface. 11. Start the NanoScope software. 12. Tune the cantilever by selecting the Autotune button. This step allows finding the resonance peak of the cantilever and adjusting the cantilever oscillation to the appropriate amplitude. 13. Engage the probe to the surface by clicking on Motor on the top panel followed by Engage. 14. Set the scan size to 1–3 μm and the samples/line at 512 (see Notes 10 and 11). 15. Adjust the scan parameters to obtain a good topographic image of the field of interest and acquire the image (see Note 12).

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The binding position can be directly related to the specificity of a protein for a particular sequence or a DNA structure. The binding of a ssDNA-binding protein to the constructs 30 -STC, 30 -DTC, and 50 -DTC is expected to occur at the end of the DNA molecules, where the ssDNA is located. On the contrary, no bound protein is expected to be found on the BEC DNA fragment. However, since the preparation of the AFM samples requires the cross-linking of the DNA-protein complexes, therefore stabilizing even transient interactions, the occurrence of proteins bound to dsDNA portions of the constructs is not rare. A statistical analysis of the frequency of protein binding in each position of the construct can be used to highlight the presence of unequal distribution of the protein along the DNA molecule. Analysis of the binding position of a protein of interest on a DNA molecule can be obtained by measuring the contour length (CL) of the DNA from the center of the bound protein to the closer DNA end (L1, Fig. 4a). A frequency distribution of the protein binding along the DNA construct is obtained by dividing the DNA length in small intervals and assigning each binding event into the appropriate interval according to the measured length L1 (see Note 13). The proteins bound at the end of the DNA molecules are considered with a L1 value of 0, and they are grouped in the first interval (End) (Fig. 4b–e). After that, the distribution is compared with a uniform distribution on the same DNA length using a χ 2 test, to determine if the distribution of bound proteins is significantly different from a uniform distribution. Due to its high flexibility and smaller filament height, visualization of the ssDNA portion of the constructs is difficult to obtain with the standard silicon probe. Therefore, measurements of the DNA lengths are usually made on dsDNA. Since proteins bound at the ssDNA are located at the edge of the molecules, it is important to verify that their binding does not occur on the final dsDNA portion of the construct. To address this issue, it is sufficient to measure the length L2 of the DNA molecules with a terminal bound protein, corresponding to the CL of the DNA fragment from the edge of the protein to the other DNA end. This length is then compared with the CL of the free DNA construct L using a Student’s t-test. A L2 value significantly smaller than L means that the protein is covering the terminal dsDNA portion of the construct, while if the protein is bound to the ssDNA no statistical difference between the two lengths is found (Fig. 4f, g). 1. Flatten the raw AFM images to remove the background noise and export them as TIFF files. 2. Open the TIFF images using the freeware ImageJ. 3. Using the Set scale option in the Analyze menu convert the pixel units into the corresponding μm size, based on the scan size used for each image.

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Fig. 4 Analysis of Ver-binding position along the constructs. (a) Representative AFM images showing DNA molecules bound in an internal position (blue arrows) or at the end (red arrow). The CL from the center of the bound protein to the closest DNA end (L1) was measured using the Segmented Line tool in ImageJ. (b–e) Histograms showing the number of Ver molecules binding in each segment of the BEC (b), 30 -STC (c), 30 -DTC (d), and 50 -DTC (e) fragments. The proteins bound at the end of the constructs were assigned to the first position (End, in red). Counts represent the number of DNA molecules bound in each position. The number of bound DNA molecules measured (n) is shown. Below each graph, a schematic representation of the constructs shows the position of each interval along half DNA molecule. (f) Schematic of the analysis performed to confirm that Ver binds the ssDNA at the end of the constructs rather than the final portion of dsDNA. The CL of each construct in the absence of the protein was measured as described above to obtain the length L (left). For the terminally bound DNA molecules, the contour length was measured starting from the edge of the protein (L2). When the protein is associated to the single-stranded DNA portion of the construct, the length L2 is equal to L (middle), while when the protein covers the final portion of the dsDNA region, L2 is less than L (right). (g) The CLs L and L2 of each construct were measured to obtain a Gaussian distribution. The table shows the mean and standard deviation values of the measured lengths for each DNA fragment. Comparison of the length distribution of bound and unbound DNA molecules was performed using a Student’s t-test. The high p values obtained for the 30 -STC, 30 -DTC, and 30 -DTC confirm that Ver is preferentially associated to the ssDNA portion, while in the few terminally bound BEC fragments it associates with the dsDNA

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4. Measure the distance between the center of the bound protein and the closest DNA end (L1) using the Segmented Line tool. 5. Divide the construct length in small intervals and assign each value L1 to the corresponding section to derive the frequency distribution. Since the measurement is performed between the protein and the nearest DNA end, the greater section will correspond to half the construct length. 6. Divide the total number of bound DNA molecules for the number of sections to obtain the expected frequencies for a uniform distribution on the same construct. 7. Compare the two distributions using a χ 2 test. A p value smaller than 0.05 indicates that the protein distribution along the DNA construct is not uniform due to the presence of a specific binding site. 8. To verify that the terminally bound proteins recognize the ssDNA measure the CL of the DNA molecules bound at either one or both ends (L2) and the CL of the unbound construct (L) following steps 1–4. 9. Compare the two distributions with a Student’s t-test. A p value greater than 0.05 means that the two distributions are not statistically different and suggests that the protein is binding the ssDNA at the end of the DNA construct. 3.7 Analysis of the Stoichiometry of ssDNA-Binding Proteins

Many ssDNA-binding proteins (such as SSB or OB fold-containing proteins) interact with the DNA substrate as multimers, suggesting that protein stoichiometry determines the binding efficiency. Comparison of the volume of free protein molecules with that of proteins bound to the DNA substrates can provide useful information about their binding stoichiometry (see Note 14). When measuring protein volumes in AFM images, some sample deformations must be taken into account. The diameters of objects are usually overestimated in AFM samples due to the tip convolution [50]. On the other hand, object heights are underestimated due to the force exerted by the tip on the sample. To compensate for these effects, the volume of a globular protein can be approximately determined using the formula for the volume of a hemiellipsoid object: V ¼

4 πr r h 6 1 2

where h is the height of the protein and r1 and r2 represent, respectively, the major and minor axes measured at half height of the protein [51, 52]. These parameters can be measured using the freeware Image SXM (Fig. 5a–c, e–g). Frequency distribution of the measured volumes can be plotted in a histogram to reveal the main peaks corresponding to the different multimerization forms

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Fig. 5 Protein volume analysis. (a) Representative AFM image of free Ver molecules on the mica surface. (b) Analysis of the protein height using Image SXM. The Density Slice tool (bar on the left) was used to set a threshold (in red) above the mica surface. The software analyzes the objects in the image that are above the selected threshold and measures their height. (c) The density slice was adjusted to half the height of the measured proteins in order to obtain the r1 and r2 axis values. (d) Histogram showing the volume distribution for unbound Ver. (e) Representative image of a DNA molecule with Ver complexes associated at both its ends. The protein complex at the bottom appears brighter than the one on the top, suggesting that its volume might be greater. (f) The protein complexes are highlighted with the Density Slice tool to determine their height. The threshold was set above the DNA surface. (g) The threshold is adjusted to half height of the protein complex at the top to determine its axes. To measure the parameters of the complex at the bottom, a second adjustment of the density slice is needed to set the threshold at half its height. (h) Volume distribution of the Ver complexes bound to the DNA. This distribution is broader than the one showed in panel (d), suggesting that Ver binds the DNA in multimeric form. Based on Ver molecular weight, the peaks correspond to a dimer and a tetramer volume, as determined using a volume/molecular weight correlation curve

of the protein, providing data on the stoichiometry of the protein of study when present in free form or when bound to the DNA (Fig. 4d, h) (see Note 15). The number of protein monomers forming the multimers in each peak can be determined using a calibration curve that correlates the protein molecular weight with its volume measured by AFM (see Note 16). 1. Open the flattened AFM image with the Image SXM software. 2. Select Set Scale in the Analyze menu to convert the pixel image size to the corresponding μm value. 3. Go to the SPM menu and select Calibration and Set Z Scale to specify the z value of the black (lower) and white (higher) pixels. The z value is dependent on the sample height selected during the image flattening process. For most DNA-protein analysis a sample height of 3 nm is recommended, therefore assigning to the black and white pixels the values of 0 nm and 3 nm, respectively.

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4. Open the Set Measurements window from the Analyze menu and select the Mean Density, the Ellipse Major Axis, and the Ellipse Minor Axis parameters, corresponding, respectively, to the parameters h, r1, and r2 in the volume formula described above. 5. Use the Selection tool to select a small area around one protein that is bound to the DNA. 6. Use the Density Slice tool in the Options menu to set a threshold that includes the pixel density above the mica surface in the samples with the free protein and above the DNA surface in the DNA-protein complex sample. The protein of interest will be marked in red. 7. Select Analyze Particles in the Analyze menu. The software will generate a text file containing the parameters of the protein. However, only the Mean Density (h) value will be used at this point. 8. Use the Density Slice tool to set a threshold that corresponds to half the height measured in the previous step and select Analyze Particles again to obtain the values r1 and r2 at half height of the protein. 9. Use the measured values h (from step 7) and r1 and r2 (from step 8) to calculate the volume of the analyzed protein. 10. Repeat steps 5–9 for different proteins from the same sample to obtain a good distribution of the protein volume. 11. Plot the protein volumes (both for the free and the bound protein) as histograms of frequencies. The peaks in the histogram represent the multimerization forms of the protein.

4

Notes 1. All the solutions used for AFM imaging must be devoid of contaminants; therefore they should be prepared with ultrapure water. All the buffers should also be filtered with 0.02 μm filters. 2. To make sure that the PCR products have real blunt ends, an end repair reaction can be performed before digestion. This step is optional and can be performed to generate the 30 -STC. Use 10 U of the DNA Polymerase I (Klenow fragment) to perform the fill-in reaction on 10 μg of PCR product. Carry out the reaction in the buffer provided with the enzyme supplemented with 100 μM each dNTPs for 30 min at 37  C followed by 20 min at 75  C to inactivate the enzyme. Perform ethanol precipitation, air-dry the DNA pellet, dissolve it in 100 μL of water and quantify, and then proceed with the

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digestion with PstI. End repair is not required if the PCR reaction is carried out with a proofreading Taq DNA polymerase. 3. The quality of the annealed product can be assessed using native polyacrylamide gel electrophoresis (PAGE). Load ~100 pmol of the product on a 12% polyacrylamide (19:1) gel in 1 Tris/borate/EDTA (TBE) buffer, along with an equal amount of the ss60 and Ada-30 oligos in the adjacent wells to assess the reduced electrophoretic mobility of the 30 -tail adapter 1. 4. The ligation reaction should be performed in a small volume (no more than 30 μL) to increase the chances of ligation between the two molecules. An excess of the 30 -tail adapter reduces the chances of ligation between two different 842 bp fragments. 5. The ideal DNA concentration should be experimentally determined, in order to obtain a good number of nonoverlapping DNA molecules in each field. 6. The protein concentration for the binding to the DNA constructs should be adjusted according to its ssDNA-binding affinity. For example, a 200 nM concentration of 6 His-Ver was used for the binding to each DNA construct. Analysis of the DNA-protein complexes using single-stranded binding protein (SSB) from E. coli was performed as a positive control. Since SSBs bind ssDNA with higher affinity compared to Ver, a 10 nM concentration of protein was used for the binding. The binding conditions and the following steps were performed using the same protocol described for Ver. 7. To obtain the final concentration of 0.1% glutaraldehyde make a 0.3% dilution with ultrapure water starting from the 25% glutaraldehyde stock. Add 10 μL of 0.3% glutaraldehyde solution to 20 μL of binding solution prepared in step 1 to obtain a final concentration of 0.1% glutaraldehyde in 30 μL of solution. The 0.3% glutaraldehyde solution can be stored at 4  C for up to 1 month. 8. Since AFM imaging is obtained through the cantilever oscillation, isolation of the microscope from any source of vibration, both in the acoustic and sub-acoustic frequencies, is required to obtain a good image. Structural vibrations can be reduced by placing the microscope on a mechanical vibration isolation table or, alternatively, on a heavy marble plate suspended from elastic bungee cords (the cords must not reach their elastic limit). Acoustic vibration isolation can be achieved by placing the microscope in an acoustic isolation chamber.

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9. Once the tip holder is removed, place it upside down on a table to expose the cantilever holder groove and gently press it against the table surface to lift the spring clip. Carefully insert the probe into the tip holder groove making sure that the edges of the probe are perfectly aligned with the inner edges of the groove. Release the pressure on the tip holder to lower the springer clip that will keep the probe in place. 10. When engaging the surface, it is recommended to set the scan size to 0 and then gradually increase it to 1–3 μm. This operation will prevent damaging the tip if big-size contaminants are present on the surface, such as dust particles. A scan size of 3 μm corresponds to a scanned area of 3 μm2 and it is generally used to obtain a good number of DNA molecules (up to 100) in every single field. Reducing the scan area to 1 μm2 allows obtaining more detailed images of molecules of interest. 11. The sample/line parameter determines the number of lines scanned in each field, and consequently the number of pixels that compose the image. This parameter can be initially set to 128 or 256 to quickly scan the field and determine the presence of interesting features in the area. However, a resolution of 512  512 pixels is recommended when acquiring an image. 12. Parameters such as the amplitude set point, the gains, and the scan rate can be adjusted during the scan to obtain a better quality image. The amplitude set point determines the amount of voltage for the feedback loop. Changes in the set point value affect the response of the cantilever vibration and the amount of force between the sample and the tip. In order to minimize this force, increase the set point until the tip is no longer interacting with the surface (the Z center position indicator in the software interface will move to retracted), and then gradually decrease it until the tip engages the sample again. The gains are three parameters that control the efficiency of the feedback loop in maintaining the cantilever oscillation amplitude in response to the roughness of the sample surface. Increasing these parameters results in a higher quality image; however if their value is too high the image will be distorted due to the appearance of feedback noise. The scan rate determines the speed at which the cantilever scans the surface and affects the efficiency of the feedback loop response. A low scan rate (0.5–2 Hz) is recommended for 1–3 μm2 areas, as it gives enough time to the feedback loop to compensate for the oscillation amplitude changes induced by topographic variations of the sample surface. 13. Since the two ends of constructs are indistinguishable, the binding positions were obtained by measuring the distance between the protein and the nearest DNA end. Consequently,

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the maximum measured length was about half the length of the construct. Each construct was then divided into 10.2 nm long sections (~30 bp) to obtain a good frequency distribution of the protein on half construct. A different size of the sections can be used to divide the measured DNA length. Reducing the section size increases the number of intervals and the accuracy of the distribution of frequencies. 14. The presence of a tag used to purify the protein can significantly alter the protein volume measured with the AFM. Proteolytic removal of the tag or the use of a small-molecularweight tag, such as the poly-histidine tag (6  His), allows obtaining a more reliable average volume value. 15. Deposition of free proteins on the mica can be performed as described for the DNA in Subheading 3.3. The concentration of protein necessary to obtain a good molecule spread on the mica surface should be empirically determined (typically ~1–10 nM). Deposition of proteins does not require MgCl2, due to the presence of partially negative charges on the amino acids that allow interaction with the negative mica surface. 16. There is a positive correlation between protein molecular weights and their volumes determined with the AFM, allowing the distinction of different oligomeric states of a protein [53]. To obtain the correlation curve, simply measure the average volume of at least three globular proteins of known size (follow the steps described in Subheading 3.7) and plot their molecular weight and their measured average volume on the x- and y-axes of a correlation plot, respectively. The correlation curve can then be used to determine the expected volume for a protein of known molecular weight. However, this correlation applies only to samples prepared with the same deposition method and scanned with the same AFM setup and the same tip. References 1. Binnig G, Quate CF, Gerber C (1986) Atomic force microscope. Phys Rev Lett 56:930–933. https://doi.org/10.1103/PhysRevLett.56. 930 2. Binnig G, Rohrer H, Gerber C, Weibel E (1982) Surface studies by scanning tunneling microscopy. Phys Rev Lett 49:57–61. https:// doi.org/10.1103/PhysRevLett.49.57 3. Radmacher M, Tillamnn RW, Fritz M, Gaub HE (1992) From molecules to cells: imaging soft samples with the atomic force microscope. Science 257:1900–1905. https://doi.org/10. 1126/science.1411505

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AFM Analysis of ssDNA-Protein Complexes microscopy. In Scanning probe microscopy: atomic force microscopy and scanning tunneling microscopy. Springer, Berlin, Heidelberg, pp 187–204. https://doi.org/10.1007/9783-662-45240-0_14 50. Hafner JH, Cheung CL, Woolley AT, Lieber CM (2001) Structural and functional imaging with carbon nanotube AFM probes. Prog Biophys Mol Biol 77:73–110. https://doi.org/ 10.1016/s0079-6107(01)00011-6 51. Shlyakhtenko LS, Gilmore J, Portillo A et al (2007) Direct visualization of the EcoRIIDNA triple synaptic complex by atomic force

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microscopy. Biochemistry 46:11128–11136. https://doi.org/10.1021/bi701123u 52. Fuentes-Perez ME, Dillingham MS, MorenoHerrero F (2013) AFM volumetric methods for the characterization of proteins and nucleic acids. Methods 60:113–121. https://doi.org/ 10.1016/j.ymeth.2013.02.005 53. Schneider SW, Larmer J, Henderson RM, Oberleithner H (1998) Molecular weights of individual proteins correlate with molecular volumes measured by atomic force microscopy. Pflugers Arch 435:362–367. https://doi.org/ 10.1007/s004240050524

Chapter 16 Analysis of Mitochondrial SSB-DNA Complexes and Their Effects on DNA Polymerase γ Activity by Electron Microscopy and Enzymatic Assays Oya Bermek and Grzegorz L. Ciesielski Abstract The mitochondrial single-stranded DNA-binding protein (mtSSB) regulates the function of the mitochondrial DNA (mtDNA) replisome. In vitro, mtSSB stimulates the activity of enzymatic components of the replisome, namely mtDNA helicase and DNA polymerase gamma (Pol γ). We have demonstrated that the stimulatory properties of mtSSB result from its ability to organize the single-stranded DNA template in a specific manner. Here we present methods employing electron microscopy and enzymatic assays to characterize and classify the mtSSB-DNA complexes and their effects on the activity of Pol γ. Key words Mitochondrial DNA replication, Mitochondrial single-stranded DNA-binding protein, DNA polymerase γ, Electron microscopy, Nucleoprotein complexes

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Introduction Three proteins comprise the core of the human mitochondrial DNA replisome: DNA polymerase gamma (Pol γ), mitochondrial replicative DNA helicase (also known as Twinkle), and singlestranded DNA-binding protein (mtSSB) [1]. Together, these proteins facilitate rolling circle-like DNA synthesis in vitro [2]. Studies reported to date indicate that mtSSB serves as the major coordinator of the activity of the two remaining enzymes [1, 3, 4]. The critical role of mtSSB in the maintenance of the human mitochondrial genome has been confirmed recently by identification of pathogenic mutations in the corresponding gene, SSBP1 [5–7]. In vitro, mtSSB regulates DNA synthesis by stimulation of DNA template unwinding and nucleotide incorporation by mtDNA helicase and Pol γ, respectively [8–10]. In investigating the stimulatory properties of mtSSB, we have demonstrated that the efficiency of Pol γ

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_16, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Stimulation of human Pol γ activity upon increasing molar ratio of mtSSB and template DNA. The DNA polymerase assay was performed using 23 fmol (closed circles) or 230 fmol (open circles) of singly primed M13 DNA, 70 fmol of Pol γ holoenzyme, and increasing amounts of mtSSB, which remain at the indicated ratio to the amount of DNA template used. The data were normalized to the amount of nucleotide incorporated by Pol γ in the absence of mtSSB (arbitrarily set to 1 in each case). (This research was originally published in The Journal of Biological Chemistry. G.L. Ciesielski, O. Bermek, F.A. Rosado-Ruiz, S.L. Hovde, O.J. Neitzke, J.D. Griffith, L.S. Kaguni, Mitochondrial single-stranded DNA-binding proteins stimulate the activity of DNA polymerase γ by organization of the template DNA, J. Biol. Chem. 290 (2015) 28697–28707. © the American Society for Biochemistry and Molecular Biology)

activity depends on the molar ratio of mtSSB and template DNA (Fig. 1) [11]. This relationship is not linear; rather the activity of Pol γ varies upon addition of increments of mtSSB, indicating that a specific stoichiometry of mtSSB and template DNA modulates the optimal function of Pol γ. Considering that DNA binding is the major activity of mtSSB, this finding implies that the efficiency of Pol γ activity depends on the formation of specific mtSSB-DNA complexes. Notably, according to the strand displacement model of human mitochondrial DNA replication, the formation of such complexes would facilitate the synthesis of the lagging strand [12, 13]. We present an approach utilizing electron microscopy (EM) and DNA polymerase activity assay to characterize the mtSSB-DNA complexes and their effect on Pol γ activity [11]. To mimic the circular form of the mitochondrial genome, we employed the circular single-stranded M13 DNA as the DNA template. For EM analysis, the template DNA was incubated initially with specific concentrations of mtSSB. The structures formed were fixed by the addition of glutaraldehyde and the samples were chromatographed over an agarose matrix. Next, the samples were adsorbed to carbon supports and shadow-cast with tungsten. Using this approach, we observed that as the concentration of mtSSB in the

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Fig. 2 Electron microscopy of human mtSSB protein bound to the M13 DNA template. The numbers below individual images indicate the ratio of mtSSB tetramers per 100 nucleotides of template DNA. The dashed line indicates the concentration of SSB tetramers that would result in saturation of the template DNA molecules (as predicted from the ssDNA-binding site size of 35 nt/tetramer). The following species were distinguished: collapsed/beaded, collapsed, partially open, and fully open. (This research was originally published in The Journal of Biological Chemistry. G.L. Ciesielski, O. Bermek, F.A. Rosado-Ruiz, S.L. Hovde, O.J. Neitzke, J.D. Griffith, L.S. Kaguni, Mitochondrial single-stranded DNA-binding proteins stimulate the activity of DNA polymerase γ by organization of the template DNA, J. Biol. Chem. 290 (2015) 28697–28707. © the American Society for Biochemistry and Molecular Biology)

analyzed samples increases, the circular DNA template opens gradually (Fig. 2). At low (sub-saturating) concentrations of mtSSB, the template DNA forms predominantly dense and compacted structures, namely the beaded and the collapsed species. Excess mtSSB (saturating and oversaturating concentrations) promotes opening of the template DNA, through a series of partially open species. To identify the species that modulate the activity of Pol γ, we correlated the EM observations with the results of Pol γ activity assays, in which we applied the corresponding ratios of the M13 DNA template and mtSSB (Fig. 1). The template DNA was preincubated with mtSSB to allow complex formation before the addition of Pol γ. The activity of Pol γ was measured as the rate of incorporation of radiolabeled nucleotides into nascent DNA. Using this comparative approach, we determined that human Pol γ preferentially utilizes denser, partially opened template species, rather than the fully opened species. Furthermore, the sub-saturating concentrations of mtSSB have little effect on Pol γ activity, indicating that Pol γ is less efficient on templates containing uncoated stretches. Together, these findings indicate that specific organization of the single-stranded DNA template by mtSSB facilitates efficient DNA synthesis by Pol γ.

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Materials

2.1 Sample Preparation for EM

1. 1 M HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid), pH 7.6. 2. 1 M Magnesium chloride (MgCl2). 3. 1 M Potassium chloride (KCl). 4. 4 M Sodium chloride (NaCl). 5. Binding buffer: 10 mM HEPES, pH 7.6, 4 mM MgCl2, 30 mM KCl. 6. M13mp18 single-stranded circular DNA (7249 nucleotides). 7. 20 μM Human mtSSB, in 35 mM Tris–HCl pH 7.5, 150 mM NaCl, 2 mM EDTA, 5 mM DTT, 20% glycerol. 8. 25% Glutaraldehyde. 9. 1 M 2-Amino-2-(hydroxymethyl)propane-1,3-diol hydrochloride (Tris–HCl), pH 7.6. 10. 6% Agarose bead. 11. Agarose bead equilibration buffer: 10 mM Tris–HCl pH 7.6 and 0.1 mM EDTA. 12. 2.5 mM Spermidine hydrochloride (HCl). 13. Adsorption buffer: 0.125 mM Spermidine HCl, 2.5 mM KCl, 5 mM NaCl, and 0.1 mM MgCl2. 14. Carbon grids. 15. Denton high-vacuum evaporator.

2.2

EM Analysis

1. FEI Tecnai T12 electron microscope. 2. Gatan Digital Micrograph software.

2.3 Assay of Stimulation of Pol γ Activity by Human mtSSB

1. 200 nM Human Pol γ-α, in 30 mM Tris–HCl pH 7.5, 100 mM KCl, 2 mM EDTA, 5 mM β-mercaptoethanol, 20% glycerol. 2. 275 nM Human Pol γ-β, in 30 mM Tris–HCl pH 7.5, 100 mM NaCl, 1 mM EDTA, 5 mM β-mercaptoethanol, 20% glycerol. 3. 20 μM Human mtSSB, in 35 mM Tris–HCl pH 7.5, 150 mM NaCl, 2 mM EDTA, 5 mM DTT, 20% glycerol. 4. 125 nM Singly primed M13 DNA (6407 nt), in 10 mM Tris– HCl pH 8.0, 200 mM NaCl, 30 mM Na citrate, 8 mM EDTA [14]. 5. Bovine serum albumin (BSA). 6. 1 M Tris–HCl, pH 8.5. 7. 1 M MgCl2. 8. 1 M Dithiothreitol (DTT). 9. 2 M KCl.

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10. 100% Glycerol, anhydrous. 11. Dilution buffer: 30 mM Tris–HCl pH 7.5, 8% glycerol. 12. 5
γN buffer: 250 mM Tris–HCl pH 8.5, 20 mM MgCl2, 2 mg/mL BSA. 13. 50
Deoxynucleoside Triphosphates (dNTP mix): 1 mM dATP, 1 mM dTTP, 1 mM dGTP, 0.5 mM dCTP. 14. 10 μCi/μL α-32P dCTP. 15. 100% Trichloroacetic acid (TCA). 16. 100% Sodium pyrophosphate (NaPPi). 17. Stop solution: 10% TCA, 0.1 M NaPPi. 18. 12.1 M Hydrochloric acid (HCl). 19. Acid wash solution: 1 M HCl, 0.1 M NaPPi. 20. 95% Ethanol. 21. Glass-fiber filter paper. 22. 20 mL Glass vials.

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Methods

3.1 Sample Preparation for EM

1. Prepare a 50 μL sample mixture containing 0.425 nM ssM13mp18 DNA and desired concentration of mtSSB (100–600 ng) in the binding buffer (see Note 1). 2. Incubate the mixture for 20 min at room temperature. 3. Fix the protein to DNA by addition of glutaraldehyde to a final concentration of 0.6%, and incubate the mixture for 5 min at room temperature. 4. Quench the cross-linking with the addition of 2.5 μL of 1 M Tris–HCl, pH 7.6 (50 mM final). 5. Separate unbound protein by chromatography over 2 mL columns of 6% agarose beads equilibrated in 10 mM Tris–HCl pH 7.6 and 0.1 mM EDTA. 6. Mix 18.5 μL sample with 1.5 μL of 20
adsorption buffer and apply 5 μL to glow-charged carbon grids. 7. Dehydrate the sample gradually, air-dry, and rotary shadowcast with tungsten [15].

3.2

EM Analysis

1. Examine samples using EM at 40 kV. 2. Measure the contour lengths by capturing the images with a Gatan Orius CCD camera and using Gatan Digital Micrograph software. Consider 0.34 nm for one base pair.

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3. Count 200–500 molecules for each concentration to determine the classifications of the complexes. Average the results of four independent experiments. 4. Adjust the contrast in the images in Adobe Photoshop and arrange into panels. 3.3 Assay of Stimulation of Pol γ Activity by Human mtSSB

1. Adjust a water bath to 37  C. 2. Prepare a master reaction mix such that each 25 μL reaction contains 1
γN buffer, 10 mM DTT, 1
dNTPs, 2.4 nM M13 template, 2 μCi α-32P dCTP, and 15 mM KCl. 3. Dispense the mix into pre-chilled microcentrifuge tube(s) on ice. Adjust with water for a final volume of 25 μL taking into account the volumes after addition of the desired amounts of mtSSB and Pol γ. 4. Add the desired amount of mtSSB as 5 μL aliquots to the microcentrifuge tubes. Include a reaction with no mtSSB as a control (see Note 1). 5. Prepare dilutions of Pol γ-α and -β at 7.2 nM and 40 nM, respectively. Mix equal volumes of protein dilutions and dispense 10 μL into the tubes containing the master mix and mtSSB (see Note 2). 6. Incubate the tubes for 30 min at 37  C, and then transfer to ice. 7. Stop the reactions with 0.5 mL of stop solution and incubate on ice for 5 min. 8. Filter samples through glass-fiber filters. Wash the reaction tube twice with acid wash solution, and then wash the filtration funnel three times with acid wash solution and once with 95% ethanol. 9. Dry the filters under a heat lamp for 5 min, and then count in scintillation fluid. 10. Spot 1 μL of the reaction mix directly onto filters, dry, and count in scintillation fluid without filtration to calculate the specific radioactivity of the mix and determine nucleotide incorporation. 11. The DNA polymerase activity of Pol γ is measured as pmol of nucleotides incorporated into the nascent DNA strand. The molar amount of incorporated nucleotide is calculated as follows: Incorporated nucleotide ¼ average cpm of washed filters/ [(average cpm/μL of unwashed filters)/(pmol/μL of nucleotides spotted)]. 12. To obtain fold of stimulation, calculate the ratio of nucleotide incorporation in the samples containing mtSSB as compared to those without mtSSB.

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Notes 1. We have predicted the specific molar ratios of human mtSSB to DNA template from the ssDNA-binding site size of 35 nt/ mtSSB tetramer. By calculation, one mole of M13mp18 singlestranded DNA template (used in EM studies) requires 207 moles of mtSSB for saturation, and the singly primed M13 DNA template (used in enzymatic assays) will require 183 moles of mtSSB for saturation. 2. We use an ~6-fold molar excess of the Pol γ-β dimer over Pol γ-α to ensure complete reconstitution of Pol γ.

Acknowledgments G.L.C. and O.B. would like to acknowledge Dr. Laurie S. Kaguni and Dr. Jack D. Griffith, who mentored the authors in the described methodology during their postdoctoral training. G.L.C. was partially supported by a grant from the Auburn University at Montgomery Research Grant-in-Aid Program. References 1. Ciesielski GL, Oliveira MT, Kaguni LS (2016) Animal mitochondrial DNA replication. Enzyme 39:255–292 2. Korhonen JA, Pham XH, Pellegrini M, Falkenberg M (2004) Reconstitution of a minimal mtDNA replisome in vitro. EMBO J 23:2423–2429 3. Cerro´n F, de Lorenzo S, Lemishko KM, Ciesielski GL, Kaguni LS, Cao FJ, Ibarra B (2019) Replicative DNA polymerases promote active displacement of SSB proteins during lagging strand synthesis. Nucleic Acids Res 47:5723–5734 4. Morin JA, Cerro´n F, Jarillo J, Beltran-HerediaE, Ciesielski GL, Arias-Gonzalez JR, Kaguni LS, Cao FJ, Ibarra B (2017) DNA synthesis determines the binding mode of the human mitochondrial single-stranded DNA-binding protein. Nucleic Acids Res 45:7237–7248 5. Piro-Me´gy C, Sarzi E, Tarre´s-Sole´ A, Pe´quignot M, Hensen F, Quile`s M, Manes G, Chakraborty A, Se´ne´chal A, Bocquet B, Cazevieille C, Roubertie A, Mu¨ller A, Charif M, Goudene`ge D, Lenaers G, Wilhelm H, Kellner U, Weisschuh N, Wissinger B, Zanlonghi X, Hamel C, Spelbrink JN, Sola` M, Delettre C (2020) Dominant mutations in mtDNA maintenance gene

SSBP1 cause optic atrophy and foveopathy. J Clin Invest 130(1):143–156 6. Gustafson MA, McCormick EM, Perera L, Longley MJ, Bai R, Kong J, Dulik M, Shen L, Goldstein AC, McCormack SE, Laskin BL, Leroy BP, Ortiz-Gonzalez XR, Ellington MG, Copeland WC, Falk MJ (2019) Mitochondrial single-stranded DNA binding protein novel de novo SSBP1 mutation in a child with single large-scale mtDNA deletion (SLSMD) clinically manifesting as Pearson, Kearns-Sayre, and Leigh syndromes. PLoS One 14: e0221829 7. Del Dotto V, Ullah F, Di Meo I, Magini P, Gusic M, Maresca A, Caporali L, Palombo F, Tagliavini F, Baugh EH, Macao B, Szilagyi Z, Pe´ron C, Gustafson MA, Khan K, La Morgia C, Barboni P, Carbonelli M, Valentino ML, Liguori R, Shashi V, Sullivan JA, Nagaraj S, El-Dairi M, Iannaccone A, Cutcutache I, Bertini E, Carrozzo R, Emma F, Diomedi-Camassei F, Zanna C, Armstrong M, Page MJ, Boesch S, Wortmann SB, Kopajtich R, Stong N, Sperl W, Davis E, Copeland WC, Seri M, Falkenberg M, Prokisch H, Katsanis N, Tiranti V, Pippucci T, Carelli V (2020) SSBP1 mutations cause mtDNA depletion underlying a complex optic

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atrophy disorder. J Clin Invest 130 (1):108–125 8. Korhonen JA, Gaspari M, Falkenberg M (2003) TWINKLE Has 50 -> 30 DNA helicase activity and is specifically stimulated by mitochondrial single-stranded DNA-binding protein. J Biol Chem 278:48627–48632 9. Oliveira MT, Kaguni LS (2010) Functional roles of the N- and C-terminal regions of the human mitochondrial single-stranded DNA-binding protein. PLoS One 5:e15379 10. Oliveira MT, Kaguni LS (2011) Reduced stimulation of recombinant DNA polymerase γ and mitochondrial DNA (mtDNA) helicase by variants of mitochondrial single-stranded DNA-binding protein (mtSSB) correlates with defects in mtDNA replication in animal cells. J Biol Chem 286:40649–40658 11. Ciesielski GL, Bermek O, Rosado-Ruiz FA, Hovde SL, Neitzke OJ, Griffith JD, Kaguni LS (2015) Mitochondrial single-stranded DNA-binding proteins stimulate the activity

of DNA polymerase γ by organization of the template DNA. J Biol Chem 290:28697–28707 12. Clayton DA (1982) Replication of animal mitochondrial DNA. Cell 28:693–705 13. Miralles Fuste´ J, Shi Y, Wanrooij S, Zhu X, ¨ , Sabouri N, Gustafsson Jemt E, Persson O CM, Falkenberg M (2014) In vivo occupancy of mitochondrial single-stranded DNA binding protein supports the strand displacement mode of DNA replication. PLoS Genet 10:e1004832 14. Farr CL, Matsushima Y, Lagina AT, Luo N, Kaguni LS (2004) Physiological and biochemical defects in functional interactions of mitochondrial DNA polymerase and DNA-binding mutants of single-stranded DNA-binding protein. J Biol Chem 279:17047–17053 15. Griffith JD, Christiansen G (1978) Electron microscope visualization of chromatin and other DNA-protein complexes. Annu Rev Biophys Bioeng 7:19–35

Chapter 17 Optical Tweezers to Investigate the Structure and Energetics of Single-Stranded DNA-Binding Protein-DNA Complexes Jose´ A. Morin, Fernando Cerro´n, Francisco J. Cao-Garcı´a, and Borja Ibarra Abstract Optical tweezers enable the isolation and mechanical manipulation of individual nucleoprotein complexes. Here, we describe how to use this technique to interrogate the mechanical properties of individual proteinDNA complexes and extract information about their overall structural organization. Key words Optical tweezers, Single-molecule manipulation, SSB, Single-stranded DNA, ProteinDNA organization

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Introduction Optical tweezers is a versatile single-molecule manipulation technique that is being used to investigate the mechanical properties and dynamics of operation of an increasing number of biochemical and biophysical processes [1–4]. The physical principles behind optical trapping and the instrumentation necessary to build standard optical tweezers have been well established [5–7]. Optical tweezers rely on the property of light to exert force on matter. The magnitude of optical forces allows trapping dielectric micronsized polystyrene beads. These beads are biochemically linked to the molecules of interest, and they serve in this way as handles to a biological system. Mechanical manipulation of the system under study requires attaching it to a second attachment point (such as a second bead held atop a micropipette in the experiments described in this chapter, Fig. 4). In this way, the system can be stretched by moving the bead in the optical trap relative to the bead fixed on top of the micropipette. Typically, optical traps can generate forces on the system under study from ~0.5 to 150 piconewtons (pN).

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_17, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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In the last few years, the optical tweezers technique has proven useful to investigate the site size, equilibrium constants, and energetics of the binding of various single-stranded DNA (ssDNA)binding proteins (SSBs) to ssDNA [8–14]. SSBs are ubiquitous and essential components of every DNA metabolic transaction requiring single-stranded intermediates. They bind to ssDNA with high affinity in a sequence-independent manner, where they perform several functions [15]. The vast majority of SSB proteins interact in vitro with a variable number of ssDNA nucleotides depending on the salt concentration and type, as well as protein-binding density on the ssDNA [16]. These different binding modes have different biochemical and structural properties and might be used selectively for different functions [13, 16]. Determination of the structure and/or organization of large SSB-DNA complexes, such as those formed during DNA replication (Okazaki fragments may range from ~100 to ~1000 nucleotides), is challenging due to the high flexibility and dynamics of these nucleoprotein complexes. Here, we show how to use optical tweezers to interrogate the structure and energetics of different binding modes of human mitochondrial SSB (HmtSSB) to long, preformed ssDNA molecules and to determine the dominant binding mode when protein binding is coupled to DNA replication. In a typical assay, the interaction of HmtSSB with a single ssDNA molecule held at a constant force between the beads will compact the ssDNA polymer, decreasing its end-to-end distance. Upon protein binding, stretching of the resulting SSB-ssDNA complex will reveal its mechanical properties, from which the actual proteinDNA organization can be extracted using models of polymer physics. Overall, our results [13] showed that HmtSSB binds to preformed ssDNA in two main modes depending on protein and salt concentration; each mode organizes either ~30 or ~50 ssDNA nucleotides per tetramer. However, when binding of HmtSSB to ssDNA is coupled to DNA synthesis, the low-site-size binding mode (~30 nt/tetramer) is selected under all experimental conditions, suggesting this as the relevant mode for DNA replication. For both binding modes, the average Gibbs energy required to unwrap one nucleotide from each HmtSSB tetramer (ΔGw) decreased linearly with NaCl concentrations from ΔGw ~ 0.7 kBT/nt at 10 mM NaCl to ~0.4 kBT/nt at 300 mM NaCl.

2

Materials Prepare all stock solutions using ultrapure water and store at room temperature unless otherwise indicated.

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1. Polystyrene beads of 3.18 μm diameter functionalized with protein G. 2. Polystyrene beads of 2 μm diameter functionalized with streptavidin. 3. PBS 7.4: 140 μM NaCl, 2.7 mM KCl, 81 mM K2HPO4, 19 mM KH2PO4. Sterilize with autoclave and filter with 0.2 μm polypropylene filter. 4. Anti-digoxigenin antibody: Dissolve 200 μg of antidigoxigenin antibody in 200 μL PBS 7.4. Store at
20  C. 5. Cross-linking buffer: 100 mM Na2HPO4, 100 mM NaCl. Sterilize with autoclave and filter with 0.2 μm polypropylene filter. 6. Dimethyl pimelimidate (DMP): 20 mg DMP into 400 μL of cross-linking buffer. 7. Eppendorf vortex mixer.

2.2

Fluidic Chambers

1. Laser engraver. 2. 24  60 mm #2 Coverslips. 3. Parafilm tape. 4. To make micropipettes use glass capillaries of 40 by 80 μm (internal–external diameters, respectively). Use a pipette puller to pull and heat simultaneously individual glass capillaries (http://tweezerslab.unipr.it for more details). The aperture of the micropipette at the tip should be 0.5–1 μm. 5. Bypassing tubes (or dispensers): Use glass capillaries of 25–100 μm internal–external diameters. 6. Standard heating plate.

2.3

Reaction Buffers

1. Binding mode reaction buffer: 10 or 100 nM HmtSSB diluted in 50 mM Tris–HCl pH 7.5, 2 mM dithiothreitol (DTT), and desired concentrations of NaCl and MgCl2. 2. DNA synthesis reaction buffer, 5, 50, 100, or 200 nM HmtSSB diluted in 50 mM Tris–HCl pH 7.5, 2 mM DTT, 50 mM NaCl, 4 mM MgCl2, 50 μM dNTPs, 2 nM Phi29 DNA polymerase (Phi29Pol). 3. Phi29Pol reaction buffer: 50 mM Tris–HCl pH 7.5, 2 mM DTT, 50 mM NaCl, 4 mM MgCl2, 50 μM dNTPs.

2.4 Preparation of DNA Hairpins

1. PCR tubes. 2. 10 mM dNTP mix. 3. 10 PCR buffer. 4. Microcentrifuge tubes. 5. Thermo-shaker.

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6. DNA purification kit. 7. 0.5 μg/μL and 0.1 μg/μL pUC18 DNA vector. 8. 1 μg/mL pUC19 DNA vector. 9. 20 U/μL SalI restriction enzyme. 10. 20 U/μL EcoRI restriction enzyme. 11. 20 U/μL Bam HI-HF restriction enzyme. 12. 20 U/μL PstI restriction enzyme. 13. 400 U/μL T4 DNA ligase. 14. 5 U/μL Taq DNA polymerase. 15. Thermal cycler. 16. 10 mM each dATP, dCTP, dTTP, dGTP, and digoxigeninlabeled dUTP. 17. Oligonucleotide primers for preparation of digoxigenin-labeled dsDNA handles: 50 - CCTCTGACACATGCAGCTCC -30 and 50 -CGCGGCCTTTTTACGGTTCC-30 . 18. Oligonucleotide primers for unwinding segment: 50 - TCTTT AACGGCAACGTAATCAGGAT -30 and 50 - CGATAATGG TATTACTCTTTGGCATGTCGA-30 . 19. Oligonucleotide primers for preparation of linker DNA segment: 50 -biotin-(dT)20 CAATCACTTCAGGTAGCATC -30 and 50 (P)- AATTGATGCTACCTGAAGTGATTGGCT GATGCA-30 . 20. Self-annealing oligonucleotide: 50 - TCGAGCAGATGCAG CAATAACGTGCATCTGC-30 . 21. Self-annealing oligonucleotide containing one abasic site: 50 -T CGAGCAGATGCAGCAA[abasic]TAACGTGCATCTGC-30 . 22. Oligonucleotide complementary to the apex of the hairpin: 50 GCCGATGCACGTTATTCGTGCATCGGCTCG-30 .

3

Methods

3.1 Preparation of Fluidic Chambers for Optical Tweezers Studies

Counter-propagating optical tweezers employ the light momentum conservation principle to calculate forces accurately [17]. This method guarantees calibration independently from parameters difficult to estimate accurately (i.e., buffer viscosity, bead radius, and temperature) and from the laser power. Figure 1 shows a schematic representation of the counter-propagating optical tweezers setup used for the experiments described in this chapter (Mini-OT). Details of construction, calibration, and operation of the MiniOT device are available at the “Tweezers Lab” website (http:// tweezerslab.unipr.it).

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Fig. 1 Schematics of the optical tweezers setup. Two counter-propagating lasers (green and yellow lines) are focused inside the flow chamber using high numerical aperture objectives to form optical traps. The light leaving from each trap is collected by a second objective and sent to a position-sensitive detector (PSD) to measure forces (by measuring changes in light momentum [17]). Beam splitters are used to deviate ~5% of the incoming light of each laser to a second set of PSDs, which are used to monitor the optical trap position. The position of each laser is controlled by piezo actuators (wigglers). A CCD camera and a blue LED light (blue line) are used to visualize the interior of the flow cell

Follow the next steps for the fabrication of a fluidic chamber compatible with a Mini-OT device: 1. Drill three 2 mm diameter holes at each side of a clean 24  60 mm coverslip #2 by using a laser engraver, as shown in Fig. 2. 2. Use the laser engraver to cut three slits (2 mm width) into the parafilm tape as shown in Fig. 2. 3. Make a fluidic chamber by sandwiching two coverslips (modified as in step 1) with two layers of the parafilm (modified as in step 2). The sandwich should include a micropipette in the middle and two bypass tubes as shown in Fig. 2. 4. Place the chamber on top of a heating plate preheated to 80  C and wait until the parafilm becomes transparent (3–4 min). During this step, the two parafilm layers fuse creating three channels connected by bypass tubes (see Fig. 2). Then, remove chamber from the heating plate using tweezers (see Note 1). 5. Mount the fluidic chamber onto a metal frame of the Mini-OT by matching the fluidic holes.

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Fig. 2 Schematic representation of the flow chamber. Use two layers of parafilm (engraved as described in the main text) and two coverslips (one of them containing three holes at each side) to form a sandwich including in between the parafilm layers, the micropipette, and the dispenser tubes, as depicted in the figure. The top and bottom channels of the flow chamber contain functionalized micron-size beads (yellow and blue spheres), which are flown to the central channel through short dispenser tubes. The direction of the main flow in the central channel (black arrows) prevents the beads from invading the micropipette position (where experiments are carried out). The optical trap (red cross) is used to trap and manipulate the beads in order to assemble the desired experimental geometry (see Fig. 2 in this volume’s Chapter 18). Additional dispenser tubes can be used to flow proteins and/or buffers to the central channel

6. Fill the top channel with a solution containing DNA-bead preparation (see below), the middle channel with reaction buffer, and the bottom channel with a solution containing 1 μL of streptavidin-covered beads diluted in 300 μL of reaction buffer. 3.2 Preparation of Functionalized Polystyrene Beads

1. Add 750 μL of protein G-coated polystyrene beads to a 1.5 mL microcentrifuge tube and centrifuge for 6 min at 1000 RCF. Discard supernatant. 2. Dilute the pellet with 750 μL of cross-linking buffer. 3. Add 22.5 μL of 50 μg/μL DMP and 45 μL of 1 μg/μL antidigoxigenin antibody, mix thoroughly, and incubate for 1 h at room temperature in a microcentrifuge tube shaker set at 110 RCF. The cross-linking reaction will ensure covalent bonds between the anti-digoxigenin antibody- and the protein G-covered polystyrene beads. 4. Centrifuge for 6 min at 1000 RCF. Discard supernatant. 5. Add 750 μL of 1 M Tris–HCl pH 7.5 and incubate for 2 h in a microcentrifuge tube shaker set at 110 RCF to quench the cross-linking reaction. 6. Divide the total volume of the sample into two different 1.5 mL microcentrifuge tubes and add 750 μL of ultrapure water in each tube (see Note 2).

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Fig. 3 Diagram of DNA hairpin construction. The assembly of a DNA hairpin suitable for manipulation in the optical tweezers is carried out in two consecutive ligation steps: In Ligation I, one end of the unwinding segment (red) is ligated to a self-complementary DNA oligonucleotide (DNA loop, green) and the other end to a short dsDNA linker (light blue) labeled with biotin (yellow) at the 50 end. In Ligation II, the product of Ligation I is ligated to one end of a ~2664 bp dsDNA spacer labeled with a digoxigenin DNA handle at the other end

7. Centrifuge for 6 min at 1000 RCF, discard supernatant, and elute the bead pellet in 750 μL of PBS 7.4. Repeat this step three times (see Note 3). 8. Mix the content of the two tubes in a single tube and centrifuge again for 6 min at 1000 RCF. Discard supernatant. 9. Elute in 1 mL PBS 7.4, make 50 mL aliquots, and store at 4  C. 3.3 Preparation of DNA Hairpin

3.3.1 Preparation of the Unwinding Segment

To study the mechanical properties of the HmtSSB-ssDNA complexes, individual ssDNA molecules were generated in the optical tweezers upon mechanical unwinding of DNA hairpins (Figs. 3 and 4). 1. In a 250 μL PCR tube, mix 1 μL of 10 mM dNTP mix with 2 μL each of 10 mM PCR oligonucleotide primers, 5 μL of 10 PCR buffer, 1 μL of 0.5 ng/μL Phi29 DNA, and 0.5 μL of 5 U/μL Taq DNA polymerase and fill to 50 μL with ultrapure water. 2. Configure the thermal cycler as follows: step 1: 3 min, 94  C; step 2: 1 min, 94  C; step 3: 1 min, 57  C; step 4: 2.5 min, 72  C; repeat steps 2–4 30 times; step 5: 10 min at 72  C. 3. Purify DNA using a commercial DNA purification column and elute in a final volume of 30 μL. 4. Digest 500 ng of the PCR product with 1.5 μL of 20 U/μL EcoRI in a final reaction volume of 30 μL containing 3 μL of 10 EcoRI restriction buffer for 2 h at 37  C. 5. Purify the DNA in a final volume of 30 μL using a commercial DNA purification column. 6. Digest the purified DNA with 1.5 μL of 20 U/μL SalI in a final reaction volume of 50 μL containing 5 μL of 10 SalI restriction buffer for 2 h at 37  C. 7. Purify the unwinding segment digested with EcoRI and SalI in a final volume of 30 μL using a commercial DNA purification column.

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Fig. 4 Schematics of the experimental setup to measure SSB-binding modes of preformed ssDNA molecules. The complementary strands of the DNA hairpin construct are tethered between a bead in the optical trap (brown) and a bead on top of the micropipette (blue). Mechanical unwinding of the DNA hairpin in the presence of a 30 mer oligonucleotide (blue line) complementary to the apex of the loop prevents re-winding of the hairpin at low forces, providing a method to obtain long ssDNA segments in the optical tweezers. After washing out the oligonucleotide, the SSB protein (brown tetramer) is introduced inside the flow cell. At a constant force, organization and compaction of ssDNA by SSB binding are followed by a gradual decrease in the distance between the beads (Δx). The plot in the right shows the consecutive force-extension curves (FECs) and distance changes obtained during the course of the experiment. Gray line: Initial FEC of the mechanical unwinding of the unzipping segment. Red line: FEC of the resulting DNA molecule upon total opening of the hairpin and annealing of the oligonucleotide at the apex. Green line: Change in distance between the beads due to SSB binding and ssDNA compaction when force is held constant at ~6 pN. Purple line: Final FEC of the SSB-DNA complex. Dotted blue line represents the fit to the reversible section of the FEC of the SSB-DNA complex with Eq. (1) 3.3.2 Preparation of Digoxigenin-Labeled dsDNA Handles

1. Preparation of dNTP-DIG mix: Mix in a 1.5 mL microcentrifuge tube 6 μL of dATP, 6 μL of dCTP, 6 μL of dGTP, and 3 μL dTTP (all dNTPs at 10 mM) with 3 μL of digoxigenin-labeled dUTP (DIG-dUTP 10 mM). Fill up to 75 μL with ultrapure water. 2. In a 250 μL PCR tube, mix 25 μL of the dNTP-DIG mix with 2 μL each of 1 mM PCR oligonucleotide primers, 10 μL of 10 PCR buffer, 1 μL of 0.1 μg/μL pUC18 DNA, and 2 μL of 5 U/μL Taq DNA polymerase and fill to 100 μL with ultrapure water. 3. Configure the thermal cycler as follows: step 1: 3 min, 94  C; step 2: 1 min, 94  C; step 3: 1 min, 60  C; step 4: 2 min, 72  C; repeat steps 2–4 29 times; step 5: 10 min at 72  C. 4. Purify DNA using a commercial DNA purification column and elute in a final volume of 30 μL. 5. Add 4 μL of 10 BamHI restriction buffer, 1 μL of 20 U/μL BamHI, and 5 mL of ultrapure water and digest for 2 h at 37  C.

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6. Purify the DNA in a final volume of 30 μL using a commercial DNA purification column. 7. Optionally, the resulting DNA products can be checked by standard agarose gel electrophoresis. 3.3.3 Preparation of dsDNA Spacer

A DNA segment of 2664 bp (pUC19 DNA vector) was used as a spacer to separate the two beads and to facilitate the manipulation and identification of single-DNA hairpins. 1. Digest 600 ng of the pUC19 DNA vector with 1.5 μL of 20 U/μL BamHI in a final volume of 30 μL containing 3 μL of BamHI 10 reaction buffer for 1 h at 37  C. 2. Purify the DNA in a final volume of 30 μL using a commercial DNA purification column. 3. Digest the purified DNA with 1.5 μL of 20 U/μL PstI in a final reaction volume of 50 μL containing 5 μL of 10 PstI restriction buffer for 2 h at 37  C. 4. Purify the BamHI-PstI pUC19 DNA in a final volume of 30 μL using a commercial DNA purification column.

3.3.4 Preparation of a Linker DNA Segment

Prepare the DNA linker by annealing two partially complementary DNA oligonucleotides (see Subheading 2.3). The DNA product of the annealing reaction will contain two distinct ends (Fig. 3). One end contains a 50 protruding terminus complementary to EcoRI restriction sequence. The other end contains a 30 protruding terminus complementary to PstI restriction sequence and a (polydT)20 50 tail labeled with biotin. 1. Mix in a 1.5 mL microcentrifuge tube equimolar concentrations of the two partially complementary oligonucleotides and take the reaction to a final volume of 100 μL with annealing buffer. The recommended stock concentration of oligonucleotides is 100 μM. 2. Heat to 95  C for 10 min in a thermo-shaker, switch off the heating, and wait until the block reaches room temperature (~2 h). 3. Dilute the sample 1/500 and store at
20  C in 20 μL aliquots.

3.3.5 Preparation of a DNA Loop for Hairpin Apex

1. Dilute a self-annealing DNA oligonucleotide (see Note 7) to a final concentration of 100 ng in annealing buffer. 2. Heat to 95  C for 10 min in a thermo-shaker, switch off the heating, and wait until the block reaches room temperature (~2 h). 3. Dilute the sample further 1/10 and store at
20  C in 20 μL aliquots.

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3.3.6 Preparation of Final DNA Hairpin Constructs

Assemble the final hairpin constructs using two consecutive ligation steps (Ligation I and Ligation II, Fig. 3). 1. For Ligation I, mix in a 1.5 mL microcentrifuge tube 230 ng of the unwinding segment digested with BamHI and EcoRI (Subheading 3.3.1), 1.5 μL of linker DNA (Subheading 3.3.4), 2 μL of DNA loop (Subheading 3.3.5), 3 μL of 10 T4 reaction buffer, and 1.5 μL of T4 DNA ligase and fill to 30 μL with ultrapure water. 2. Incubate overnight at 16  C. 3. Purify DNA using a commercial DNA purification column. Elute DNA in 30 μL of water. 4. Store Ligation I at 4  C for weekly use or store at
20  C for future use. 5. For Ligation II, mix in a 1.5 mL microcentrifuge tube 15 μL of Ligation I, 7 ng of DNA spacer (Subheading 3.3.3), 0.5 μL of digoxigenin-labeled DNA handles (Subheading 3.3.2), 3 μL of 10 T4 reaction buffer, and 1.5 μL of T4 DNA ligase and fill to 30 μL with ultrapure water. 6. Incubate overnight at 16  C. 7. Purify DNA using a commercial DNA purification column and elute DNA in 30 μL of water. 8. Store Ligation II at 4  C for weekly use or store at
20  C for future use.

3.4 Generation and Manipulation of Single ssDNA Molecules in the Optical Tweezers

The first step to a successful optical tweezers experiment is to generate a tether. To this end, the anchor points of the molecule (biotin and digoxigenin, Figs. 3 and 4) have to be linked to plastic microspheres functionalized with the complementary chemical groups. The link between digoxigenin and anti-digoxigenin is made by preincubation of the construct with anti-digoxigenincoated beads, while the link between biotin and streptavidin-coated beads is done at the optical tweezers (see below).

3.4.1 Incubation of the DNA Hairpin Construct with Anti-digoxigenin-Coated Beads

1. In a test tube, mix 3 μL of the DNA hairpin construct obtained after the Ligation II step with 3 μL of anti-digoxigenin-coated beads and 3 μL of reaction buffer for 10 min at room temperature (see Note 4). 2. Dilute the mixture to 300 μL with the reaction buffer (see Note 5).

3.4.2 Generation of Single-DNA Hairpin Tethers in the Optical Tweezers

To generate a tether, prepare optical tweezers and fluidic chamber as described above (Subheadings 3.1–3.3). A schematic representation of single tether generation is shown in Fig. 2 of this volume’s Chapter 18.

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1. Use the optical trap to trap a streptavidin-coated bead delivered by the corresponding dispenser tube. 2. Place the bead on top of the micropipette by air suction. 3. Use the optical trap to trap a bead coated with the DNA hairpin construct and bring it closer to the streptavidin-coated bead. 4. Reset force sensor to 0 piconewtons (pN) (see Note 6). 5. Approach and retract the pipette from the trapped bead until an increase in force along the y-axis is observed (see Note 7). Attachment of a DNA molecule between the two beads will be monitored from the motion of the bead in the optical trap as the two beads are pulled apart. 3.4.3 Identification of Individual DNA Hairpins

A single hairpin presents well-defined mechanical properties, including a force-extension signal corresponding to the stretching of the dsDNA handle (see Note 8) and a sawtooth pattern that reflects the mechanical unzipping of the hairpin (Fig. 4). For an individual hairpin, mechanical unzipping occurs at an average force of ~15 pN (Fig. 4). In the case of a double tether, unzipping occurs at twice the expected force. The mechanical unzipping process is fully reversible (see Note 9).

3.4.4 Generation of ssDNA in the Optical Tweezers

Figure 4 shows a schematic representation of the experimental procedure. 1. Ensure that a single-DNA hairpin is tethered between the beads by visually inspecting its force-extension curve. 2. Unzip the molecule and keep the force constant at ~20 pN (hairpin fully unzipped). 3. Flow in a 250 nM solution of an oligonucleotide complementary to the region at the apex of the hairpin. Hybridization of the oligonucleotide at the apex generates a sizeable kinetic barrier to hairpin reannealing when tension is released. 4. Release the tension below the reannealing force. The signature of a successful hybridization is the loss of reversibility in the hairpin reannealing process. 5. Wash out the oligonucleotide by flowing the appropriate reaction buffer into the central channel of the flow chamber.

3.5 Mechanical Characterization of SSB-DNA Complexes 3.5.1 Preparation of SSB Solution

The following procedures were optimized to determine the human mitochondrial SSB-binding properties and can be adapted to study other SSB proteins. 1. Dilute SSB in 500 μL of reaction buffer to the final concentration at which its ssDNA-binding properties are studied.

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2. Filter the solution with a 0.22 μm nylon filter to eliminate possible protein aggregates and/or bulky contaminants. 3. Flow the protein solution into the dispenser tube connected to the central channel of the flow chamber (Fig. 2). 3.5.2 Determination of Force-Extension Curves of SSB-ssDNA Complexes

Figure 4 shows a schematic representation of the experimental procedure. 1. Upon isolating a single ssDNA molecule between the beads, keep a constant force on the system (usually below 3 pN), and flow the SSB solution into the central channel of the flow chamber. 2. SSB binding and organization of the ssDNA will decrease the extension between the beads. 3. After the binding reaction is accomplished (no further decrease in extension), flow the reaction buffer into the middle channel to wash free SSB. 4. Pull the protein-DNA complex at a constant rate to obtain its force-extension curve (see Note 10). 1. Calculate the area between the SSB-free and SSB-bound forceextension curves (see Note 11).

3.6 Calculation of the Energy to Unwrap a Single Nucleotide from the SSB Tetramer

2. Divide the area by the total number of nucleotides of the ssDNA (see Note 12).

3.7 Determination of the SSB-Binding Mode to ssDNA

1. Fit the reversible section of the force-extension curve of the SSB-ssDNA complex (see Fig. 4, Note 13) to the following equation for the extension per nucleotide [13, 18]: L x complex ðF Þ ¼ ð1
c Þ x ssDNA ðF Þ þ c ends m     F L ends kB T  coth ,
F L ends kB T

ð1Þ

where c is the fraction of ssDNA nucleotides bound to the SSB tetramer, xssDNA(F) is the force-extension curve of the free ssDNA measured experimentally, m is the number of nucleotides bound per SSB, and Lends is the effective distance between the nucleotides bound to the SSB (kB is the Boltzmann constant and T is the temperature in Kelvin). 2. By fixing the value of Lends to the crystallographic length of a SSB tetramer, the fit gives the coverage, c, and the number of monomers bound per ligand, m (see Note 14).

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Fig. 5 Schematics of the experimental setup to measure the co-replicational binding modes of SBB to ssDNA. Use the optical tweezers to attach between the beads a single-DNA hairpin construct containing an abasic site at the apex. Keep force constant at ~3–4 pN and flow the Phi29 DNA polymerase (green triangle) and the desired concentration of SSB (brown tetramer). As strand displacement synthesis proceeds, the displaced ssDNA strand is gradually covered by the SSB. DNA synthesis terminates at the abasic site, yielding a hybrid DNA molecule with 5226 bp of dsDNA and 2540 nt of ssDNA gradually covered by the SSB. The plot shows the consecutive force-extension curves (FECs) and distance changes obtained during the course of the experiment. Gray line: Initial FEC of the mechanical unwinding of the unzipping segment. Green line: Change in distance between the beads as the DNA polymerase elongates the dsDNA handle and displaces the complementary ssDNA, which is gradually covered by SSB (constant force ~4.5 pN). Purple line: Final FEC of the SSB-DNA complex. Dashed blue line corresponds to the wormlike chain model for a hybrid DNA molecule containing 5226 bp of dsDNA (persistence length P ¼ 53 nm, stretch modulus S ¼ 1200 pN/nm) and 2540 nt of free ssDNA (P ¼ 0.75 nm, S ¼ 800 pN/nm) 3.8 Determination of SSB-Binding Mode During DNA Replication

Figure 5 shows a schematic representation of the experimental procedure. 1. Dilute Phi29 DNA polymerase to 2 nM and SSB to the desired final concentration in the Phi29Pol reaction buffer. 2. Use the optical tweezers to isolate a single-DNA hairpin as described in Subheading 3.4. In this case, the selfcomplementary oligonucleotide that forms the hairpin’s apex contains an abasic site to stop replication. 3. Keep the force constant on the hairpin at ~3 pN (hairpin is closed) using the force feedback mode of the optical tweezers. 4. Flow the polymerase-SSB solution into the central channel of the flow chamber. 5. The combined effect of DNA replication and SSB binding will be detected in real time as a gradual increase in the distance between the beads (Fig. 5). 6. Wash out protein excess by flowing Phi29Pol reaction buffer (without dNTPs) into the central channel of the flow chamber. 7. Pull the protein-DNA complex at a constant rate in order to obtain its force-extension curve (Fig. 5).

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8. After subtraction of the dsDNA contribution, fit with Eq. (1) the equilibrium force-extension curves of the protein-DNA complexes to obtain the values of the average number of wrapped nucleotides per tetramer and the fraction of ssDNA covered by the SSB under these conditions (as described in Subheading 3.7).

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Notes 1. Application of gentle pressure on the coverslip helps the sealing process. Note that too much pressure will break the coverslips and/or foster the progression of the melted parafilm layer too deep into the channel (which in turn may block the dispenser tubes and the pipette tip). 2. Water is used to dilute the high concentration of Tris–HCl and to facilitate precipitation of the beads onto the bottom of the test tubes during centrifugation. 3. During centrifugation steps a significant amount of beads will be bound to and detach easily from the walls of the tube. Remove the supernatant carefully. 4. The DNA:bead ratio should be optimized based on the success to obtain single tethers in the optical tweezers. Note that for each DNA-bead preparation there are fluctuations in the final concentration, which makes the optimum ratio to change slightly. 5. The anti-digoxigenin-coated bead dilution ratio should be adjusted based on the density of beads in the corresponding channel in the flow cell. Note that for each anti-digoxigenincoated bead preparation there are fluctuations in the final concentration. An over-diluted preparation will result in unnecessary waiting times to capture a bead, while an overconcentrated situation will result in difficulties to capture single beads. 6. Zeroing the detectors is a sensitive step; take care that no external flows are acting on the trapped bead, and separate the two beads ensuring that there are no tethers between them. 7. The sensitive detection of the bead position in the trap and, by extension, the calibrated measurement of displacement and force are determined by back-focal-plane interferometry. 8. Contour length is calculated as Lbp*Nbp_handle, where Lbp is the length of a base pair (0.34 nm) and Nbp_handle ¼ 2664 bp is the number of base pairs of the dsDNA. 9. The force at which mechanical unzipping of the hairpin occurs depends on the ionic strength of the solvent. The characteristic sawtooth pattern that reflects the mechanical unzipping is

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highly correlated with the underling base pair sequence of the hairpin. Once the hairpin is fully unzipped, the resulting molecules have a characteristic length given by (in this case) Lbp*Nbp_handle + Lnt*(2*Nbp_hairpin) ¼ 3.84 μm, where Lnt ~ 0.58 nm is the average distance between two consecutive ssDNA nucleotides at 15 pN [19] and Nbp_hairpin ¼ 2540 bp is the number base pairs of the unzipping segment. 10. In our study, a pulling rate of 200 nm/s was used. 11. Areas were calculated numerically using the trapezoidal rule. The relaxation curve was used instead of the stretching free ssDNA curve to avoid the energetic contribution of long-range ssDNA secondary structure induced by salt. 12. The average work to unwrap one nucleotide from each HmtSSB tetramer, ΔGw, in pN·nm, was converted to kBT units (Boltzmann constant multiplied by the absolute temperature) using the relationship kBT ¼ 4.11 pN·nm at 25  C, which corresponds to the energy contribution from thermal fluctuations. Ideally, the average work should be corrected by the protein coverage value (calculated from Eq. 1). 13. To determine the reversible section of the force-extension curve of the SSB-ssDNA complex, stretch and relax the complex in consecutive cycles (starting from 0 pN). In each cycle, reach a slightly higher force (~1 pN) than in the previous one. The reversible section corresponds to that in which the pulling and relaxing force-extension curves overlap. 14. Usually Lends cannot be fitted independently of m as the bracket in Eq. (1) goes to 1 for F  kBT/ Lends (  4 pN nm/8 nm  0.5 pN).

Acknowledgments This work was supported by Spanish Ministry of Science, Innovation and Universities grants BFU2015-63714-R and PGC2018099341-B-I00 to BI and MINECO/FEDER grants FIS201017440, FIS2015-67765-R, and RTI2018-095802-B-I00 to FJCG. References 1. Neuman KC, Nagy A (2008) Single-molecule force spectroscopy: optical tweezers, magnetic tweezers and atomic force microscopy. Nat Methods 5(6):491–505 2. Bustamante C, Cheng W, Mejia YX (2011) Revisiting the central dogma one molecule at a time. Cell 144(4):480–497 3. Capitanio M, Pavone FS (2013) Interrogating biology with force: single molecule high-

resolution measurements with optical tweezers. Biophys J 105(6):1293–1303 4. Choudhary D, Mossa A, Jadhav M, Cecconi C (2019) Bio-Molecular applications of recent developments in optical tweezers. Biomol Ther 9(1):E23 5. Perkins TT (2014) Ångstro¨m-precision optical traps and applications. Annu Rev Biophys 43:279–302

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6. van Mameren J, Wuite GJ, Heller I (2011) Introduction to optical tweezers: background, system designs, and commercial solutions. Methods Mol Biol 783:1–20 7. Moffitt JR, Chemla YR, Smith SB, Bustamante C (2008) Recent advances in optical tweezers. Annu Rev Biochem 77:205–228 8. Suksombat S, Khafizov R, Kozlov AG, Lohman TM, Chemla YR (2015) Structural dynamics of E. coli single-stranded DNA binding protein reveal DNA wrapping and unwrapping pathways. elife 4:e08193 9. Bell JC, Liu B, Kowalczykowski SC (2015) Imaging and energetics of single SSB-ssDNA molecules reveal intramolecular condensation and insight into RecOR function. elife 4: e08646 10. Hatch K, Danilowicz C, Coljee V, Prentiss M (2007) Direct measurements of the stabilization of single-stranded DNA under tension by single-stranded binding proteins. Phys Rev E 76:021916 11. Pant K, Karpel RL, Rouzina I, Williams MC (2005) Salt dependent binding of T4 gene 32 protein to single and double-stranded DNA: single molecule force spectroscopy measurements. J Mol Biol 349:317–330 12. Shokri L, Marintcheva B, Richardson CC, Rouzina I, Williams MC (2006) Single molecule force spectroscopy of salt-dependent bacteriophage T7 gene 2.5 protein binding to single-stranded DNA. J Biol Chem 281:38689–38696

13. Morin JA, Cerro´n F, Jarillo J, Beltran-HerediaE, Ciesielski GL, Arias-Gonzalez JR, Kaguni LS, Cao FJ, Ibarra B (2017) DNA synthesis determines the binding mode of the human mitochondrial single-stranded DNA-binding protein. Nucleic Acids Res 45(12):7237–7248 14. Cerro´n F, de Lorenzo S, Lemishko KM, Ciesielski GL, Kaguni LS, Cao FJ, Ibarra B (2019) Replicative DNA polymerases promote active displacement of SSB proteins during lagging strand synthesis. Nucleic Acids Res 47:5723–5734 15. Shereda RD, Kozlov AG, Lohman TM, Cox MM, Keck JL (2008) SSB as an organizer/ mobilizer of genome maintenance complexes. Crit Rev Biochem Mol Biol 43:289–318 16. Lohman TM, Ferrari ME (1994) Escherichia coli single-stranded DNA-binding protein: multiple DNA-binding modes and cooperativities. Annu Rev Biochem 63:527–570 17. Smith S, Cui Y, Bustamante C (2003) Opticaltrap force transducer that operates by direct measurement of light momentum. Methods Enzymol 361:134–162 18. Jarillo J, Morı´n JA, Beltra´n-Heredia E, Villaluenga JP, Ibarra B, Cao FJ (2017) Mechanics, thermodynamics, and kinetics of ligand binding to biopolymers. PLoS One 12(4): e0174830 19. Smith S, Cui Y, Bustamante C (1996) Overstretching B-DNA: the elastic response of individual double-stranded and single-stranded DNA molecules. Science 271(5250):795–799

Chapter 18 Measurements of Real-Time Replication Kinetics of DNA Polymerases on ssDNA Templates Coated with Single-Stranded DNA-Binding Proteins Fernando Cerro´n and Borja Ibarra Abstract Optical tweezers can monitor and control the activity of individual DNA polymerase molecules in real time, providing in this way unprecedented insight into the complex dynamics and mechanochemical processes that govern their operation. Here, we describe an optical tweezers-based assay to determine at the singlemolecule level the effect of single-stranded DNA-binding proteins (SSB) on the real-time replication kinetics of the human mitochondrial DNA polymerase during the synthesis of the lagging strand. Key words Optical tweezers, Single-molecule manipulation, DNA replication, Protein-protein interactions, Real-time kinetics

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Introduction In recent years, optical tweezers have been amply used to measure the real-time replication kinetics of various DNA polymerases moving on DNA templates held at constant mechanical tension between two beads [1–8]. Under these conditions, due to the difference in the elastic properties between single- and doublestranded DNA (ssDNA and dsDNA, respectively) [9], as the polymerase coverts the flexible ssDNA template to the more rigid dsDNA helix, the distance between the beads changes. This change in distance is measured with a resolution of 1–10 nm (and 100 Hz), allowing, in this way, to monitor the activity of individual polymerase molecules in real time [10]. Therefore, using DNA extension as a reaction coordinate, it is possible to detect and quantify transient features of the reaction as rare events (i.e., transient inactive states) or heterogeneous behavior. This possibility is being instrumental in unveiling the complex dynamics of DNA polymerases during DNA synthesis. In addition, optical tweezers allows the application of controlled mechanical tensions to the ends of the DNA template.

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_18, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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The effect of increasing tensions on the real-time kinetics of the polymerase provides unprecedented insight into the mechanochemical reactions that control DNA synthesis and proofreading reactions [3, 8]. However, DNA polymerases do not work on naked or free ssDNA; instead, DNA synthesis is carried out on templates heavily coated with ssDNA-binding proteins (SSB). Far from inhibiting DNA synthesis, the tightly bound SSB proteins provide a platform that allows replicative DNA polymerases to process efficiently [11, 12]. To date, the mechanism used by replicative DNA polymerases to dislodge from the template the array of tightly bound SSBs without compromising their replication rates remains unclear. Here, we describe a method aimed at shedding light on how the apparently competing functions of DNA polymerase and SSB are coordinated during the synthesis of the lagging strand [13]. We use optical tweezers to measure the combined effect of mechanical tension and SSB on the instantaneous replication rate of the human mitochondrial DNA polymerase, Pol γ [14], in a single-molecule assay that mimics lagging strand synthesis. As mentioned before, mechanical tension modulates the kinetic rates of DNA polymerases and, when applied to the ends of ssDNA-SSB complexes, regulates the binding modes of SSB proteins [15, 16]. Consequently, mechanical tension is a useful variable that can be readily used to determine the mechanistic processes that govern the replication of SSB-DNA complexes and to quantify the energies involved in the mechanism of SSB release by DNA polymerases. Overall, our results suggest that SSB promotes the maximum replication rate of the DNA polymerase by the elimination of the secondary structure of the template. However, for this to occur, species-specific repulsive interactions (probably electrostatic) between the polymerase-SSB pair are required to overcome the energy of binding of SSB to ssDNA.

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Materials Prepare all stock solutions using ultrapure water and store at room temperature unless otherwise indicated.

2.1 Polystyrene Bead Functionalization

1. Polystyrene beads of 3.18 μm diameter functionalized with protein G (Spherotech, Lake Forest, IL, USA). 2. Polystyrene beads of 2 μm diameter functionalized with streptavidin (Spherotech, Lake Forest, IL, USA). 3. PBS 7.4: 140 μM NaCl, 2.7 mM KCl, 81 mM K2HPO4, 19 mM KH2PO4. Sterilize with autoclave and filter with 0.2 μm polypropylene filter.

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4. Anti-digoxigenin antibody: Dissolve 200 μg of antidigoxigenin antibody in 200 μL PBS 7.4. Store at
20  C. 5. Cross-linking buffer: 100 mM Na2HPO4, 100 mM NaCl. Sterilize with autoclave and filter with 0.2 μm polypropylene filter. 6. Dimethyl pimelimidate (DMP): 20 mg DMP into 400 μL of cross-linking buffer. 7. Eppendorf vortex mixer. 2.2

Flow Chambers

1. Laser engraver. 2. 24  60 mm #2 Coverslips. 3. Parafilm tape. 4. To make micropipettes use glass capillaries of 40 by 80 μm (internal–external diameters, respectively). Use a pipette puller to pull and heat simultaneously individual glass capillaries (http://tweezerslab.unipr.it for more details). The aperture of the micropipette at the tip should be 0.5–1 μm. 5. Bypassing tubes (or dispensers): Glass capillaries of 25–100 μm internal–external diameters. 6. Heating plate.

2.3 Hybrid SingleDouble-Stranded DNA (ssdsDNA) Preparation

1. pBacgus11 double-stranded DNA plasmid. 2. 20 mg/mL Proteinase K stock solution in sterile water. 3. 0.5 M EDTA: Adjust pH to 8.0 with NaOH pellets and sterile filter. 4. Biotin-DNA handle, oligonucleotide sequences: 50 -[Biotin]-GGGTTTGTAAGCCTGAT-30 . 50 -[Phosphate]-AGCTATCAGGCTTACAAAC-30 . 5. Annealing buffer: 10 mM Tris (pH 7.5), 50 mM NaCl, 1 mM EDTA. 6. Thermal cycler. 7. 10 mM (each) dATP, dCTP, dTTP, dGTP, and digoxigeninlabeled dUTP. 8. 100 ng/μL pUC18 plasmid. 9. PCR oligonucleotide primers: Oligo1: 50 -CCTCTGACACATGCAGCTCC-30 . Oligo2: 50 -CGCGGCCTTTTTACGGTTCC-30 . 10. 10 U/μL NBbvCI nicking enzyme. 11. 100 U/μL Exonuclease III. 12. 20 U/μL HindIII-HF restriction enzyme.

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13. 20 U/μL BamHI-HF restriction enzyme. 14. 400 U/μL T4 DNA Ligase. 15. 5 U/μL Taq DNA polymerase. 2.4 Denaturing Alkaline Gel Electrophoresis

1. 10 Denaturing buffer: 300 mM NaOH, 10 mM EDTA. Sterilize by autoclaving.

2.5 Replication Reaction

1. Replication reaction buffer: 50 mM Tris pH 8.5, 30 mM KCl, 10 mM DTT, 4 mM MgCl2, 0.2 mg/mL BSA. Add the four deoxynucleoside triphosphates (dNTPs) to a final concentration of 50 μM when needed. Filter with a 0.22 μm polypropylene filter.

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2. Agarose gel: Dissolve 1 g of agarose in 90 mL of boiling ultrapure water. Let it cool down to 55  C and add 10 mL of 10 denaturing buffer. Mix softly.

Methods

3.1 Preparation of Fluidic Chambers for Optical Tweezers Studies

Counter-propagating optical tweezers employ the light momentum conservation principle to calculate forces accurately. This method guarantees calibration independently of parameters challenging to be estimated precisely (i.e., buffer viscosity, bead radius, and temperature) and of the laser power. Details of construction, calibration, and operation of the counter-propagating optical tweezers setup (Mini-OT) used for the experiments described in this chapter are available at the “Tweezers Lab” website (http:// tweezerslab.unipr.it). A figure describing the fluidic chamber can be found in this volume’s Chapter 17. Follow the next steps for the fabrication of a flow chamber compatible with a Mini-OT device. 1. Drill three 2 mm diameter holes at each side of a clean 24  60 mm coverslip #2 by using the laser engraver. 2. Use the laser engraver to cut three slits (2 mm width) into the parafilm tape. 3. Make a fluidic chamber by sandwiching two coverslips (modified as in step 1) with two layers of the parafilm (modified as in step 2). The sandwich should include a micropipette in the middle and two bypass tubes. 4. Place the chamber on top of a heating plate preheated to 80  C and wait until the parafilm becomes transparent (3–4 min). During this step, the two parafilm layers fuse creating three channels connected by bypass tubes (see Note 1). Then, remove the chamber from the heating plate using tweezers. 5. Mount the microfluidics chamber onto the Mini-OT metal frame by matching fluidic holes.

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6. Fill the top channel with a solution containing DNA-bead preparation (see below), the middle channel with reaction buffer, and the bottom channel with a solution containing 1 μL of streptavidin-covered beads diluted in 300 μL of reaction buffer. 3.2 Preparation of Functionalized Polystyrene Beads

1. Add 750 μL of protein G-coated polystyrene beads to a 1.5 mL microcentrifuge tube and centrifuge for 6 min at 1100  g. Discard supernatant. 2. Dilute the pellet with 750 μL of cross-linking buffer. 3. Add 22.5 μL of DMP (50 μg/μL) and 45 μL of antidigoxigenin antibody (1 μg/μL), mix thoroughly, and incubate for 1 h at room temperature in a microcentrifuge tube shaker set at 123  g. The cross-linking reaction will ensure covalent bonds between the anti-digoxigenin- and the protein G-covered polystyrene beads. 4. Centrifuge for 6 min at 1100  g. Discard supernatant. 5. Add 750 μL of 1 M Tris–HCl pH 7.5 and incubate for 2 h in a microcentrifuge tube shaker set at 123  g to quench the crosslinking reaction. 6. Divide the total volume of the sample into two different 1.5 mL microcentrifuge tubes and add 750 μL of ultrapure water into each tube (see Note 2). 7. Centrifuge for 6 min at 1100  g, discard supernatant, and elute the bead pellet in 750 μL of PBS 7.4. Repeat this step three times (see Note 3). 8. Mix the content of the two tubes in a single tube and centrifuge again for 6 min at 1100  g. Discard supernatant. 9. Elute in 1 mL PBS 7.4 and make 50 mL aliquots. Store at 4  C.

3.3 Preparation of Hybrid ssdsDNA Molecules for Manipulation with Optical Tweezers 3.3.1 Production of ssdsDNA Molecules

Carry out all procedures at room temperature unless otherwise specified. Figure 1 shows a schematic diagram of the preparation protocol.

1. Prepare four test tubes of 1.5 mL. In each tube, add 2 μg of pBacgus11 DNA plasmid, 3 μL of 10 NBbvCI reaction buffer, and 1.8 μL of the 10 U/μL NBbvCI nicking enzyme. Bring the reaction to a final volume of 30 μL with ultrapure water. Mix the solution gently, avoiding the generation of bubbles. 2. Incubate the mixture at 37  C for 1 h. 3. Inactivate NBbvCI by heating the sample for 20 min at 80  C (see Note 4).

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Fig. 1 Schematic representation of the ssdsDNA preparation protocol. Start with a dsDNA vector containing a unique NBbvIC nicking site. Upon nicking reaction, use Exonuclease III (ExoIII) to remove about 1000 nucleotides from the 30 -hydroxyl terminus. Stop reaction and treat sample with proteinase K (Prot K) to digest completely ExoIII. Cut dsDNA with BamHI and HindIII restriction enzymes. Ligate the resulting ends to the complementary ends of dsDNA handles labeled with digoxigenin (red) and biotin (yellow)

4. Add 0.5 μL of 100 U/μL Exonuclease III and incubate for 6 min at room temperature (see Note 5). 5. Add 0.5 M EDTA to a final concentration of 40 mM to stop Exonuclease III reaction. Mix gently without agitation. 6. Add 20 mg/mL proteinase K to a final concentration of 50 μg/ mL, and incubate at 55  C for 1 h and 30 min. Then, allow the sample to cool down to room temperature. 7. Combine the content of the four test tubes. Purify the ssdsDNA using one column of a commercial DNA purification kit. Elute ssdsDNA in a final volume of 30 μL. 8. Digest ssdsDNA with 0.8 μL of 20 U/μL HindIII and 0.8 μL of 20 U/μL BamHI restriction enzymes for 2 h at 37  C in a final volume of 50 μL. 9. Purify ssdsDNA using a commercial DNA purification column.

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10. Store at 4  C for immediate use or keep at
20  C for future use. Estimate the length of the ssDNA gap generated by Exonuclease III by alkaline agarose gel electrophoresis. 3.3.2 Preparation of Digoxigenin-Labeled dsDNA Handles (DIG-DNA Handle)

1. Preparation of dNTP-DIG mix: Mix in a 1.5 mL microcentrifuge tube 6 μL of dATP, 6 μL of dCTP, 6 μL of dGTP, and 3 μL of dTTP (all dNTPs at 10 mM) with 3 μL of digoxigeninlabeled dUTP (10 mM DIG-dUTP). Fill up to 75 μL with ultrapure water. 2. In a 250 μL PCR tube, mix 25 μL of the dNTP-DIG mix with 2 μL each of PCR oligonucleotide primers (1 mM), 10 μL of 10 PCR buffer, 1 μL of 100 ng/μL pUC18 DNA, and 2 μL of 5 U/μL Taq DNA polymerase and fill to 100 μL with ultrapure water. 3. Configure the thermal cycler as follows: step 1: 3 min, 94  C; step 2: 1 min, 94  C; step 3: 1 min, 60  C; step 4: 2 min, 72  C; repeat steps 2–4 29 times; step 5: 10 min at 72  C. 4. Purify DNA using a commercial DNA purification column and elute in a final volume of 30 μL. 5. Add 4 μL of 10 BamHI restriction buffer, 1 μL of 20 U/μL BamHI, and 5 mL of ultrapure water and digest for 2 h at 37  C. 6. Purify the DNA in a final volume of 30 μL using a commercial DNA purification column. 7. Optionally, the resulting DNA products can be checked by agarose gel electrophoresis.

3.3.3 Biotin-Labeled DNA Handle

1. Mix in a 1.5 mL test tube equimolar concentrations of complementary oligo 1 and biotin-labeled oligo 2. Bring the final volume to 100 μL using annealing buffer. 2. Heat to 95  C for 10 min in a thermo-shaker, switch off heating, and wait until the block reaches room temperature (~2 h). 3. Dilute the sample to 1 ng/μL and store at
20  C in 20 μL aliquots.

3.3.4 DNA Ligation

This step is required to label one end of the ssdsDNA with biotin and the other end with digoxigenin for posterior manipulation with the optical tweezers. 1. In a 1.5 mL test tube, mix 50 ng of ssdsDNA, 0.5 μL of 1 ng/μ L biotin-DNA handle, 0.4 μL of 20 ng/μL DIG-DNA handle, 3 μL of 10 T4 reaction buffer, and 1.5 μL of 400 U/μL T4 DNA ligase and fill to 30 μL with ultrapure water. 2. Incubate overnight at 16  C.

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3. Purify the DNA ligation product using a commercial DNA purification column. Elute DNA in 30 μL of water. 4. Store at 4  C for weekly use or store at
20  C for future use. 3.4 Isolation and Manipulation of Single ssdsDNA Molecules in the Optical Tweezers

1. Mix 1–10 ng of the DNA ligation product of ssdsDNA (Subheading 3.3.4), 3 μL of reaction buffer, and 6 μL of antidigoxigenin-covered polystyrene beads in a 1.5 mL test tube and incubate for 10 min at room temperature (see Note 6). 2. Dilute the mixture in 300 μL of the reaction buffer (without dNTPs).

3.4.1 DNA-Bead Incubation 3.4.2 Isolation of Individual ssdsDNA Molecules in the Optical Tweezers

Figure 2 shows a schematic representation of the experimental procedure. Prepare flow chamber and optical tweezers as described in Subheadings 3.1–3.3. 1. Use the optical trap to trap a streptavidin bead and place it on top of the micropipette by air suction. 2. Use the optical trap to trap one bead functionalized with ssdsDNA; place the bead near the streptavidin bead on top of the micropipette. 3. Reset force sensor to 0 piconewtons (pN) (see Note 7). 4. Approach the two beads by moving the optical trap along the yaxis. 5. Attachment of a single ssdsDNA molecule between the two beads will be monitored from the motion of the bead in the optical trap as the two beads are pulled apart (see Note 8). 6. A single ssdsDNA molecule presents unique mechanical properties (a unique force-extension curve) that can be used to identify individual attachments [17, 18].

3.5 Detection of Individual DNA Replication Activities on SSB-Free and SSB-Bound ssDNA 3.5.1 Preparation of a Reaction Solution Containing the DNA Polymerase with/ Without SSB

The following procedures were optimized to study the activity of the human mitochondrial DNA polymerase on ssDNA covered by different types of SSBs. These procedures can be adapted to study other replicative DNA polymerases.

1. Dilute DNA polymerase to 2 nM in 500 μL of reaction buffer (containing 50 μM dNTPs). 2. To measure the activity of DNA polymerase in the presence of SSB, add to the above mixture the desired SSB at a final concentration of 5–50 nM (final concentration may vary depending on the ssDNA-binding properties of the SSB under study).

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Fig. 2 Diagram illustrating the steps to isolate a single ssdsDNA molecule between two beads in the flow chamber of the optical tweezers. (1–3) In the central channel of the flow chamber, use the optical trap (red X) to trap a streptavidin-covered bead (in blue, coming from the bottom channel) and hold it on top of the micropipette by suction. (4–5) Trap a DNA-bead (dark orange) coming from the top channel and place it close to the streptavidin bead. (5–6) Approach the two beads by moving the optical trap along the y-axis. Attachment of a single ssdsDNA molecule between the two beads will be monitored from the motion of the bead in the optical trap as the two beads are pulled apart

3. Filter the solution with a 0.22 μm cellulose acetate syringe filter to eliminate possible protein aggregates and bulky contaminants. 4. Flow the solution into the central channel of the flow chamber through the protein dispenser tube (Fig. 2). 3.5.2 Measurement of Replication Activities on SSB-Free and SSB-Bound ssDNA

Figure 3 shows a schematic representation of the experimental procedure. 1. Upon isolating a single ssdsDNA molecule between the beads, keep a constant force (usually below 20 pN) on the system, and flow the solution containing the polymerase with or without SSB into the central channel of the flow chamber (see Note 9). 2. Generally, SSBs have higher affinities for ssDNA than DNA polymerases, and they will bind to ssDNA first. Binding of SSB to ssDNA will be detected as a change in extension between the beads (usually compaction, see this volume’s Chapter 17).

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Fig. 3 Experimental setup and detection of individual replication activities. (a) Experimental setup to measure individual replication activities in the presence of SSB. Use the optical tweezers to isolate an individual ssdsDNA molecule between two beads: one held in the optical trap (red cone) and the other on top of a micropipette (in light blue). SSB (brown) binds to ssDNA (Δx1) and the DNA polymerase (blue-red) is loaded at the 30 end of the primer template. In the presence of dNTPs, the polymerase starts DNA synthesis releasing SSBs as it advances along the template. At constant force, this activity is measured as a change in the distance between the beads (Δx2). (b) At constant force, the initial force-extension curve of the ssdsDNA molecule (red curve) is gradually converted to dsDNA (blue curve) by the activity of the polymerase (green dots). (c) Representative replication traces of the mitochondrial DNA polymerase on SSB-free ssDNA (blue) and SSB-bound ssDNA (red). Traces were shifted along the time axis for clarity of display

3. The activity of a single-DNA polymerase molecule is followed in real time by measuring the change in extension between beads that occurs as the polymerase converts the flexible SSB-free or SSB-bound ssDNA to the more rigid dsDNA molecule. 4. To convert the change in distance between beads to the number of replicated nucleotides apply the expression

Optical Tweezers to Measure Real-Time DNA Replication Kinetics

Δx Nt ð f Þ ¼

Δx exp ð f Þ , x dsDNA ð f Þ
x ð f Þ

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ð1Þ

where ΔxNt( f ) is the number of nucleotides replicated by the polymerase at a force f, Δxexp( f ) is the experimental change in distance measured between the beads, and xdsDNA( f ) is the average extension of a single dsDNA base pair stretched at a force f [17]. In the absence of SSB, x( f ) corresponds to the average extension of a single ssDNA nucleotide stretched at a force f, whereas in the presence of SSB, x( f ) is the average extension of a single nucleotide wrapped around the SSB at a force f (see Note 10). 5. The average replication rates at each force are determined by a line fit to the traces showing the number of replicated nucleotides versus time (Fig. 3c). 6. The effects of mechanical tension and the presence of SSB bound to the DNA template on the real-time kinetics of a single-DNA polymerase (see Note 11) provide information about the mechanism used by polymerase to release SSB protein during DNA synthesis [13].

4

Notes 1. The application of gentle pressure on the coverslip helps the sealing process. Note that too much pressure will break the coverslips and will favor the progression of the melted parafilm layer too deep into the channel (which, in turn, may block the dispenser tubes and the pipette tip). 2. Use water to dilute the high concentration of Tris–HCl and to facilitate precipitation of the beads onto the bottom of the test tubes during centrifugation. 3. During centrifugation steps, a significant amount of beads will stick to and detach easily from the walls of the tube. Remove the supernatant carefully. 4. After incubation, spin down the water condensed at the lid of the tube. 5. Exonuclease III will digest the 30 end of the nicked DNA plasmid at a rate of ~170 nucleotides/s in the NBbvCI reaction buffer. 6. The concentration of functionalized beads can vary from one preparation to the other. Therefore, the amount of DNA can be varied from 1 to 10 ng per reaction in order to obtain a proper DNA:bead ratio for manipulation of single molecules in the optical tweezers. 7. Avoid physical contact between the beads in this step.

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8. The sensitive detection of the bead position in the trap and, by extension, the calibrated measurement of displacement and force are determined by back-focal-plane interferometry. 9. In the absence of SSB, ss and dsDNA present similar extensions at tensions of ~5–7 pN. This similarity will prevent detection of activities within this fore range. To measure DNA replication on SSB-DNA, hold the protein-DNA complex at tensions below ~8 pN, where SSB remains stably bound to the ssDNA [15, 16]. 10. Calculate xssDNA( f ) and x( f ) from their corresponding experimental force-extension curves. xdsDNA( f ) is obtained from the wormlike chain model for semiflexible polymers with a persistence length of 53 nm and a stretch modulus of 1200 pN/ nm [19]. 11. Typical values that can be measured are average velocity, velocity without pauses, and pause occupancy. To compute the velocity without pauses, use the method described in [13]. In this method, a variable time window is used to calculate the instantaneous replication rate of the polymerase, thus enabling the identification of the optimal time window range to calculate the maximum replication rate of the enzyme.

Acknowledgments This work was supported by Spanish Ministry of Science, Innovation and Universities grants BFU2015-63714-R and PGC2018099341-B-I00 to BI. References 1. Wuite GJL, Smith SB, Young M, Keller D, Bustamante C (2000) Single-molecule studies of the effect of template tension on T7 DNA polymerase activity. Nature 404:103–106 2. Maier B, Bensimon D, Croquette V (2000) Replication by a single DNA polymerase of a stretched single-stranded DNA. Proc Natl Acad Sci U S A 97:12002–12007 3. Ibarra B, Chemla YR, Plyasunov S, Smith SB, La´zaro JM, Salas BC (2009) Proofreading dynamics of a processive DNA polymerase. EMBO J 28:2794–2802 4. Kim S, Schroeder CM, Xie XS (2010) Singlemolecule study of DNA polymerization activity of HIV-1 reverse transcriptase on DNA templates. J Mol Biol 395:995–1006 5. Morin JA, Cao FJ, La´zaro JM, Arias-Gonzalez JR, Valpuesta JM, Carrascosa JL, Salas M, Ibarra B (2012) Active DNA unwinding

dynamics during processive DNA replication. Proc Natl Acad Sci U S A 109:8115–8120 6. Manosas M, Spiering MM, Ding F, Bensimon D, Allemand JF, Benkovic SJ, Croquette V (2012) Mechanism of strand displacement synthesis by DNA replicative polymerases. Nucleic Acids Res 40:6174–6186 7. Naufer MN, Murison DA, Rouzina I, Beuning PJ, Williams MC (2017) Single-molecule mechanochemical characterization of E. coli pol III core catalytic activity. Protein Sci 26:1413–1426 8. Hoekstra TP, Depken M, Lin SN, CabanasDane´s J, Gross P, Dame RT, Peterman EJG, Wuite GJL (2017) Switching between exonucleolysis and replication by T7 DNA polymerase ensures high fidelity. Biophys J 112:575–583

Optical Tweezers to Measure Real-Time DNA Replication Kinetics 9. Bustamante C, Smith SB, Liphardt J, Smith D (2000) Single-molecule studies of DNA mechanics. Curr Opin Struct Biol 10:279–285 10. Moffitt JR, Chemla YR, Smith SB, Bustamante C (2008) Recent advances in optical tweezers. Annu Rev Biochem 77:205–228 11. Shereda RD, Kozlov AG, Lohman TM, Cox MM, Keck JL (2008) SSB as an organizer/ mobilizer of genome maintenance complexes. Crit Rev Biochem Mol Biol 43:289–318 12. Oliveira MT, Kaguni LS (2011) Reduced stimulation of recombinant DNA polymerase γ and mitochondrial DNA (mtDNA) helicase by variants of mitochondrial single-stranded DNA-binding protein (mtSSB) correlates with defects in mtDNA replication in animal cells. J Biol Chem 286(47):40649–40658 13. Cerro´n F, de Lorenzo S, Lemishko KM, Ciesielski GL, Kaguni LS, Cao FJ, Ibarra B (2019) Replicative DNA polymerases promote active displacement of SSB proteins during lagging strand synthesis. Nucleic Acids Res 47 (11):5723–5734 14. Kaguni LS (2004) DNA polymerase gamma, the mitochondrial replicase. Annu Rev Biochem 73:293–320

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15. Morin JA, Cerro´n F, Jarillo J, Beltran-HerediaE, Ciesielski GL, Arias-Gonzalez JR, Kaguni LS, Cao FJ, Ibarra B (2017) DNA synthesis determines the binding mode of the human mitochondrial single-stranded DNA-binding protein. Nucleic Acids Res 45(12):7237–7248 16. Suksombat S, Khafizov R, Kozlov AG, Lohman TM, Chemla YR (2015) Structural dynamics of E. coli single-stranded DNA binding protein reveal DNA wrapping and unwrapping pathways. elife 4:e08193 17. Smith SB, Finzi L, Bustamante C (1992) Direct mechanical measurements of the elasticity of single DNA molecules using magnetic beads. Science 258(5085):1122–1126 18. Ibarra B, Chemla YR, Plyasunov S, Smith SB, La´zaro JM, Salas M, Bustamante C (2009) Proofreading dynamics of a processive DNA polymerase. EMBO J 28(18):2794–2802 19. Bouchiat C, Wang MD, Allemand JF, Strick T, Block SM, Croquette V (1999) Estimating the persistence length of a worm-like chain molecule from force-extension measurements. Biophys J 76(1):409–413

Chapter 19 Selective Suppression of Endogenous Gene Expression Using RNAi in Drosophila Schneider S2 Cells Yuichi Matsushima Abstract RNA interference (RNAi) is a posttranscriptional gene silencing method that is triggered by doublestranded RNA (dsRNA). RNAi is used to inactivate genes of interest and provides a genetic tool for lossof-function studies in a variety of organisms. I have used this method to reveal the physiological roles of a number of endogenous proteins involved in mitochondrial DNA metabolism in Schneider cells, including the mitochondrial single-stranded DNA-binding protein. Here, I present experimental schemes of selective suppression of endogenous gene expression using RNAi in Drosophila Schneider S2 cells. With this method, the function of exogenous wild-type or mutant genes can be evaluated. Key words RNAi, RNA interference, Drosophila, Schneider cells, mtSSB

1

Introduction RNA interference (RNAi) is a powerful tool that allows researchers to investigate gene function by “knocking down” or reducing the levels of a particular protein [1]. In RNAi, the introduction of target-specific double-stranded RNA (dsRNA) results in the silencing of target genes. This phenomenon was first observed in C. elegans [2]. RNAi technology is used in various species, from protozoa to animals [1]. In Drosophila, RNAi has been applied to both cell culture and animals. Transfection of dsRNA into Drosophila Schneider cells results in effective gene-specific knockdown [3], but the effect of transfected dsRNA is transient. However, if the dsRNA is produced from a genome-integrated expression vector (i.e., in an established RNAi cell line), the gene silencing effect can be monitored continuously. Additionally, exon-specific RNAi has been developed in Drosophila cells for the selective suppression of each alternative splicing variant [4, 5]. Interestingly, exon-specific RNAi does not work in worms and plants [6]. Worms and plants contain RNA-dependent RNA polymerase which synthesizes

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_19, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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secondary siRNAs associated with exon-specific siRNA-mediated target mRNA degradation [7]. These secondary siRNAs work against all mRNA-containing regions derived from common exons of the target mRNA. However, exon-specific RNAi can be used in higher animals which lack RNA-dependent RNA polymerase, including fly, mouse, and human. In this chapter, I describe the use of untranslated region (UTR)-specific dsRNA instead of exonspecific dsRNA. The use of UTR-specific dsRNA allows for the degradation of endogenous mRNA, but not of exogenous mRNA lacking the UTR region (Fig. 1). I use RNAi to study the function of several essential constituents of the mitochondrial DNA replication and transcription machinery in Drosophila cells [8, 9]. To evaluate the function of exogenous wild-type or mutant genes, I selectively suppress endogenous gene expression. In this chapter, I describe how to establish cell lines for the selective suppression of endogenous gene expression using RNAi in Drosophila Schneider S2 cells and the gene for the mitochondrial single-stranded DNA-binding protein (mtSSB) as an example. To this end, I develop doubly transfected cell lines with both RNAi and exogenous gene expression plasmids. To inhibit the expression of endogenous genes, I developed a strategy to produce double-stranded interfering RNAs targeted to the UTR regions of the target genes (Fig. 1). These dsRNAs do not degrade mRNA lacking the UTR region.

mtSSB expression vector

mtSSB gene endogenous promoter

mRNA

inducible promoter

endogenous mtSSB

endogenous mtSSB

mRNA degradation

AAAAAAAAAAAA

mRNA

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expression RNA inducible promoter

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Fig. 1 Schematic diagram depicting selective suppression of endogenous Drosophila mtSSB expression by RNAi in Schneider S2 cells. A 30 -UTR mtSSB fragment was cloned as an inverted repeat into the pMt/Hy inducible vector. In induced conditions, exogenous mtSSB mRNA and inverted repeat RNA are expressed. The induced inverted repeat RNA forms dsRNA which triggers the specific degradation of endogenous mtSSB mRNA. However, the dsRNA does not affect the induced exogenous mtSSB mRNA, which lacks its original 30 -UTR

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Materials

2.1 RNAi in Schneider Cells

1. pMt/Hy expression vector [10].

2.1.1 Construction of Exogenous mtSSB Expression Vectors and the RNAi Vector

3. Gel extraction kit.

2. Restriction enzymes XhoI, SpeI, and EcoRI. 4. PrimeSTAR DNA polymerase. 5. PCR purification kit. 6. Quick Ligation Kit. 7. SURE 2 Supercompetent Cells. 8. LB plate containing 100 μg/mL ampicillin. 9. Plasmid midi kit. 10. 3 M Sodium acetate (pH 5.2). 11. 100% Ethanol. 12. 70% Ethanol. 13. Phenol/chloroform/isoamyl alcohol (25:24:1, v/v). 14. TE buffer: 10 mM Tris–HCl pH 8.0, 1 mM EDTA. 15. Primers for the construction of the mtSSB expression vector: Forward (50 –30 ) GCGCCTCGAGTGCAACCACAAGGCG CATGCTGAATCCTCTGTTGACC and reverse (50 –30 ) GC GCACTAGTTTAGTTGTTGGCATCACGG. 16. Primers for the sense fragment of RNAi vector: Forward (50 –30 ) GCGCCTCGAGACTAGTAATTTAAGCCCAGAT CAC and reverse (50 –30 ) GCGCGAATTCGGGATCGATG GAGTACGACTACGCATG. 17. Primers for the antisense fragment of RNAi vector: Forward (50 –30 ) GCGCCTCGAGACTAGTAATTTAAGCCCAGAT CAC and reverse (50 –30 ) GCGCGAATTCAAAAAGCTTG GAGTACGACTACGCATG.

2.1.2 Establishment of the Cell Lines

1. Schneider S2 cells. 2. Schneider’s Drosophila medium supplemented with 10% fetal bovine serum. 3. 60 mm Dishes. 4. Effectene Transfection Reagent. 5. 96-Well culture plates. 6. Hygromycin. 7. 75 cm2 Culture flasks. 8. 125 cm2 Culture flasks. 9. 100 mM CuSO4

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10. Phosphate-buffered saline (PBS): 135 mM NaCl, 10 mM Na2HPO4, 2 mM KCl, 2 mM KH2PO4. 11. Lysis buffer: 10 mM Tris–HCl pH 8.0, 5 mM EDTA, and 1% SDS. 12. BCA Protein Assay Kit.

3

Methods

3.1 Selective Suppression of Endogenous Drosophila mtSSB in Schneider Cells 3.1.1 Construction of the mtSSB Expression Vector

A schematic diagram depicting the construction map of exogenous Drosophila mtSSB (wild-type or mutant) gene expression plasmids is shown in Fig. 2a. 1. Digest 1 μg of pMt/Hy expression vector with 10 units each of XhoI and SpeI in a reaction volume of 20 μL. 2. Separate the digested DNA fragments by electrophoresis using a 0.7% agarose gel. 3. Purify the digested vector from the agarose gel using a gel extraction kit. 4. Amplify the sense fragment with PrimeSTAR DNA polymerase and the following primer pair: 50 -GCGCctcgag TGCAACCA CAAGGCGCATGCTGAATCCTCTGTTGACC -30 and 50 -GCGCactagtTTAGTTGTTGGCATCACGG-30 . 5. Purify the sense PCR products using a PCR purification kit. 6. Digest the sense PCR fragment with 10 units each of XhoI and SpeI in a reaction volume of 20 μL. 7. Electrophorese the reaction mixture using a 1% agarose gel. 8. Purify the sense fragment from the agarose gel using a gel extraction kit. 9. Ligate the vector DNA with the sense fragment using the Quick Ligation Kit. Use a molecular ratio of vector:sense fragment of 1:3. 10. Transform E. coli SURE 2 cells with 1 μL of ligation mixture using the E. coli pulser, and plate on an LB plate containing 100 μg/mL ampicillin. 11. Recover plasmids from the colonies and check their identity and integrity by restriction endonuclease digestion of a small aliquot, following steps 1 and 2. 12. Select positive colonies and purify the plasmid DNA using a plasmid midi kit. 13. Dissolve DNA in 400 μL of TE buffer. 14. Add 400 μL of phenol/chloroform/isoamyl alcohol (25:24:1, v/v), mix well, and then separate the phases via centrifugation at 20,000  g for 5 min.

Selective RNAi in Drosophila Cells

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Fig. 2 (a) Schematic representation of an exogenous Drosophila mtSSB gene expression plasmid without the 30 -UTR region. Wild-type or mutant mtSSB was cloned into the pMt/Hy vector. (b) Schematic representation of a dsRNA expression plasmid targeted to the 30 -UTR region of the Drosophila mtSSB gene. Gray arrows show sense and antisense sequences. The open box indicates the loop region. The loop region is 24 nucleotides in length and contains three restriction enzyme sites, ClaI—CCC—EcoRI—AAA—HindIII

15. Transfer the aqueous phase to a new tube, add 40 μL of 3 M sodium acetate (pH 5.2) and 800 μL of 100% ethanol, mix, and incubate at room temperature for 10 min. 16. Centrifuge at 20,000  g for 10 min and discard the supernatant. Rinse the pellet with 600 μL of 70% ethanol. Centrifuge at 20,000  g for 5 min and discard the supernatant. 17. Dry the pellet at room temperature for 10 min and dissolve the pellet in 50 μL of TE buffer. 3.1.2 Construction of the RNAi Vector

A schematic depicting the structure of a dsRNA expression plasmid targeting the 30 UTR region of Drosophila mtSSB gene in Schneider cells is shown in Fig. 2b (see Note 1). 1. Digest 1 μg of pMt/Hy plasmid with 10 units each of XhoI and SpeI in a 20 μL reaction volume. 2. Load the reaction mixture onto a 0.7% agarose gel and electrophorese. 3. Purify the digested vector from the agarose gel using a gel extraction kit. 4. Amplify the sense fragment with PrimeSTAR DNA polymerase and the primer pair 50 -GCGCctcgagACTAGTAATTTAAGCC CAGATCAC -30 and 50 -GCGCgaattc GGGATCGATGGAG TACGACTACGCATG-30 .

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5. Amplify the antisense fragments with PrimeSTAR DNA polymerase and the primer pair 50 - GCGCCTCGAG actagt AATT TAAGCCCAGATCAC -30 and 50 -GCGCgaattc AAAAAGCTTGGAGTACGACTACGCATG-30 . 6. Purify the sense and antisense PCR products using a PCR purification kit. 7. Digest sense PCR fragments with 10 units each of XhoI and EcoRI, and digest antisense PCR fragments with 10 units each of SpeI and EcoRI. 8. Load the reaction mixture onto a 2.0% agarose gel and electrophorese. 9. Purify the sense and antisense fragments from the agarose gel using a gel extraction kit. 10. Ligate the vector DNA with sense and antisense fragments using the Quick Ligation Kit. Use molecular ratio of vector: sense fragment:antisense fragment of 1:3:3. 11. Transform E. coli SURE 2 cells with 1 μL of ligation mixture using the E. coli pulser, and plate on an LB plate containing 100 μg/mL ampicillin. 12. Recover plasmids from the colonies and check their identity and integrity by restriction endonuclease digestion of a small aliquot, following steps 1 and 2. 13. Select positive colonies and purify the plasmid using a plasmid midi kit. 14. Dissolve DNA in 400 μL of TE buffer. 15. Add 400 μL of phenol/chloroform/isoamyl alcohol (25:24:1, v/v), mix well, and separate the phases via centrifugation at 20,000  g for 5 min. 16. Transfer the aqueous phase to a new tube, add 40 μL of 3 M sodium acetate (pH 5.2) and 800 μL of ethanol, mix, and incubate at room temperature for 10 min. 17. Centrifuge at 20,000  g for 10 min and discard the supernatant. Rinse the pellet with 600 μL of 70% ethanol. Centrifuge at 20,000  g for 5 min and discard the supernatant. 18. Dry the pellet at room temperature for 10 min and dissolve the pellet in 50 μL of TE buffer. 3.1.3 Establishment of Cell Lines

1. Culture Drosophila Schneider S2 cells at 25  C in Schneider’s Drosophila medium supplemented with 10% fetal bovine serum. Subculture cells to 3–5  106 cells/mL every 3rd to 5th day. 2. Place 4 mL of cells at a density of 3–5  106 cells/mL into a 60 mm dish 24 h prior to transfection.9

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3. Mix in a tube the expression vector and the RNAi vector in a 1:3 ratio for use in transfection. For the establishment of RNAi cell lines, use the RNAi vector alone for transfection. 4. Transfect the cells using Effectene Transfection Reagent, following the manufacturer’s instructions. 5. Incubate for 24–48 h at 25  C. 6. Transfer cells from the 60 mm dish into a 96-well culture plate containing 20 mL of fresh Schneider’s Drosophila medium supplemented with 10% fetal bovine serum and 200 μg/mL hygromycin. 7. Incubate at 25  C for 8–15 days until hygromycin-resistant colonies grow. 8. Transfer single colonies each into a 75 cm2 culture flask containing 10 mL of fresh Schneider’s Drosophila medium supplemented with 10% fetal bovine serum and 100–200 μg/mL hygromycin. 9. Incubate at 25  C until cells reach a density of 15–20  106 cells/mL or for 5–7 days. 10. Transfer the cells into a 125 cm2 culture flask containing 20 mL of fresh Schneider’s Drosophila medium supplemented with 10% fetal bovine serum and 100–200 μg/mL hygromycin. 11. Incubate at 25  C until cells reach a density of 15–20  106 cells/mL or for 5–7 days. 12. Transfer 7 mL of the culture into a 125 cm2 flask containing 20 mL of fresh Schneider’s Drosophila medium supplemented with 10% fetal bovine serum and 100–200 μg/mL hygromycin. 13. Incubate at 25  C until cells reach a density of 15–20  106 cells/mL or for 5–7 days. 14. Culture the selected Schneider S2 cells at 25  C in Schneider’s Drosophila medium supplemented with 10% fetal bovine serum, subculturing 3–5  106 cells/mL every 3rd to 5th day. 3.2 Induction of dsRNA Expression with or Without Exogenous mtSSB

1. Dilute cells to 3–5  106 cells/mL and add CuSO4 to a final concentration of 0.4 mM. 2. Incubate at 25  C and subculture to 3–5  106 cells/mL every 3rd day. 3. After 10 days of culture, harvest cells via centrifugation at 2000  g for 5 min. 4. Wash with PBS and centrifuge at 2000  g for 5 min. 5. Add lysis buffer, sonicate the lysate, heat at 100  C for 5 min, and centrifuge at 20,000  g for 10 min.

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Fig. 3 Expression of Drosophila mtSSB in Schneider cells. (a) Immunoblot analysis of cell extracts probed with rabbit antiserum against the Drosophila mtSSB [8]. Schneider cells with no plasmid (control) or carrying an empty pMt/Hy (vector), RNAi vector (RNAi), RNAi vector + wild-type mtSSB (RNAi + WT), or RNAi vector + mutant mtSSB (W79T/F85A) (RNAi + Mut) were cultured for 10 days in the presence or absence of inducer (0.4 mM CuSO4). Protein extracts (10 μg) were fractionated by 17% SDS-PAGE, transferred to nitrocellulose filters, and probed with rabbit antiserum against d-mtSSB [8] as indicated. After induction, the endogenous mtSSB levels were depleted in the cells expressing RNAi. Induction of exogenous mtSSB was confirmed in cells with both an RNAi vector and exogenous expression vector (WT or Mut) (see Notes 3 and 4). (b) The mitochondrial DNA:nuclear DNA (mtDNA:nDNA) ratio in Schneider S2. The mtDNA:nDNA ratio in Schneider S2 was determined by Southern blot analysis. DNA was isolated after growth for 10 days in the presence or absence of 0.4 mM CuSO4. A fragment of the ATPase6 gene was used as the probe for determination of mtDNA content, and a DNA fragment from the nuclear histone gene cluster was used for normalization. The value of the S2 cell line grown in the absence of 0.4 mM CuSO4 was taken as 1. The data was analyzed using the Student’s two-tailed t-test and is presented as means  SD (*P < 0.05; ***P < 0.001). The depletion of endogenous mtSSB reduced the relative mtDNA/nDNA ratio. Expression of exogenous wild-type mtSSB rescues the mtDNA depletion phenotype. The expression of exogenous mutant mtSSB does not rescue the mtDNA depletion phenotype

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6. Transfer the lysate to a fresh tube and assay the protein concentration, using the BCA Protein Assay Kit. 7. Check the level of the target protein via immunoblot analysis (Fig. 3a) (see Notes 3 and 4). 3.3 Mitochondrial DNA Analysis

1. Dilute cells to 3–5  106 cells/mL and add CuSO4 to a final concentration of 0.4 mM. 2. Incubate at 25  C and subculture to 3–5  106 cells/mL every 3rd day. 3. After 10 days of culture, harvest cells via centrifugation at 2000  g for 5 min. 4. Wash with PBS and centrifuge at 2000  g for 5 min. 5. Purify genomic DNA from Drosophila Schneider S2 cells via standard methods. 6. Determine the relative mtDNA copy number from the cells using qPCR or Southern blot (Fig. 3b).

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Notes 1. Due to restrictions in the selectable region, a short dsRNA expression vector with a stem region less than 100 bp is used. If there are fewer restrictions on the selectable area it is often better to use a longer stem region (>300 bp). 2. Continuous transfection of in vitro-synthesized dsRNA may be used instead of an RNAi vector. 3. The RNAi construct may repress the target transcript under uninduced conditions because of leaky expression from the metallothionein promoter (see Fig. 3a). 4. The cell lines are stable for at least 3 months. The target protein level should be regularly checked by immunoblot. If sufficient suppression/induction is not apparent, it is best practice to establish a new cell line.

Acknowledgments I would like to thank Dr. Rebecca Porter from Edanz Group (www. edanzediting.com/ac) for editing a draft of this manuscript. This work was supported by the Japan Society for the Promotion of Science (JSPS) KAKENHI Grant Numbers 15H03096, 18H03180, and 18K19303.

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References 1. Wilson RC, Doudna JA (2013) Molecular mechanisms of RNA interference. Annu Rev Biophys 42:217–239. https://doi.org/10. 1146/annurev-biophys-083012-130404 2. Fire A, Xu S, Montgomery MK, Kostas SA, Driver SE, Mello CC (1998) Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature 391 (6669):806–811 3. Hammond SM, Bernstein E, Beach D, Hannon GJ (2000) An RNA-directed nuclease mediates post-transcriptional gene silencing in Drosophila cells. Nature 404(6775):293–296 4. Celotto AM, Lee JW, Graveley BR (2005) Exon-specific RNA interference: a tool to determine the functional relevance of proteins encoded by alternatively spliced mRNAs. Methods Mol Biol 309:273–282. https://doi. org/10.1385/1-59259-935-4:273 5. Celotto AM, Graveley BR (2002) Exonspecific RNAi: a tool for dissecting the functional relevance of alternative splicing. RNA 8 (6):718–724. https://doi.org/10.1017/ s1355838202021064 6. Hutvagner G, Zamore PD (2002) RNAi: nature abhors a double-strand. Curr Opin

Genet Dev 12(2):225–232. https://doi.org/ 10.1016/s0959-437x(02)00290-3 7. Sijen T, Fleenor J, Simmer F, Thijssen KL, Parrish S, Timmons L, Plasterk RH, Fire A (2001) On the role of RNA amplification in dsRNA-triggered gene silencing. Cell 107 (4):465–476. https://doi.org/10.1016/ s0092-8674(01)00576-1 8. Farr CL, Matsushima Y, Lagina AT 3rd, Luo N, Kaguni LS (2004) Physiological and biochemical defects in functional interactions of mitochondrial DNA polymerase and DNA-binding mutants of single-stranded DNA-binding protein. J Biol Chem 279 (17):17047–17053 9. Matsushima Y, Goto Y, Kaguni LS (2010) Mitochondrial Lon protease regulates mitochondrial DNA copy number and transcription by selective degradation of mitochondrial transcription factor A (TFAM). Proc Natl Acad Sci U S A 107(43):18410–18415. https://doi. org/10.1073/pnas.1008924107 10. Koelle MR, Talbot WS, Segraves WA, Bender MT, Cherbas P, Hogness DS (1991) The Drosophila EcR gene encodes an ecdysone receptor, a new member of the steroid receptor superfamily. Cell 67(1):59–77

Chapter 20 Stimulation of Variant Forms of the Mitochondrial DNA Helicase Twinkle by the Mitochondrial Single-Stranded DNA-Binding Protein Ana P. C. Rodrigues and Marcos T. Oliveira Abstract Defects in mitochondrial DNA (mtDNA) maintenance may lead to disturbances in mitochondrial homeostasis and energy production in eukaryotic cells, causing diseases. During mtDNA replication, the mitochondrial single-stranded DNA-binding protein (mtSSB) stabilizes and protects the exposed singlestranded mtDNA from nucleolysis; perhaps more importantly, it appears to coordinate the actions of both the replicative mtDNA helicase Twinkle and DNA polymerase gamma at the replication fork. Here, we describe a helicase stimulation protocol to test in vitro the functional interaction between mtSSB and variant forms of Twinkle. We show for the first time that the C-terminal tail of Twinkle is important for such an interaction, and that it negatively regulates helicase unwinding activity in a salt-dependent manner. Key words Twinkle, P66, mtSSB, Mitochondrial DNA replication, dsDNA unwinding assay

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Introduction Mitochondria are unique organelles because they have their own genome, which has its own inheritance mode, and is replicated and maintained independently of the cell’s nuclear genome [1]. The mitochondrial DNA (mtDNA) in humans and most animals is a circular double-stranded molecule of approximately 16 kb, composed of 37 genes, 13 of which encode central subunits of four multienzymatic complexes involved in mitochondrial oxidative phosphorylation (OXPHOS) [2, 3]. This process is known to produce about 90% of the ATP consumed by cells in diverse, vital enzymatic reactions [4]. In addition, mitochondria are also involved in apoptosis, calcium signaling, autophagy, and production of a variety of intermediates for a plethora of cellular metabolic pathways [5].

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_20, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Replication of the mitochondrial genome is accomplished by a machinery containing at least four nuclear-encoded proteins, assembled at the mtDNA replication fork. The replicative mtDNA helicase Twinkle, DNA polymerase gamma (Pol γ, subunits α and β), and mitochondrial single-stranded DNA-binding protein (mtSSB) are sufficient for in vitro synthesis of new DNA molecules that can be as long as the entire mitochondrial genome. For this reason, together they are called the minimum mitochondrial replisome [1, 3, 6]. Each of these replisome proteins appears to be equally necessary, because their deletion or depletion, or mutations affecting their function, can lead to aging, and clinical or deleterious phenotypes both in humans and in animal models [3]. Twinkle is a homohexameric/heptameric ring-shaped enzyme, which is a homolog of the bacteriophage T7 primase-helicase gp4. It is responsible for unwinding the double-stranded (ds) mtDNA most likely using a steric exclusion mechanism. In this model, Twinkle binds one parental single-stranded (ss) DNA, forcing it through its central pore as the enzyme translocates in the 50 –30 direction, at the same time that the complementary ssDNA is forced out [1, 3, 7]. ssDNA translocation, which requires the energy of nucleotide triphosphate hydrolysis, leads to the release of the ssDNA that will serve as template for a putative mitochondrial primase and Pol γ to synthesize new mtDNA strands [1, 3]. Human Pol γ has a heterotrimeric configuration, with one catalytic subunit (Pol γ-α) comprising the 50 –30 DNA polymerase, 30 –50 exonuclease, and 50 -dRP lyase activities of the enzyme, and a dimeric accessory subunit (Pol γ-β) that enhances the activities of Pol γ-α [1, 3]. During replication, the homotetrameric mtSSB binds the exposed parental ssDNA to stabilize and protect it from nucleolysis, but also to coordinate both Twinkle and Pol γ activities [1, 8–11]. mtSSB stimulates the rate of dsDNA unwinding by Twinkle in vitro [8–11], and the concerted action of Pol γ and Twinkle in strand-displacement DNA synthesis assays [6]. Although the mechanisms that regulate these events are still unclear, it is possible that mtSSB physically interacts with Twinkle, recruiting it to DNA, and/or enhancing its translocation properties. An alternative explanation is that mtSSB simply suppresses the reannealing of unwound ssDNA by binding to it, thus representing a passive mechanism for Twinkle stimulation [11, 12]. Here, we describe a protocol of helicase stimulation assay to quantify the functional interaction between Twinkle and mtSSB (which could actually be applied to any helicase-SSB system), and provide evidence that both physical interaction and passive stimulation take place at the mtDNA replication fork. We use a C-terminal truncation variant of Twinkle, called P66 [13], that has mtSSB-independent, intrinsically higher ATPase and dsDNA unwinding activities under low-salt conditions, but that is almost completely inactive at a salt concentration closer

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to the physiological conditions. Under the latter conditions, the fold stimulation of P66 by mtSSB is comparable to that of the fulllength, wild-type Twinkle (Twinkle WT), although the overall unwinding by P66-mtSSB is still low. P66 was originally designed to be the recombinant version of the first product generated upon limited trypsin proteolysis of Twinkle WT. It retains the ability to form homo-oligomers, and each subunit has a predicted size of 66 kDa [13]. The C-terminal 44 amino acids that are absent in P66 represent an unstructured tail in our homology model of Twinkle (Fig. 1), with no sequence similarity with the C-terminal tail of T7 gp4, but well conserved among most mammals [14]. Further investigation of the properties of this Twinkle variant will certainly provide valuable information on how the enzyme functions in concert with the other mitochondrial replisome components, to help maintain mitochondrial and cellular homeostasis.

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Materials 1. 1.5 mL microcentrifuge tubes. 2. Water bath or heat block (37–95  C). 3. Purified recombinant human mtSSB (see Note 1). 4. Purified recombinant human Twinkle WT (see Note 1). 5. Purified recombinant human Twinkle P66 (see Note 1). 6. Radiolabeled DNA unwinding substrate: circular ss pBluescript KS(+) DNA (2958 nt) hybridized to 20 nucleotides of a 50 -radiolabeled 60-mer oligodeoxyribonucleotide (50 T(40) AGG TCGTTCGCTCCAAGCT30 ) (see Note 2). 7. Reaction mixture 20: 20 mM Tris–HCl pH 7.5, 10% glycerol, 500μg/mL bovine serum albumin, 10 mM dithiothreitol, 4 mM MgCl2, 3 mM ATP, 0.4 nM of DNA unwinding substrate, and 20 mM KCl. 8. Reaction mixture 100: 20 mM Tris–HCl pH 7.5, 10% glycerol, 500μg/mL bovine serum albumin, 10 mM dithiothreitol, 4 mM MgCl2, 3 mM ATP, 0.4 nM of DNA unwinding substrate, and 100 mM KCl. 9. 10 Stop solution: 6% SDS, 100 mM EDTA, pH 8.0. 10. 10 Loading buffer: 50% Glycerol, 0.25% bromophenol blue. 11. 1 TBE: 90 mM Tris–HCl–borate, 2 mM EDTA. 12. 22% Native polyacrylamide gel: 59:1 acrylamide/bisacrylamide in 1 TBE. 13. Vacuum gel drier with heat. 14. Phosphor Screen and Storm 820 Scanner.

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Fig. 1 Structural model of Twinkle and its possible interactions with DNA and mtSSB. (a) Homology model of a single Twinkle subunit [14], showing the zinc-binding-like (ZBD), the RNA polymerase-like (RPD), the linker, and the helicase domains [14]. The unstructured 44-amino acid-long C-terminal tail is shown in red. (b) In the model of a homoheptameric Twinkle (cyan) bound to a forklike DNA (gray), the C-terminal tails would be positioned ahead of the enzyme, toward dsDNA. There are no laboratory or in silico data to support the configuration of the C-terminal tails presented; in fact, we speculate that they are flexible and dynamic structures that could interact with dsDNA, mtSSB (purple), the helicase domain of adjacent subunits, and possibly other mitochondrial factors. Although mtSSB could stimulate Twinkle activity by binding both ssDNA strands released upon unwinding, the putative physical interaction mechanism, suggested by our results (see

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Methods

3.1 dsDNA Unwinding Assay

1. Prepare reaction mixtures 20 and 100 on ice, and dispense 50μL into 1.5 mL microcentrifuge tubes. Incubate tubes for 5 min in a water bath or heat block at 37  C (see Note 3). 2. Set two tubes aside which will be referred to as “boil –” and “boil +” controls. Do not add any helicase in these tubes. The “boil –” control represents the unreacted, intact substrate, and the “boil +” control represents completely melted substrates (see Fig. 2a) after boiling (see step 5 below). 3. Add 1.8, 3.5, 7.0, and 14.0 nM Twinkle WT or P66 (both calculated as hexamers) into the remaining tubes containing the reaction mixtures, and incubate all tubes for 30 min at 37  C. 4. Stop the reaction by addition of 5μL of 10 stop solution, followed by 5μL of 10 loading buffer. 5. Incubate the “boil +” control in a water bath or heat block for 5 min at 95  C. 6. Subject 20–25μL of each reaction sample to electrophoresis at room temperature in a 22% native polyacrylamide gel at 600 V until the bromophenol blue dye travels approximately 3 cm, using 1 TBE as running buffer (see Note 4). 7. Dry the gel under vacuum with heat, and expose it to a phosphor screen for at least 1 h (see Note 5). 8. Scan the phosphor screen using a Storm 820 Scanner (see Note 6). A representative outcome of this analysis is shown in Fig. 2a, in which the upper bands in the gels represent the unreacted, intact (non-unwound) substrate molecules, and the lower bands represent the 60-mer oligonucleotide product of the unwinding reaction. 9. Determine the volume of each band by applying computer integration analysis of the radioactive signals using the ImageQuant software (see Note 6) and subtracting the background. The background is the apparent signal of the unreacted, intact substrate in the “boil +” sample, and of the 60-mer oligonucleotide product in the “boil –” sample (see Fig. 2a).

ä Fig. 1 (continued) Fig. 3), could only take place if the Twinkle C-terminal tails interact with mtSSB molecules bound to the opposite ssDNA strand. Such interactions have not been shown to date. The software Pymol (www.pymol.org) was used to analyze the structures and models, which are represented to scale, and to create the figure

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Fig. 2 Representative results of DNA unwinding assays to measure the helicase activity of full-length Twinkle WT and P66, in the absence of mtSSB. Unlike Twinkle WT that has salt-independent properties (lanes 3–6 in both gels in a), P66 only maintains a basal activity at 100 mM KCl (lanes 7–10 in lower gel in a). At 20 mM KCl, however, P66 enzymatic efficiency increases on average 7.5-fold (lanes 7–10 in upper gel in a), indicating that the C-terminal tail (see Fig. 1) stabilizes the enzyme under fluctuating ionic conditions. This stabilization has a negative effect on dsDNA unwinding, which may be overcome by interactions with mtSSB (see Fig. 3). (a) Example of radioactive signals provided by the unreacted, intact DNA unwinding substrates and partially or fully unwound reaction products. (b) Analysis of dsDNA unwinding activity from a composite of three independent experiments. The points represent the average values of unwound substrate, and the error bars represent the standard deviation

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10. To quantify the unwinding activity, consider the fraction of radiolabeled DNA species that is single stranded (60-mer oligonucleotide product, lower bands in Fig. 2a), following the formula % unwinding ¼ (VP/(VS + VP))  100, in which VP represents the volume of the 60-mer oligonucleotide product and VS the volume of unreacted, intact substrate in the sample lane of interest. A representative outcome of this quantification is shown in Fig. 2b. 3.2 Helicase Stimulation Assay

1. Prepare reaction mixtures 20 and 100 on ice and dispense 50μL into 1.5 mL microcentrifuge tubes. 2. Add 0, 25, 50, and 100 nM mtSSB (calculated as tetramers) into tubes (see Note 7). 3. Set two tubes aside which will be referred to as “boil –” and “boil +” controls. Add 100 nM mtSSB and no helicase in these tubes. The “boil –” control represents the unreacted, intact substrate bound by mtSSB (see Note 8), and the “boil +” control represents completely melted substrates after boiling. 4. Incubate tubes for 5 min in a water bath or heat block at 37  C (see Note 3). 5. Add 3.5 nM Twinkle WT or P66 (both calculated as hexamers) into the tubes with dispensed reaction mixtures (except “boil – ” and “boil +” controls), and incubate all tubes for 30 min at 37  C. 6. Follow the same procedures described in steps 4–10 of Subheading 3.1 to process the samples, reveal the results, and quantitate unwinding activity. Figure 3 shows a representative quantification of the experimental results.

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Notes 1. Various expression and purification strategies have been applied to obtain recombinant mtSSB, Twinkle WT, and P66 [15– 17]. We use the protocols described previously in [10, 18]. 2. The best results for these assays are obtained when the DNA unwinding substrate is used immediately after radiolabeling of the 50 -end of the 60-mer oligonucleotide, and its hybridization with the circular ss pBluescript KS(+) DNA. Different protocols may be used for the radiolabeling reaction, but we use that recommended by New England BioLabs with the T4 polynucleotide kinase and 32P-γ-ATP, followed by purification using a Bio-Rad microBiospin P-30 Tris–chromatography column (to remove unincorporated 32P-γ-ATP) [9]. It is imperative to test how much purified radiolabeled 60-mer oligonucleotide should be mixed with pBluescript KS(+) DNA (to guarantee

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Fig. 3 Effects of mtSSB on Twinkle unwinding activity. mtSSB does not stimulate the already elevated unwinding activity of P66 at 20 mM KCl, but increases that of Twinkle WT up to fivefold, independent of salt concentration. A substantial stimulation of P66 is observed at 100 mM KCl, a condition under which this variant does not function properly (see Fig. 2). These results suggest that P66 is stimulated passively by mtSSB in vitro. Further stimulation may be prevented by the lack of physical interactions between Twinkle and mtSSB, which is putatively mediated by the helicase C-terminal tail (see Fig. 1). The bars represent the average values of unwound substrate from three independent experiments, and the error bars represent the standard deviation

that all circular ssDNA molecules are hybridized by an oligonucleotide molecule) by adding increasing concentrations of the oligonucleotide to a fixed concentration of ssDNA circles. Mixtures should be incubated at 65  C for 1 h, and then 37  C for 30 min. Aliquots should be analyzed on a 22% native polyacrylamide gel to help determine the mixture that contains mostly hybridized oligonucleotide.

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3. To optimize the results, we do not recommend adding the enzyme in the reaction mixtures while the reaction tubes are still on ice, because Twinkle is more stable at 37  C [13]. 4. Although this is a short time, it is sufficient for the unreacted, intact DNA substrate to fully penetrate the gel. 5. If the radioactive signal of the DNA unwinding substrate is not strong enough due to radioisotope decay, the gel can be exposed overnight. If overnight exposure is not sufficient to obtain clear signals, prepare a fresh batch of substrate. 6. We use equipment and software from Amersham Biosciences, but any similar scanners and software can be as effective. 7. 100 nM mtSSB is expected to bind all available ssDNA of the DNA unwinding substrate in the reaction, based on a calculated binding site size of 55–60 nucleotides per tetramer. 8. This “boil –” control is important to show that mtSSB does not have intrinsic strand-displacement activity. Although some SSBs have such an activity in vitro, we have never observed it using the mtSSB concentrations used here.

Acknowledgments We would like to thank Dr. Roberto Negro for help with data collection, Murilo F. Othonicar for help with the figures, and Dr. Laurie S. Kaguni for supervision of this work and critical reading of the manuscript. A.P.C.R. was supported by a fellowship from the Conselho Nacional de Desenvolvimento Cientı´fico e Tecnolo´gico (CNPq, grant number 140788/2018-2). M.T.O. would like to acknowledge funding from the Fundac¸˜ao de Amparo a` Pesquisa do Estado de Sa˜o Paulo (FAPESP, grant number 2017/ 04372-0), and the CNPq (grant numbers 424562/2018-9 and 306974/2017-7). References 1. Ciesielski GL, Oliveira MT, Kaguni LS (2016) Animal mitochondrial DNA replication. Enzyme 39:255–292 2. Zeviani M, Donato DS (2004) Mitochondrial disorders. Brain 127(3):2153–2172 3. McKinney EA, Oliveira MT (2013) Replicating animal mitochondrial DNA. Genet Mol Biol 36(3):308–315 4. Wallace DC (1992) Mitochondrial genetics: a paradigm for aging and degenerative diseases? Science 256(5057):628–632

5. Nunnari J, Suomalainen A (2012) Mitochondria: In sickness and in health. Cell 148 (6):1145–1159 6. Korhonen JA, Pham XH, Pellegrini M, Falkenberg M (2004) Reconstitution of a minimal mtDNA replisome in vitro. EMBO J 23 (12):2423–2429 7. Peter B, Falkenberg M (2020) Twinkle and other human mitochondrial DNA helicases: structure, function and disease. Gene 11 (418):01–21

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8. Farr CL, Wang Y, Kaguni LS (1999) Functional interactions of mitochondrial DNA polymerase and single-stranded DNA-binding protein. J Biol Chem 274(21):14779–14785 9. Oliveira MT, Kaguni LS (2010) Functional roles of the N- and C-Terminal regions of the human mitochondrial single-stranded DNA-binding protein. PLoS One 5(10):e15379 10. Oliveira MT, Kaguni LS (2011) Reduced stimulation of recombinant DNA Polymerase γ and mitochondrial DNA (mtDNA) helicase by variants of mitochondrial single-stranded DNA-binding protein (mtSSB) correlates with defects in mtDNA replication in animal cells. J Biol Chem 286(47):40649–40658 11. Kaur P, Longley MJ, Pan H et al (2020) Singlemolecule level structural dynamics of DNA unwinding by human mitochondrial Twinkle helicase. J Biol Chem 295:5564–5576 12. Fuste´ JM, Shi Y, Wanrooij S et al (2014) In vivo occupancy of mitochondrial single-stranded DNA binding protein supports the strand displacement mode of DNA replication. PLoS Genet 10(12):e1004832 13. Ziebarth TD, Farr CL, Kaguni LS (2007) Modular architecture of the hexameric human

mitochondrial DNA helicase. J Mol Biol 367 (5):1382–1391 14. Kaguni LS, Oliveira MT (2016) Structure, function and evolution of the animal mitochondrial replicative DNA helicase. Crit Rev Biochem Mol Biol 51(1):53–64 15. Korhonen JL, Gaspari M, Falkenberg M (2003) Twinkle has 5’ -> 3’ DNA helicase activity and is specifically stimulated by mitochondrial single-stranded DNA-binding protein. J Biol Chem 278(49):48627–48632 16. Longley MJ, Humble MM, Sharief FS, Copeland WC (2010) Disease variants of the human mitochondrial DNA helicase encoded by C10orf2 differentially alter protein stability, nucleotide hydrolysis, and helicase activity. J Biol Chem 285(39):29690–29702 17. Longley MJ, Smith LA, Copeland WC (2009) Preparation of human mitochondrial singlestranded DNA-binding protein. Methods Mol Biol 554:73–85 18. Makowska-Grzyska MM, Ziebarth TD, Kaguni LS (2010) Physical analysis of recombinant forms of the human mitochondrial DNA helicase. Methods 51(4):411–415

Chapter 21 Measuring the Complex Effects of the Single-Stranded DNA-Binding Protein gp2.5 on Primer Synthesis and Extension by the Bacteriophage T7 Replisome Alfredo J. Hernandez Abstract The single-stranded DNA-binding protein gp2.5 of bacteriophage T7 plays myriad functions in the replication of phage genomes. In addition to interacting with ssDNA, gp2.5 binds to the T7 DNA polymerase and primase/helicase proteins, regulating their enzymatic activities. Here we describe in vitro methods to examine the effects of gp2.5 on primer synthesis and extension by the T7 replisome. Key words DNA replication, Primase, Polymerase, Denaturing, Acrylamide

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Introduction Bacteriophage T7 has evolved as an efficient machinery for DNA replication that nevertheless performs all the DNA transactions of more complex organisms [1–3]. The T7 genome encodes all but one of its replication proteins. In vitro, only four proteins are required to reconstitute coordinated leading- and lagging-strand DNA synthesis: the DNA polymerase, gp5, and its processivity factor, E. coli thioredoxin (Trx); the bifunctional primase-helicase gp4; and the single-stranded DNA-binding protein, gp2.5 [4, 5]. The simple composition of the T7 replisome makes it an attractive model system to study conserved biochemical mechanisms involved in DNA replication. gp2.5 regulates the activities of T7 DNA polymerase, the primase/helicase gp4, and it is crucial for coordinating the rates of DNA synthesis of the leading and lagging strands [6]. The presence of gp2.5 enhances primer formation and RNA-dependent DNA synthesis using ss M13 DNA as a substrate [7]. Using short, synthetic ssDNA templates, gp2.5 enhances the efficiency of primer synthesis by reducing the dissociation of oligoribonucleotide intermediates from the primase/helicase active site, increasing the

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_21, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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fraction of full-length primer produced. While the presence of gp2.5 tempers the amount of primer synthesis, decreasing the proportion of primers extended by DNA polymerase by approximately twofold, primers are utilized with faster kinetics than in reactions lacking gp2.5, or those that contain either E. coli SSB or a mutant version of gp2.5 which lacks its acidic C-terminal tail (gp2.5 Δ26). The decrease in primer utilization may allow the coordination of leading- and lagging-strand DNA synthesis by modulating the timing of initiation/termination events during synthesis of Okazaki fragments [8]. Here we provide methods to examine the effect of gp2.5 on primer synthesis and extension by T7 DNA replication enzymes. The electrophoretic assays described here are highly sensitive and can be performed in a relatively short time. However, they require the use of radioactive nucleotide substrates and are low-throughput assays. While the assays here are designed for a kinetic examination of these reactions, they can be easily adapted to end-point assays.

2

Materials Prepare all solutions using Milli-Q water and reagent-grade chemicals, and filter sterilize using 0.2 μm polyethersulfone membranes (e.g., Nalgene Rapid-Flow™ or similar). Follow all institutional regulations for safe use and disposal of radioactive material, including but not limited to usage of personal protective equipment, such as gloves, safety glasses, lab coat, and appropriate acrylic shields.

2.1 Effect of gp2.5 on Primer Synthesis

1. Purified T7 gp2.5, as reported [9]. 2. Purified T7 gp4, as reported [8] (see Notes 1 and 2). 3. 5
Reaction buffer: 200 mM HEPES-KOH, pH 7.5, 250 mM potassium glutamate, 50 mM DTT, 0.5 mM EDTA. 4. 10 mM ATP. 5. 10 mM CTP. 6. 0.1 M MgCl2. 7. [α-32P CTP], 3000 Ci/mmol, 10 mCi/mL. 8. [γ-32P ATP], 3000 Ci/mmol, 10 mCi/mL. 9. Single-stranded DNA containing a single T7 primase recognition site (see Note 3). 10. 25 mM dNTPs. 11. Formamide dye buffer: 93% (v/v) Formamide, 50 mM EDTA, 0.01% xylene cyanol, and 0.01% bromophenol blue. 12. Rapid-quench flow apparatus. 13. 10 mM Tris pH 7.5, 1 mM EDTA.

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14. Quench solution: 250 mM EDTA, 0.25% SDS. 15. 0.2 N NaOH. 16. 0.3 N H3PO4. 17. Methanol. 18. 10 mL Syringes. 19. 1 mL Syringes. 20. 20G needles. 2.2 Effect of gp2.5 on Primer Extension

1. Purified T7 gp2.5, as reported [9]. 2. Purified T7 gp4, as reported [8]. 3. Purified T7 DNA polymerase, as reported [10] (see Note 4). 4. 5
Reaction buffer: 200 mM HEPES-KOH pH 7.5, 250 mM potassium glutamate, 50 mM DTT, 0.5 mM EDTA. 5. 10 mM ATP. 6. 10 mM CTP. 7. 0.1 M MgCl2. 8. [α-32P CTP], 3000 Ci/mmol, 10 mCi/mL. 9. [γ-32P ATP], 3000 Ci/mmol, 10 mCi/mL. 10. Single-stranded DNA containing a single T7 primase recognition site. 11. 25 mM dNTPs. 12. Formamide dye buffer: 93% (v/v) Formamide, 50 mM EDTA, 0.01% xylene cyanol, and 0.01% bromophenol blue. 13. Rapid-quench flow apparatus. 14. 10 mM Tris pH 7.5, 1 mM EDTA. 15. Quench solution: 250 mM EDTA, 0.25% SDS. 16. 0.2 N NaOH. 17. 0.3 N H3PO4. 18. Methanol. 19. 10 mL Syringes. 20. 1 mL Syringes. 21. 20G Needles.

2.3 Denaturing Polyacrylamide Gel Electrophoresis

1. 10
TBE: 1 M Tris base, 1 M boric acid, 20 mM EDTA. 2. Denaturing acrylamide solution: 25% Acrylamide (19:1 acrylamide:bis-acrylamide), 3 M urea, 1
TBE. 3. 10% (w/v) APS. 4. N,N,N0 ,N0 -tetramethylethane-1,2-diamine (TEMED). 5. Sequencing gel electrophoresis apparatus.

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6. Sequencing gel glass plates. 7. Sequencing gel combs and spacers. 8. Metal binder clips. 9. Chromatography paper. 10. Plastic cling wrap. 11. Gel dryer. 2.4

Data Analysis

1. Phosphorimager and storage screens. 2. Phosphorimager scanner and analysis software. 3. Spreadsheet software. 4. Nonlinear, least squares fitting software.

3

Methods

3.1 Effect of gp2.5 on Primer Synthesis

1. Aliquot 2 μL of formamide dye buffer into eight 1.5 mL polypropylene tubes.

3.1.1 Multiple-Turnover Primer Synthesis: Manual Sampling

2. Prepare a 20 μL mix composed of 1
buffer, 0.1 mM ATP, 0.1 mM CTP, 0.2 mCi/mL [α-32P] CTP, 0.1 μM ssDNA template, 3 μM gp2.5, and 0.1 μM gp4 hexamer (0.6 μM monomer). 3. Incubate the mixture at room temperature for 5 min. 4. Remove 2 μL of the mixture using a micropipette and mix with 2 μL formamide dye buffer in a 1.5 mL tube prepared in step 1 for the no-reaction control (time zero). 5. Add 2 μL of 0.1 M MgCl2 to reaction mixture (to a final concentration of 10 mM). 6. Manually withdraw 2 μL samples of the reaction mixtures using a micropipette at 10-s intervals and mix with formamide dye predispensed in 1.5 mL tubes prepared in step 1. 7. Heat samples at 95  C for 5 min.

3.1.2 Multiple-Turnover Primer Synthesis Reaction: Rapid-Quench Instrument

1. Prepare 400 μL of solution A, containing 1
buffer, 6 μM gp2.5, 0.2 μM gp4 hexamer, 0.2 μM ssDNA template, 0.6 mM dNTPs, 0.2 mM ATP, and CTP (including 0.4 mCi/mL [α-32P] CTP) (see Notes 5 and 6). 2. Prepare 400 μL of solution B, containing 1
buffer and 20 mM MgCl2. 3. Turn on rapid-quench flow instrument; move the syringe driver to the home position. 4. Open buffer syringe position to “LOAD.” 5. Attach a 10 mL syringe containing buffer to the drive syringe port. Change syringe knob position to “LOAD.”

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6. Using 10 mL syringes, fill instrument buffer reservoirs A and B with 10 mM Tris, pH 7.5, and 1 mM EDTA and fill syringe C with quench solution. Remove air bubbles by pushing plungers in and out. 7. Change knob positions to “FIRE.” Adjust the position of the syringe driver bar. Lower the syringe until buffer comes out the exit loop. 8. Attach exit line to vacuum. Change sample knobs to “FLUSH.” Close buffer syringe knobs. Turn on vacuum, dip the two flush loops in H2O for 10 s, and then dip in MeOH for 10 s. Dry loops by sucking air for ~15 s, or until dry. 9. Change sample knob position to “LOAD.” Place solution A into 1 mL syringe (see Note 7). Plug syringe into one of the sample load ports. Repeat for solution B, and place in the opposite sample load port. 10. On main menu, type 1 for quench-flow run. Enter reaction time, in seconds (see Note 8), and press “ENTER.” The reaction loop number to use will be displayed. Switch to required reaction loop. Repeat wash procedure in step 8. 11. Turn sample knobs to “LOAD.” Load reaction mixes in sample loops until just outside the central mixing compartment. Turn all knobs to “FIRE.” Confirm that all knobs are in the correct position. 12. Place 1.5 mL tube onto end of exit loop. Press “GO” to run the reaction. Repeat wash procedure in step 8. Reload buffer syringes A and B until they hit the driver. 13. Repeat steps 9–12 for each desired time point. For the no-reaction control, do not load solution containing the start reagent (i.e., MgCl2 in this case). 14. When finished collecting samples, return buffer syringe driver to its home position. 15. Remove reaction mixture syringes. Turn sample loading knob to “LOAD” position. Under vacuum, wash each sample loop with 10 mL 0.2 N NaOH, followed by 10 mL 0.3 N H3PO4 and 10 mL MeOH. 16. Turn syringe knobs to “LOAD” position. Press down on all three syringe plungers to retrieve reaction and quench buffers. Wash syringes with 5 mL H2O at least twice. 17. Fill syringes again with 5 mL H2O and turn knobs to “FIRE” position. Change knob positions and turn on vacuum to remove H2O. Flush as in step 8. Turn off instrument. 18. Mix samples 1:1 with formamide dye, and heat at 95  C for 5 min.

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3.1.3 Single-Turnover Primer Synthesis Reaction: Rapid-Quench Instrument

1. Prepare 400 μL of solution A, containing 1
buffer, 40 μM gp2.5, 3 μM gp4 hexamer, 6 μM ssDNA template, 0.6 mM dNTPs, 2 mM CTP, and 2 nM [γ-32P] ATP (see Note 9). 2. Follow steps 2–18 of Subheading 3.1.2.

3.2 Effect of gp2.5 on Primer Extension

1. Aliquot 2 μL of formamide dye buffer into 8 1.5 mL polypropylene tubes.

3.2.1 Multiple-Turnover Primer Synthesis and Extension Reaction: Manual Sampling

2. Prepare a 20 μL mix composed of 1
buffer, 0.1 mM ATP, 0.1 mM CTP, 0.2 mCi/mL [α-32P] CTP, 0.1 μM ssDNA template, 3 μM gp2.5, 0.5 μM T7 DNA polymerase, and 0.1 μM gp4 hexamer (0.6 μM monomer). 3. Follow steps 3–7 of Subheading 3.1.1.

3.2.2 Multiple-Turnover Primer Synthesis and Extension Reaction: Rapid-Quench Instrument

1. Prepare 400 μL of solution A, containing 1
buffer, 6 μM gp2.5, 1 μM T7 DNA polymerase, 0.2 μM gp4 hexamer, 0.2 μM ssDNA template, 0.6 mM dNTPs, 0.2 mM ATP, and CTP (including 0.4 mCi/mL [α-32P] CTP). 2. Follow steps 2–18 of Subheading 3.1.2.

3.2.3 Single-Turnover Primer Synthesis and Extension Reaction Using a Rapid-Quench Instrument (See Note 10)

1. Prepare 400 μL of solution A, containing 1
buffer, 40 μM gp2.5, 30 μM T7 DNA polymerase, 3 μM gp4 hexamer, 6 μM ssDNA template, 0.6 mM dNTPs, 1 mM CTP, and 2 nM [γ-32P] ATP.

3.3 Denaturing Polyacrylamide Gel Electrophoresis

1. Wash glass plates (see Note 11) with water and detergent followed by ethanol. Wipe dry.

2. Follow steps 2–18 of Subheading 3.1.2.

2. Assemble the glass plates using 0.4 mm spacers according to the manufacturer’s instructions (see Note 12). 3. Add 0.3 mL 10% APS and 30 μL TEMED to 30 mL denaturing acrylamide gel solution. 4. Pour denaturing acrylamide gel solution into gels and insert sample well comb. Allow 1 h for polymerization. 5. Remove comb, place gel onto apparatus, and fill the upper and lower buffer chambers with 1
TBE. 6. Flush each well with 1
TBE using a syringe or micropipette. 7. Pre-run gel for 30 min at 25 W. Temperature of plates should be ~55  C. 8. Load samples. Run at 25 W for 45 min or until bromophenol blue migrates halfway down the gel. 9. Transfer the entire gel onto chromatography paper and cover with plastic wrap (see Note 13). 10. Dry gel using a gel drier (see Note 14).

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Data Analysis

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1. Prepare a dilution series of the remainder of the reaction mixture and spot the same volume loaded on the gel onto chromatography paper (see Note 15). 2. Expose dry gel and dilution series onto a phosphorimager screen. 3. Scan image after an appropriate exposure time (see Note 16) and quantify bands and dilution series using software for gel analysis. 4. Use dilution series to construct a standard curve of phosphorimager counts as a function of CTP or ATP concentration. 5. Determine the concentration of products of interest by comparing their intensity with the standard curve. 6. Construct a reaction progress curve by plotting the concentration of product as a function of time (see Note 17). 7. Determine the rate of product formation by fitting progress curves using commercially available software. The details of the operation will vary with the software employed (see Note 18).

4

Notes 1. We routinely purify T7 replication proteins in an untagged form. It is also possible to use N-terminal His-tagged variants to simplify purification schemes. Placement of a His-tag at the C-terminus is not recommended, as it is deleterious in genetic and in vitro assays. Proteolytic removal of affinity tags can circumvent this problem, but cleavage is often inefficient. 2. Flash-freeze small aliquots of purified gp4 in 25 mM Tris–HCl pH 7.5, 50 mM NaCl, 0.1 mM EDTA, 1 mM TCEP, and 10% glycerol using liquid nitrogen and store at –80  C. In our hands, storage of purified gp4 under previously published conditions (20 mM potassium phosphate pH 7.4, 0.1 mM EDTA, 1 mM DTT containing 50% glycerol at –20  C) results in a gradual reduction of specific activity. 3. The T7 primase-helicase gp4 catalyzes the formation of primers for DNA synthesis at specific DNA sequences called primase recognition sites, or PRS (50 -GTC-30 ) [11]. The cytosine in the PRS is essential for recognition of the site, but it is not copied into the product. Single-stranded DNA containing a single T7 PRS encoding a tetraribonucleotide primer (50 -GGGTC-30 , 50 -GTGTC-30 , 50 -TGGTC-30 ; PRS is underlined) can be purchased from a variety of DNA synthesis companies (a pentamer is the minimum template length for primer synthesis, as the cryptic C is essential). If the primer is to be extended by DNA polymerase, additional nucleotides must be present at the 50 end of the template.

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4. T7 DNA polymerase is a tight (Kd ¼ 5 nM) 1:1 complex of T7 gp5 and E. coli thioredoxin [12]. We typically reconstitute the holoenzyme by mixing purified gp5 and thioredoxin at a 1:10 molar ratio and use as is. It is also possible to isolate the reconstituted holoenzyme by chromatography through a hydroxyapatite column [10]. 5. Reaction mixtures for rapid quench experiments are prepared at a 2
concentration. These are mixed 1:1 upon initiation of the reaction to give a 1
concentration. 6. We describe operation of a KinTek RQF-3, but rapid quench instruments from other manufacturers function similarly. 7. Use a syringe with a 20G needle attached to facilitate withdrawal of the sample from 1.5 mL tubes. 8. The range of timescales investigated should be empirically determined. In the reaction catalyzed by gp4, formation of full-length primer products is evident after a few milliseconds [8]. 9. In single-turnover experiments, a low concentration of labeled substrate is mixed with saturating concentrations of enzyme (as well as DNA, CTP, and gp2.5 in this case). 10. The extension products accumulate in a time regime that allows for manual sampling of the reaction. If the experimental objective is to examine only extension products, proceed with the manual protocol. 11. We typically use gel plates that are 16 cm wide and 26 cm long. 12. Clamp gel plates using black metal binder clips. 13. Gels containing a high percentage of acrylamide do not stick well to chromatography paper. Transfer gel first onto plastic cling wrap and then very carefully place onto chromatography paper for drying. 14. Another disadvantage of gels containing a high percentage of acrylamide is that they tend to crack in vacuum-powered gel dryers. Lowering the drying temperature from 80 to 65  C leads to less cracking of gels. Alternatively, the gels can be exposed to phosphorimager screens without drying. 15. Fivefold dilutions of the sample mixture, using TE buffer (10 mM Tris–HCl, pH 8.0, 1 mM EDTA), work well, spanning a range of 1:25 to 1:15,625. This will serve as the standard curve to quantify the concentration of products formed during the reaction. Spot the same volume of sample onto chromatography paper as the volume loaded on the gel, i.e., spot 4 μL of the dilution if 4 μL of sample was loaded.

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16. Exposure time can range from 30 min to overnight, and depends on the specific activity of the radiolabeled nucleotide. For best results, do not use radiolabeled nucleotides with reference dates older than a month. 17. Account for the correct number of labels incorporated for each product species if using CTP as the labeled nucleotide. 18. Determine the rate of product formation of multiple-turnover primer synthesis/extension reactions by fitting progress curves to a linear equation. If there is significant curvature, use the initial rate method by fitting the data to a single-exponential function: y ¼ A(e–kt), where y is the product concentration, A is the reaction amplitude, k is the observed rate constant, and t is time, in seconds [13]. The slope of the tangent line at time ¼ 0 is the initial rate. Fit pre-steady-state primer synthesis reactions to the pre-steady-state burst equation y ¼ A(e–kburst) + ratesst, where A is the reaction amplitude, kburst is the observed rate constant for the pre-steady-state burst, ratess is the steady-state rate (¼kcat[Enzyme]), and t is time, in seconds [14]. Fit progress curves of single-turnover primer synthesis/extension reactions to a single-exponential function y ¼ A(e–kt). It is also possible to fit the data by numerical integration using commercially available software [15]. References 1. Hamdan SM, Richardson CC (2009) Motors, switches, and contacts in the replisome. Annu Rev Biochem 78:205–243. https://doi.org/ 10.1146/annurev.biochem.78.072407. 103248 2. Lee SJ, Richardson CC (2011) Choreography of bacteriophage T7 DNA replication. Curr Opin Chem Biol 15:580–586. https://doi. org/10.1016/j.cbpa.2011.07.024 3. Kulczyk AW, Richardson CC (2016) The Replication System of Bacteriophage T7. Enzyme 39:89–136. https://doi.org/10.1016/bs.enz. 2016.02.001 4. Lee J, Chastain PD 2nd, Griffith JD, Richardson CC (2002) Lagging strand synthesis in coordinated DNA synthesis by bacteriophage T7 replication proteins. J Mol Biol 316:19–34. https://doi.org/10.1006/jmbi.2001.5325 5. Lee J, Chastain PD 2nd, Kusakabe T, Griffith JD, Richardson CC (1998) Coordinated leading and lagging strand DNA synthesis on a minicircular template. Mol Cell 1:1001–1010 6. Hernandez AJ, Richardson CC (2019) Gp2.5, the multifunctional bacteriophage T7 singlestranded DNA binding protein. Semin Cell

Dev Biol 86:92–101. https://doi.org/10. 1016/j.semcdb.2018.03.018 7. Nakai H, Richardson CC (1988) The effect of the T7 and Escherichia coli DNA-binding proteins at the replication fork of bacteriophage T7. J Biol Chem 263:9831–9839 8. Hernandez AJ, Lee SJ, Richardson CC (2016) Primer release is the rate-limiting event in lagging-strand synthesis mediated by the T7 replisome. Proc Natl Acad Sci U S A 113:5916–5921. https://doi.org/10.1073/ pnas.1604894113 9. Kim YT, Tabor S, Bortner C, Griffith JD, Richardson CC (1992) Purification and characterization of the bacteriophage T7 gene 2.5 protein. A single-stranded DNA-binding protein. J Biol Chem 267:15022–15031 10. Tabor S, Huber HE, Richardson CC (1987) Escherichia coli thioredoxin confers processivity on the DNA polymerase activity of the gene 5 protein of bacteriophage T7. J Biol Chem 262:16212–16223 11. Tabor S, Richardson CC (1981) Template recognition sequence for RNA primer synthesis by

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gene 4 protein of bacteriophage T7. Proc Natl Acad Sci U S A 78:205–209 12. Akabayov B, Akabayov SR, Lee SJ, Tabor S, Kulczyk AW, Richardson CC (2010) Conformational dynamics of bacteriophage T7 DNA polymerase and its processivity factor, Escherichia coli thioredoxin. Proc Natl Acad Sci U S A 107:15033–15038. https://doi.org/10. 1073/pnas.1010141107 13. Cao W, De La Cruz EM (2013) Quantitative full time course analysis of nonlinear enzyme

cycling kinetics. Sci Rep 3:2658. https://doi. org/10.1038/srep02658 14. Johnson KA (2013) A century of enzyme kinetic analysis, 1913 to 2013. FEBS Lett 587:2753–2766. https://doi.org/10.1016/j. febslet.2013.07.012 15. Johnson KA (2009) Fitting enzyme kinetic data with KinTek global kinetic explorer. Methods Enzymol 467:601–626. https://doi. org/10.1016/S0076-6879(09)67023-3

Chapter 22 Strand Displacement and Unwinding Assays to Study the Concerted Action of the DNA Polymerase and SSB During Phi29 TP-DNA Replication Alicia del Prado and Margarita Salas Abstract The Bacillus subtilis phage Phi29 has a linear double-stranded DNA with a terminal protein (TP) covalently linked to each 50 end (TP-DNA). Phi29 single-stranded DNA-binding protein (SSB) is encoded by the viral gene 5 and binds the ssDNA generated during the Phi29 genome replication, stimulating the DNA elongation rate. Here, we describe some protocols to evaluate the effect of Phi29 SSB mutants on the DNA elongation rate and their unwinding activity during replication by Phi29 DNA polymerase using as substrate TP-DNA and also singly primed M13 DNA. Key words Single-stranded DNA-binding protein, DNA binding, Helix-destabilization activity, DNA replication, Bacteriophage Phi29, DNA polymerase

1

Introduction Bacteriophage Phi29 infects the bacteria Bacillus subtilis and has served as a model to study the replication with a terminal protein (TP). Its genome is a linear double-strand DNA (dsDNA) with a TP attached at each 50 DNA end (TP-DNA) [1]. Phi29 DNA polymerase is the enzyme responsible for viral DNA replication, and due to its inherent properties like high processivity, strand displacement capacity, and high fidelity is able to replicate the whole genome without the need of processivity or unwinding factors, being an incredible biotechnological tool for isothermal DNA amplification [2–5]. Phi29 single-stranded DNA-binding protein (SSB), encoded by the viral gene 5, binds the ssDNA generated during the Phi29 replication, protecting DNA from nucleases and preventing unproductive binding of Phi29 DNA polymerase to ssDNA [6, 7]. It has been described that during replication of TP-DNA and primed M13 ssDNA, Phi29 SSB stimulates the dNMP incorporation rate

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_22, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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[8] and due to its DNA unwinding activity or helix destabilization activity it is also able to displace oligonucleotides annealed to M13 ssDNA [8, 9]. The Phi29 SSB residue Tyr57 is involved in binding the ssDNA, being the mutant derivative Y57A not only impaired in binding ssDNA but also in the unwinding activity and in the stimulation of the DNA elongation [10]. Here, we provide two protocols to evaluate the effect of Phi29 SSB mutants on the DNA elongation rate. The first protocol is carried out under conditions in which the strand opening is impaired, for that we use a Phi29 DNA polymerase mutant (D12A/D66A) that has two catalytic aspartic acids of the exonuclease active site mutated into alanine [11]. This variant lacks the exonuclease activity and its strand displacement capacity is also impaired. In these conditions the role of the SSB in stimulating the elongation rate during strand displacement DNA replication is easier to detect than when the DNA polymerase has its intrinsic strand displacement capacity. The second protocol for this goal uses as substrate the M13 DNA hybridized to a 17-mer oligonucleotide to get a primed M13 DNA. In this assay, the first replication round does not require the strand displacement, but when the DNA polymerase reaches the 50 end of the 50 terminus of the oligonucleotide primer, it is needed to couple the polymerization with the strand displacement to continue with the replication. Finally, we also provide a protocol to evaluate the unwinding activity of Phi29 SSB; for that we analyze the helix destabilization activity of the SSBs testing if they are able to displace an oligonucleotide annealed to M13 ssDNA.

2

Materials Prepare all solutions using distilled water. Prepare and store all the buffers at room temperature (except the 10 reaction buffer that must be stored at 20  C).

2.1 Incubation Reaction

1. 10 Reaction buffer: 500 mM Tris–HCl pH 7.5, 0.01 M dithiothreitol (DTT), 40% (v/v) glycerol, 1 mg/mL bovine serum albumin (BSA). 2. Buffer dissolution with Tween (BDT): 25 mM Tris–HCl pH 7.5, 0.1 M NaCl, 0.05% Tween. 3. 10 Hybridization buffer: 0.6 M Tris–HCl pH 7.5, 2 M NaCl. 4. Stop solution: 0.2% Sodium dodecyl sulfate (SDS), 20 mM EDTA, 50 mM Tris–HCl, pH 7.5. 5. Sephadex G-50 SDS: 55 g Sephadex G50, 50 mL 1 M Tris– HCl pH 7.5, 10 mL 10% SDS, and water up to 1 L. Autoclave before using.

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6. 250 mM Ammonium sulfate. 7. 100 mM MgCl2. 2.2 8% Polyacrylamide Gels, 0.1% SDS (300  250  0.5 mm)

1. Resolving gel buffer: 8.1 mL Water, 4 mL acrylamide/bis solution (37.5:1), 7.5 mL 1 M Tris–HCl pH 8.8, 200 μL 10% SDS, 170 μL 10% ammonium persulfate (APS), and 15 μL TEMED. 2. Stacking gel buffer: 7.3 mL Water, 1.25 mL acrylamide/bis solution (37.5:1), 1.25 mL 1 M Tris–HCl pH 6.8, 100 μL 10% SDS, 150 μL 10% APS, and 15 μL TEMED. 3. Running buffer Tris–glycine-SDS: 3 g Trizma base (99% titration), 14.4 g glycine, 5 mL 20% SDS, and water up to 1 L (see Note 1).

2.3 Alkaline 0.7% Agarose Gels

1. 6 Alkaline loading buffer: 30% Glycerol, 0.25% (w/v) bromophenol blue, and 0.25% xylene cyanol (w/v). 2. 10 Alkaline agarose gel electrophoresis buffer: 500 mM NaCl, 10 mM EDTA. Add 100 mL 5 M NaCl and 20 mL 0.5 M EDTA (pH 8) in 700 mL water, and then adjust the final volume to 1 L. Dilute in water before using. 3. Running buffer: 30 mM NaOH, 1 mM EDTA. Add 30 mL 5 M NaOH and 10 mL 0.5 M EDTA (pH 8) in 1 L of water, and then adjust the final volume to 5 L.

2.4 Nucleotides and DNAs

1. 1 mM Unlabeled dNTPs. 2. Radiolabeled nucleotides: [α-32P]dATP (3000 Ci/mmol) and [γ-32P]ATP (3000 Ci/mmol). 3. M13mp18 ssDNA isolated as described [10]. 4. Universal primer (17 mer): 50 GTTTTCCCAGTCACGAC 30 . 5. Phi29 TP-DNA isolated as described [12].

3

Methods The experiments are performed with radiolabeled nucleotides; all safety regulations and waste disposal regulations must be followed.

3.1 Strand Displacement Coupled to M13-DNA Replication

1. Make primed M13mp18 by hybridizing M13-DNA with the universal primer (17 mer) at 70  C for 5 min, in a mixture containing 200 mM NaCl, 80 nM M13-DNA, 700 nM universal primer, and water up to a final volume of 150 μL. Allow mixture to slowly cool to room temperature (see Note 2). 2. Prepare incubation mixture on ice in 25 μL volume by adding 2.5 μL of 10 reaction buffer (final concentration: 50 mM Tris–HCl pH 7.5, 1 mM DTT, 4% (v/v) glycerol, 0.1 mg/mL

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BSA), 1 μL 1 mM each dNTP (final concentration 40 μM), 0.1 μL [α-32P]dATP (1 μCi), 1.45 μL of the hybridized primed M13mp18 ssDNA (final concentration 5 nM), 2 μL of a dilution of 50 ng/μL of Phi29 DNA polymerase in BDT (60 mM), 1 μL of a dilution of 10 μg/μL of the Phi29 SSB or mutants in BDT (30 μM), and water up to 25 μL (see Note 3). 3. The reaction starts by adding 2.5 μL 100 mM MgCl2 (10 mM final) and the samples are incubated for 5 or 40 min at 30  C. 4. Stop the reaction by adding 30 μL 10 mM EDTA and 0.1% SDS. 5. Remove unincorporated radioactive nucleotides using Sephadex G-50 spin columns. 6. Fill a 1 mL syringe lightly with gauze (the lower end). 7. Add the Sephadex G-50 SDS into the column and centrifuge it during 5 min at 2500 g to pack the column and remove residual buffer (see Note 4). 8. Apply the sample carefully to the center of the column bed. 9. Spin column for 5 min at 2500 g. Recover the eluted. 10. Dry the samples in a SpeedVac centrifuge and resuspend it in water (see Note 5). 11. Take 15 μL of the resuspended samples and add 2.1 μL 5 M NaOH (for a final concentration of 0.7 M NaOH) to denature the DNA. Add 3 μL 6 alkaline loading buffer and run the samples in an alkaline 0.7% agarose gel. 12. Add 0.7 g agarose in 10 mL 10 alkaline agarose gel electrophoresis buffer with 90 mL of H2O. The agarose gels are prepared in an alkaline solvent to denature the DNA [13]. 13. Heat it in a microwave oven until the agarose is dissolved. 14. When the temperature decreases (up to around 50  C), pour the agarose gel into the mold with the comb. 15. Wait until the agarose has completely solidified. 16. Remove the comb and mount the gel in the electrophoresis tank with running buffer. 17. Add running buffer until the gel is completely covered. 18. Load the sample mixture into the wells of the gel. 19. Run the gel at 75 V for 2 or 2.5 h (see Note 6). 20. Turn off the electrical current and dry the gel (see Note 7). 21. For the autoradiography use a cassette that avoids the contact with light and films (film of 13  18 cm) (see Note 8). 22. Develop your film and analyze your results (see Note 9) (Fig. 1).

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Fig. 1 Strand displacement coupled to M13-DNA replication. The assay was performed as described in Subheading 3.1, using 60 nM of Phi29 DNA polymerase, 5 nM of primed M13 ssDNA, and 30 μM SSB wild type or mutant. Samples were incubated for the indicated time at 30  C and analyzed by 0.7% alkaline agarose gel electrophoresis followed by autoradiography. (Reproduced with permission from de la Torre I, Quin˜ones V, Salas M, del Prado A (2019) Tyrosines involved in the activity of Phi29 single-stranded DNA binding protein. PLoS ONE 14(5): e0217248. https://doi.org/10.1371/journal.pone.0217248) 3.2 TP-DNA Replication with an Exonuclease-Deficient Phi29 DNA Polymerase

1. Prepare incubation mixture on ice in 25 μL volume by adding 2.5 μL of 10 reaction buffer, 1 μL 500 μM each dNTPs (final concentration 20 μM), 0.1 μL [α-32P]dATP (1 μCi), 2 μL 250 mM ammonium sulfate (final concentration 20 mM), 1.5 μL 335 ng/μL TP-DNA (1.6 nM final), 2 μL of a 10 ng/ μL dilution in BDT of ϕ29 DNA polymerase wild-type or exo-mutant D12A/D66A (13 mM final), 2 μL of a 5 ng/μL dilution in BDT of TP (13 nM final), 1 μL of a 10 μg/μL dilution in BDT of ϕ29 SSB wild type or mutants (30 μM final), and water up to 25 μL (see Note 10). The exo-mutant D12A/ D66A that lacks two of the catalytic aspartic acids is described in [11]. 2. The reaction starts by adding 2.5 μL 100 mM MgCl2 (10 mM final) and the samples are incubated for 30, 60, or 90 min at 30  C (see Notes 11 and 12).

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Fig. 2 TP-DNA replication with the exonuclease-deficient Phi29 DNA polymerase mutant. The assay was performed as described in Subheading 3.2, using 13 nM of Phi29 DNA polymerase exo-mutant D12A/D66A, 1.6 nM of TP-DNA, and 30 μM SSB. Samples were incubated for the indicated times at 30  C and analyzed by 0.7% alkaline agarose gel electrophoresis followed by autoradiography. ((Reproduced with permission from de la Torre I, Quin˜ones V, Salas M, del Prado A (2019) Tyrosines involved in the activity of Phi29 single-stranded DNA binding protein. PLoS ONE 14(5): e0217248. https://doi.org/10.1371/journal. pone.0217248)

3. Stop the reaction by adding 30 μL 10 mM EDTA and 0.1% SDS. 4. Remove unincorporated radioactive nucleotides using Sephadex G-50 spin columns, as described in steps 6–9 of Subheading 3.1. 5. Dry the samples in a SpeedVac centrifuge and resuspend it in water (see Note 5). 6. Follow steps 11–21 of Subheading 3.1 to denature DNA, run the samples in an alkaline 0.7% agarose gel, dry the gel, and autoradiograph it. 7. Develop the film and analyze the results (Fig. 2). 3.3

Unwinding Assay

1. Prepare oligonucleotide labeling reaction on ice in a final volume of 30 μL. Add 590 nM oligonucleotide (M13 universal primer, 17 mer), 20 U T4 polynucleotide kinase, 70 mM Tris– HCl pH 7.5, 10 mM MgCl2, 5 mM DTT, and 10 μCi [γ-32P] ATP (3000 Ci/mmol).

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2. Incubate for 45 min at 37  C and then for 5 min at 65  C to inactivate the enzyme. 3. To remove the unincorporated radiolabeled nucleotides, use an oligo spin column of Sephadex G-25 in buffer 10 mM Tris– HCI pH 7.5, 1 mM EDTA, and 100 mM NaCl. First, centrifuge it for 1 min at 1000 g to pack the column and remove residual buffer (see Note 13). 4. Apply the sample carefully in the center of the column bed and spin the column for 4 min at 1000 g. Recover the eluate. 5. Hybridize M13-DNA with the 50 -labeled universal primer (17 mer) at 60  C for 5 min, in a mixture containing 200 mM NaCl, 60 mM Tris–HCl pH 7.5, 90 nM M13DNA, 60 nM universal primer, and water up to a final volume of 60 μL. Allow mixture to slowly cool to room temperature (see Note 14). 6. Prepare incubation mixture on ice in 12.5 μL volume by adding 2.5 μL of 10 reaction buffer, 1 μL (2 nM) labeled primed M13mp18 ssDNA, 2 μL of a dilution in BDT of ϕ29 SSB wild type or mutants (to final concentrations of 10, 20, 40, or 80 μM), and water up to 12.5 μL. 7. Incubate for 60 min at 37  C. 8. Stop the reaction with 1.25 μL of 0.25% (w/v) bromophenol blue, 0.25% (w/v) xylene cyanol, 30% (v/v) glycerol, and 0.5% SDS. 9. Subject the samples to electrophoresis in 8% acrylamide gels containing 0.1% SDS at 4  C. Mount gel glass plates (300  250  0.5 mm) with the spacers. Prepare the resolving gel buffer and pour in between plates (see Note 15). 10. When polymerized, prepare the stacking gel buffer and pour on top of resolving gel. Immediately put the desired comb on top of stacking gel. 11. When polymerized, load the sample mixture into the wells of the gel. 12. Add the running buffer and run the gel at 60 mA for 1 h. 13. Turn off the electrical current, transfer gel to a 3 MM paper, and dry the gel (see Note 7). 14. For the autoradiography use a cassette that avoids the contact with light and films (film of 24  30 cm) (see Note 8). 15. Develop the film (see Note 9). The expected results are shown in Fig. 3. 16. To quantify the percentage of displacement of the oligonucleotide by ϕ29 SSB wild type or mutants, scan the gel results and use a software (see Note 16) to apply densitometer to the 50 -labeled 17-mer band, which results from the displacement

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Fig. 3 Helix-destabilizing activity of Phi29 SSB wild type and mutant Y57A. The assay was carried out as described in Subheading 3.3, using M13 ssDNA hybridized to a 50 -labeled 17-mer oligonucleotide (as depicted in the figure) and increasing amounts of SSB wild type or mutant. 90  C: Heat-denatured substrate; c: without SSB. (Reproduced with permission from de la Torre I, Quin˜ones V, Salas M, del Prado A (2019) Tyrosines involved in the activity of Phi29 single-stranded DNA binding protein. PLoS ONE 14(5): e0217248. https:// doi.org/10.1371/journal.pone.0217248)

of the oligonucleotide. To get the percentage of displacement, divide the densitometric value for the displaced oligonucleotide (17 mer, bottom band in Fig. 3) by that for the initial substrate (M13-17 mer + 17 mer, top and bottom band, respectively, in Fig. 3).

4

Notes 1. It can also be prepared as a 10 concentrated stock, and then diluted in water before using. 2. It is important that the temperature drop slowly to get a good hybridization (usually around 3 h and a half).

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3. We usually prepare a mix with all the components, except the SSB and the MgCl2; distribute 21.5 μL of it in the Eppendorf; and add the SSB, and finally the metal to initiate the reaction. 4. The column must have more than 800 μL after the centrifugation; if not the unincorporated radioactive nucleotides would not be correctly removed. 5. The volume of water to resuspend depends on the amount of radioactivity, usually in 30 or 45 μL (more radioactivity, more water, to avoid loading the gel with too high amounts of radioactivity). 6. The bromophenol blue and xylene cyanol cannot be used as markers for DNA migration in this gel, because they lose their color due to NaOH. 7. 3 MM papers can also be used if a gel-drying equipment is not available. 8. For a better detection of the radioactivity, use amplifier screens. 9. The exposure time depends on the amount of radioactivity (usually 12 h). Low amounts require longer times. 10. Ammonium sulfate is required for the interaction between the DNA polymerase and the terminal protein. 11. We usually prepare a mix with all the components, except SSB and MgCl2. Distribute 21.5 μL of the mix in Eppendorf, and add SSB, and finally the metal to initiate the reaction. 12. When several samples contain the same amount of protein but are assayed at different times, we perform the reactions in the same Eppendorf (in this case the final volume would be 75 μL instead of 25 μL). At the indicated time, remove 25 μL of the reaction and stop it with stop solution. 13. Once can use homemade columns or purchase a mini quick spin oligo columns. 14. Add 1.5 times more molecules of M13-DNA than of radiolabeled primer to ensure the hybridization of all primer molecules. 15. The polymerization time is quite short with the amount of APS and TEMED indicated. If necessary you can use a lower concentration for a longer polymerization time. 16. We use the Image Lab Software.

Acknowledgments This chapter is dedicated to the memory of Professor Margarita Salas. This work was supported by grant BFU2014-52656-P (to Margarita Salas) from the Spanish Ministry of Economy and

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Competitiveness and institutional grants from the Fundacio´n Ramo´n Areces and Banco Santander to the Centro de Biologı´a Molecular “Severo Ochoa.” References 1. Salas M (1991) Protein-priming of DNA replication. Annu Rev Biochem 60:39–71 2. Blanco L, Bernad A, La´zaro JM, Martı´n G, Garmendia C, Salas M (1989) Highly efficient DNA synthesis by the phage ϕ29 DNA polymerase. Symmetrical mode of DNA replication. J Biol Chem 264(15):8935–8940 3. Kamtekar S, Berman AJ, Wang J, La´zaro JM, de Vega M, Blanco L, Salas M, Steitz TA (2004) Insights into strand displacement and processivity from the crystal structure of the protein-primed DNA polymerase of bacteriophage phi29. Mol Cell 16(4):609–618 4. Rodrı´guez I, La´zaro JM, Blanco L, Kamtekar S, Berman AJ, Wang J, Steitz TA, Salas M, de Vega M (2005) A specific subdomain in ϕ29 DNA polymerase confers both processivity and strand-displacement capacity. Proc Natl Acad Sci U S A 102(18):6407–6412 5. de Vega M, Salas M (2012) Chapter 11: Bacteriophage ϕ29 DNA polymerase: an outstanding replicase. In: Properties and modifications FaI, recombination and applications, editor. Encyclopedia of DNA Research. Nova Science Publishers, Hauppauge, pp 319–340. ISBN: 978-1-61324-305-32012 6. Martı´n G, La´zaro JM, Me´ndez E, Salas M (1989) Characterization of the phage ϕ29 protein p5 as a single-stranded DNA binding protein. Function in ϕ29 DNA-protein p3 replication. Nucleic Acids Res 17 (10):3663–3672

7. Gutie´rrez C, Martı´n G, Sogo JM, Salas M (1991) Mechanism of stimulation of DNA replication by bacteriophage ϕ29 single-stranded DNA-binding protein p5. J Biol Chem 266 (4):2104–2111 8. Soengas MS, Gutie´rrez C, Salas M (1995) Helix-destabilizing activity of ϕ29 singlestranded DNA binding protein: effect on the elongation rate during strand displacement DNA replication. J Mol Biol 253(4):517–529 9. Gasco´n I, La´zaro JM, Salas M (2000) Differential functional behavior of viral phi29, Nf and GA-1 SSB proteins. Nucleic Acids Res 28 (10):2034–2042 10. de la Torre I, Quinones V, Salas M, del Prado A (2019) Tyrosines involved in the activity of phi29 single-stranded DNA binding protein. PLoS One 14(5):e0217248. https://doi.org/ 10.1371/journal.pone.0217248 11. Bernad A, Blanco L, La´zaro JM, Martı´n G, Salas M (1989) A conserved 3’-5’ exonuclease active site in prokaryotic and eukaryotic DNA polymerases. Cell 59(1):219–228 ˜ alva MA, Salas M (1982) Initiation of 12. Pen phage ϕ29 DNA replication in vitro: formation of a covalent complex between the terminal protein, p3, and 5’-dAMP. Proc Natl Acad Sci U S A 79(18):5522–5526 13. McDonell MW, Simon MN, Studier FW (1977) Analysis of restriction fragments of T7 DNA and determination of molecular weights by electrophoresis in neutral and alkaline gels. J Mol Biol 110(1):119–146

Chapter 23 Structural Characterization of a Single-Stranded DNA-Binding Protein: A Case Study of the ORF6 Protein from Bacteriophage Enc34 Elina Cernooka, Janis Rumnieks, and Andris Kazaks Abstract In the quest to understand how single-stranded DNA-binding proteins function and evolve at a molecular level, determination of their high-resolution three-dimensional structure using methods such as X-ray crystallography is indispensable. Here we present a collection of methods used in crystallographic studies of the single-stranded DNA-binding protein from the bacteriophage Enc34, from designing expression constructs through to protein production, purification, and crystallization, to determination and analysis of the protein’s three-dimensional structure. The chapter aims to shed light on all the essential stages in a structural study of a single-stranded DNA-binding protein, with a spotlight on procedures specific to X-ray crystallography to aid those interested in venturing into structural biology. Key words Single-stranded DNA-binding protein, Structural biology, X-ray crystallography, Construct design, Protein production, Protein purification, Crystallization, Co-crystallization, Structure determination, Structural analysis

1

Introduction The function of any protein, be it an enzyme, an antibody, or a single-stranded DNA-binding protein (SSB), emerges as the most direct consequence of the highly complex and unique shape that its constituent string of amino acid folds into - its three-dimensional structure. Determination of the exact pattern of how the atoms are spatially arranged within a protein molecule thus provides the most fundamental insight into its characteristics and functioning. The three-dimensional structure can immediately suggest the molecular mechanism of how the protein functions, uncover distant connections to other proteins and thus reveal its evolutionary past, and enable further studies such as structure-guided site-directed mutagenesis, or investigation of different functional (i.e., free and ligand bound) states of the protein for an even deeper understanding of how it works.

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3_23, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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The most prevalent technique for determining high-resolution structures of biological macromolecules is X-ray crystallography in which the structure is elucidated from the diffraction pattern that X-rays produce as they pass through an ordered arrangement of protein molecules within a crystal. As of now, the Protein Data Bank, the global public repository of determined macromolecular structures, holds dozens of SSB structures, some of clear homology and some unique, originating from all forms of life, from viruses to humans. Among them are about 20 structures of SSBs bound to single-stranded DNA, which capture the very interactions that take place when an SSB fulfills its role in protecting DNA intermediates and maintaining genomic integrity. The long road to a crystal structure of an SSB begins with the design of an expression construct which involves standard molecular cloning techniques widely used and familiar to many, but with several aspects that need to be considered to design a suitable construct for use in structural biology. After the expression constructs have been made, the project proceeds to production and purification of the SSB of interest, which may turn out to be a quite straightforward or a rather challenging endeavor, as crystallization trials require relatively large amounts of highly purified protein. With a pure and concentrated SSB in hand, the project then enters possibly its most critical stage en route to the structure—crystallization. The conditions upon which—if at all—the protein is willing to crystallize are impossible to predict in advance; therefore hundreds of different conditions usually need to be tested until, hopefully, a suitable one is found. The crystals are then brought to a synchrotron X-ray radiation facility for testing, and in case of welldiffracting crystals (which, as might be suspected, is not always the case) a set of diffraction images are collected from which the protein’s three-dimensional structure can then be solved, i.e., an interpretable electron density map can be calculated which allows for building and analyzing a three-dimensional atomic model of the protein using specialized software tools. In this chapter we walk through all the steps that were taken to structurally characterize the SSB of the bacteriophage Enc34 [1], encoded by open reading frame (ORF) 6 in its genome. We describe how to develop an expression construct and a production and purification pipeline for the full-length ORF6 protein to test its crystallization ability. The protein readily crystallized under several conditions, so we proceeded to produce a selenomethioninesubstituted variant of the full-length ORF6 (SeMet-ORF6) which is required to provide experimental phasing data for structure solution. After optimization of purification and crystallization protocols for SeMet-ORF6, the obtained crystals were used for X-ray data collection and structure determination. Finally, a C-terminally truncated variant of ORF6 (ORF6ΔC) was constructed with the rationale that the unstructured tail might both impede

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crystallization due to its flexibility and compete for the same binding surface as the single-stranded DNA [2], and a structure of the ORF6ΔC in complex with DNA was obtained. We provide details on all major procedures involved in the study, from molecular cloning strategies to obtaining protein crystals to in silico analyses of the SSB’s three-dimensional structure.

2 2.1

Materials Construct Design

1. Reagents and equipment electrophoresis.

for

PCR

and

agarose

gel

2. Purification kit for PCR products. 3. Enzymes for molecular cloning (restriction endonucleases, DNA ligase). 4. E. coli competent cells for routine cloning (DH5α, TOP10, XL1-Blue, or a similar strain). 5. Liquid LB medium and LB agar supplemented with appropriate antibiotic. 6. Plasmid DNA minipreparation kit. 2.2 Protein Production

1. E. coli expression strains BL21(DE3) and B834(DE3). 2. Liquid LB medium and LB agar supplemented with appropriate antibiotic. 3. Liquid 2
TY medium supplemented with appropriate antibiotic. 4. Selenomethionine medium kit, e.g., Molecular Dimensions. 5. Sterile test tubes and 2 L flat-bottom flasks. 6. Cell incubation shaker with temperature control. 7. Spectrophotometer for measuring optical density of the cultures. 8. 100 mM Solution of isopropyl β-D-1-thiogalactopyranoside (IPTG) in ultrapure water. Store at 20  C. 9. Centrifuge for cell harvesting.

2.3 Protein Purification

1. TN buffer: 20 mM Tris–HCl pH 8.0, 300 mM NaCl. Store at 4  C. 2. Ultrasonic cell disruptor. 3. Refrigerated benchtop centrifuge capable of achieving 15,000
g and suitable tubes for centrifugation. 4. 5 mL Syringes.

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5. IMAC purification column, e.g., 1 mL HisTrap FF crude. 6. Elution buffer A: 20 mM Tris–HCl pH 8.0, 300 mM NaCl, 20 mM imidazole. Store at 4  C. 7. Elution buffer B: 20 mM Tris–HCl pH 8.0, 300 mM NaCl, 300 mM imidazole. Store at 4  C. 8. 1 M Solution of dithiothreitol (DTT) in ultrapure water: Store at 20  C. 9. Recombinant TEV protease (see Note 1): Store at 20  C. 10. High-performance liquid chromatography system. 11. Desalting and buffer-exchange column(s), e.g., 5 mL HiTrap Desalting. 12. Strong anion-exchange column, e.g., 1 mL Mono Q 5/50 GL. 13. Mono Q buffer A: 20 mM Tris–HCl pH 8.0, 100 mM NaCl. Store at 4  C. 14. Mono Q buffer B: 20 mM Tris–HCl pH 8.0, 1 M NaCl. Store at 4  C. 15. Standard SDS-PAGE reagents and equipment. 16. Spectrophotometer for measuring protein concentration. 17. Centrifugal ultrafiltration devices with appropriate molecular weight cutoff, e.g., Amicon Ultra. 2.4

Crystallization

1. Small-volume pipetting robot or 0.5–10 μL and 10–100 μL multichannel pipettes. 2. Swissci 96-well two-drop MRC crystallization plates. 3. Crystallization screens in 96-well deep-well blocks (see Note 2). 4. Clear sealing tape or adhesive sheets, e.g., Molecular Dimensions. 5. Light microscope. 6. Microloops of various sizes, a CryoWand, magnetic CryoCaps and vials, CryoCanes, and CryoSleeves. 7. Liquid nitrogen and suitable storage containers/dewars.

2.5 Software for Structure Determination and Analysis

1. CCP4 software suite (http://www.ccp4.ac.uk). 2. MolProbity (http://molprobity.biochem.duke.edu/). 3. Dali server (http://ekhidna2.biocenter.helsinki.fi/dali/). 4. PDBePISA (https://www.ebi.ac.uk/pdbe/pisa/). 5. PyMOL (https://pymol.org/2/).

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Methods Structural studies of an SSB or indeed any previously uncharacterized protein require successful accomplishment of a number of sequential stages: designing an expression construct for the protein, production and purification of the recombinant SSB, crystallization, determination of the three-dimensional structure of the protein using X-ray crystallography, and analysis of the structure with various software tools. The experimental steps and/or considerations that constitute each of them are detailed below, with a generalized outline of each step and more detailed descriptions of the procedures that were used in studies of the Enc34 SSB where appropriate.

3.1

Construct Design

The first matter to consider when designing your construct is the choice of a suitable expression vector which will determine the production level of the protein. Structural studies typically require large amounts of the protein; hence vectors that provide high expression levels are preferred. For expression in Escherichia coli, the T7 expression system [3] is among the most widely used and usually results in very robust expression levels and is the recommended option to produce the recombinant SSB for structural studies. A wide selection of T7-based vectors is available commercially such as the pET system developed by Novagen. The next effort in the construct design should be directed toward enabling easy means of purification for the recombinant protein. Crystallization requires a highly pure (>95%) protein preparation; hence it is usually essential to append an affinity tag for purification to the protein. A hexahistidine (6
His) tag is a widely used and suitable option, but other possibilities also exist (see Note 3). The affinity tag can be either introduced via in-frame cloning into a suitable vector or, alternatively, included in the primer sequence used for amplification. The third important matter is to assess whether there are any flexible or unstructured parts in the protein, as they will often interfere with crystallization; even if they do not, unstructured regions are usually not resolved in the structure; hence their removal from the protein should be an option to consider (see Note 4). While crystallization of the full-length protein should still be tested, in case a long unstructured stretch at the N- or C-terminus is identified, a respective truncation of the protein might be necessary. The purification tag, although usually rather short, might also interfere with crystallization, and its removal after affinity purification is generally advisable; this can be enabled by engineering a sequence for a site-specific protease between the tag and the rest of the protein. When the desired expression construct has been envisioned, the procedure then follows routine molecular

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Fig. 1 Cloning strategy for the Enc34 SSB. Fw denotes the forward primer, Rv1 the reverse primer for the fulllength SSB, and Rv2 the reverse primer for the truncated protein; start and stop codons are colored in gray, nucleotide sequences encoding the 6
His tag in red, the TEV protease site in yellow, and the Enc34 SSB in green, cyan, and blue; the amino acid sequence is given below the nucleotide sequence. A secondary structure prediction from Jpred is indicated below the Enc34 SSB sequence

cloning steps involving design of appropriate primers, PCR amplification, digestion of the cloning vector and insert by restriction endonucleases, ligation, transformation, and screening of colonies. For cloning of the Enc34 SSB gene, we chose the pETDuet-1 vector which contains a sequence encoding the 6
His tag upstream the first multiple cloning site (Fig. 1). To enable removal of the tag after affinity purification, a sequence corresponding to a tobacco etch virus (TEV) protease cleavage site was added to the forward primer, preceded by a BamHI site for cloning in-frame with the 6
His tag in the vector, and followed by a StuI restriction site to facilitate screening of clones with inserts (see Note 5).

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Secondary structure prediction revealed a stretch of unstructured residues at the C-terminus of the protein and it was decided that two constructs, one of the full-length protein and the other with a 21-residue truncation from the C-terminus, shall be made. Two different reverse primers were designed containing a BspTI site for cloning and used in a PCR together with the forward primer to amplify the respective fragments from the phage genomic DNA. Subsequent to PCR amplification, the expression constructs were created using standard molecular cloning techniques [4]. 3.2 Protein Production

When using a T7-driven system in E. coli, the standard protocol is to grow the cells in rich medium until OD600 is 0.6–0.8, induce expression with 1 mM IPTG, and cultivate the bacteria for 4 h at 37  C before harvesting them by centrifugation. These conditions usually result in very high expression levels, but a not too uncommon issue is a complete or partial insolubility of the recombinant protein, which unfortunately was the observation also for the Enc34 SSB. However, during our efforts to produce a wide variety of different recombinant proteins over the years, we have developed an alternative protocol in our laboratory which uses lower cultivation temperature and inducer concentration and has a high success rate in producing soluble protein while maintaining comparable or better protein yields than the standard protocol (Fig. 2). Both the full-length and C-terminally truncated Enc34 SSB were produced according to the following protocol:

Fig. 2 SDS-PAGE analysis of production and solubility of the Enc34 SSB using standard or optimized expression protocols. Lane 1, molecular weight marker; lane 2, lysate of uninduced cells; lanes 3 and 6, total cellular protein at the end of protein production; lanes 4 and 7, insoluble protein fraction; lanes 5 and 8, soluble protein fraction. The arrow points to the Enc34 SSB

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1. Transform E. coli BL21(DE3) competent cells with the protein-encoding plasmid, plate on selective LB agar plates, and incubate overnight at 37  C. 2. On the following day, pick eight colonies and inoculate each in 5 mL of LB medium supplemented with the appropriate antibiotic. Incubate the cultures at 37  C overnight without agitation. 3. In a 2 L flat-bottom flask, inoculate 400 mL of antibioticsupplemented 2
TY medium with the overnight cultures. Cultivate at 25  C with agitation at 200 rpm until OD600 of the culture reaches 0.6–0.8. 4. Reduce temperature to 22  C and add IPTG to a final concentration of 0.01 mM. Continue cultivation overnight (16–18 h). 5. Harvest cells by centrifugation for 20 min at 6000
g. The cells can then be stored at 20  C or used immediately to proceed with purification. If the SSB of interest shares less than 35% sequence identity with a protein of known three-dimensional structure, solution of its structure using X-ray crystallography will likely require experimental phasing. This is most often done through single- or multiplewavelength anomalous diffraction (SAD or MAD, respectively) experiments which require collecting crystal diffraction data at wavelength(s) close to an X-ray absorption edge of “heavy” (i.e., with a number of electrons significantly higher than the “light” carbon, oxygen, nitrogen, and hydrogen) atoms incorporated in the protein. The approach almost exclusively used for achieving this is substitution of methionine residues in the protein with selenomethionine, a methionine analog in which the sulfur atom has been exchanged for selenium. In practice this entails production of the protein of interest by a methionine-auxotroph strain in a medium containing selenomethionine; with the T7 expression system, the E. coli strain B834(DE3) is routinely used and the expression is done in M9-based minimal medium supplemented with selenomethionine (see Note 6). The Enc34 SSB did not have any homologs with a known three-dimensional structure at the time, which required production of a selenomethionine-substituted protein to solve the structure. To achieve this, the full-length Enc34 SSB was produced in E. coli B834(DE3) cells using a commercially available selenomethionine kit (medium base with glucose, nutrient mixture, and selenomethionine solution). Expression was done according to the instructions provided by the manufacturer with the only modifications that 600 mL of medium was used, and that the cultivation was done at 25  C overnight (see Note 7).

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The purification begins with lysing the bacteria and centrifugation of the crude lysate to remove cellular debris, and proceeds with several column chromatography steps to progressively separate the SSB of interest from the plethora of other proteins in the lysate to eventually arrive at a homogeneous and concentrated SSB preparation ready for crystallization. As the first step, the clarified lysate is usually passed through an affinity purification column which quickly removes more than 90% of the contaminating proteins. After optional removal of the purification tag, the protein preparation is then routinely applied to a high-resolution ion-exchange column that separates out most of the remaining contaminants. As the final purification step, the protein is sometimes loaded onto a “polishing” gel filtration column to further remove minor contaminants and possible aggregates of the protein of interest, and, if necessary, to exchange the buffer to something suitable for crystallization (see Note 8). Purification of the Enc34 SSB was done according to the following protocol: 1. Resuspend 0.5 g cells in 5 mL of ice-cold TN buffer (see Note 9) and place on ice. Disrupt the cells by sonication, taking care not to heat up the suspension. Clarify the lysate by centrifugation at 15,000
g for 30 min at 4  C in a refrigerated centrifuge and collect the supernatant. 2. Pre-equilibrate a 1 mL HisTrap FF crude column with TN buffer. Using a syringe, pass the clarified lysate through the column and collect the flow-through. Wash the column with 5 mL of TN buffer and collect the eluate together with the previous fraction. Elute weakly bound proteins with 3 mL of elution buffer A, followed by 3 mL of elution buffer B to elute the SSB. Collect each fraction in a separate tube. 3. Supplement elution fraction B with 1 mM DTT, add 200–250 μg of recombinant TEV protease, and incubate at 4  C overnight. 4. Remove imidazole using a 5 mL HiTrap Desalting column pre-equilibrated with TN buffer (see Note 10). Pass the desalted preparation through a 1 mL HisTrap column and collect the flow-through that contains the cleaved SSB. Wash the column with additional 3 mL of TN buffer and add the eluate to the previous flow-through fraction (see Note 11). 5. Reduce NaCl concentration in the flow-through fraction to 100 mM by dilution with 20 mM Tris–HCl pH 8.0, and apply the preparation onto a 1 mL MonoQ 5/50 GL column pre-equilibrated with MonoQ buffer A. Elute bound proteins with 20 column volumes of a linear gradient from 0% to 100% MonoQ buffer B. Visualize the stages of protein purification by SDS-PAGE (Fig. 3).

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Fig. 3 Purification of the Enc34 SSB visualized by SDS-PAGE. Lane 1, molecular weight marker; lane 2, clarified bacterial lysate; lane 3, HisTrap flow-through; lane 4, HisTrap elution fraction A; lane 5, HisTrap elution fraction B; lane 6, HisTrap elution fraction B after overnight digestion with TEV protease; lane 7, HisTrap flow-through after digestion with TEV protease; lane 8, HisTrap-bound protein fraction after digestion with TEV protease; lanes 9–12, MonoQ peak fractions; the arrow points to the Enc34 SSB

6. Pool the MonoQ peak fractions and estimate the concentration of the purified protein (see Note 12). Reduce NaCl concentration to ~100 mM and concentrate to 10 mg/mL using a centrifugal ultrafiltration device. 3.4

Crystallization

Crystals diffracting to high resolution are the key for a successful protein crystallography project, but crystallization is unfortunately the stage of the least control and the most trial-and-error within the whole endeavor. Crystallization is achieved by slowly concentrating a protein solution until there is too little solvent left to fully hydrate the protein, which at this point starts to make contacts with other protein molecules. This phenomenon usually manifests itself as the protein precipitating out of solution; however, in some cases the molecules can instead pack into an ordered three-dimensional lattice—a crystal. Crystallization is dependent on the presence of different polymer precipitants, salts, buffers, and other components of the crystallization mixture which are impossible to predict in advance and must be determined experimentally in crystallization trials. In practice, certain combinations of precipitants, salts, and other molecules have proven to be more successful in promoting crystallization than others, and these are compiled and provided commercially as high-throughput crystallization screens by several companies. After setting up trials with several such screens, the crystallization plates are incubated at a constant temperature and periodically inspected under a microscope for crystal formation

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under any of the conditions. The time required for crystals to grow may range from hours to days to sometimes weeks and months. When—to much delight—a crystal is discovered, an attempt to further optimize the conditions usually takes place by systematically changing the concentrations of the constituent components of the mixture to grow more and better crystals. Finally, the crystals are collected and prepared for shipping to a synchrotron radiation facility for X-ray diffraction experiments. Separate crystallization trials were performed for the full-length ORF6 protein and the deletion variant ORF6ΔC, both in the absence and presence of oligonucleotides of various lengths. All samples were initially screened with several commercially available 96-well crystallization screens and the discovered conditions were then manually replicated and/or optimized to produce more crystals for data collection (Fig. 4). Crystallization trials are most conveniently set up using a pipetting robot that can handle small sample volumes (1 μL or smaller) but can also be done using appropriate multichannel pipettes. Make sure to have at least 40 μL of concentrated (~10 mg/mL) SSB solution per screen if using a small-volume robot, or at least 100 μL per screen otherwise. As much as possible, set up several screens at once to increase chances of success. The process of co-crystallizing the SSB with ssDNA is technically the same as for the protein on its own, with the only difference that the DNA and protein are mixed together immediately before crystallization (see Note 13).

Fig. 4 Crystals used for structure determination of the Enc34 SSB. (a) A crystal of the full-length protein, (b) crystals of the C-terminally truncated ORF6ΔC protein, (c) crystals of the ORF6ΔC-oligothymidine complex

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If using a pipetting robot, make sure that the robot can handle a 2 mL 96-well deep-well block and a 96-well two-drop MRC crystallization plate; if necessary, consult the manual of your pipetting robot to configure it accordingly. Then create a workflow to execute the following protocol: 1. Place a deep-well block with the crystallization screen, an empty crystallization plate, and a microtube with a concentrated (~10 mg/mL) protein solution at appropriate positions in the pipetting station. 2. Transfer 50 μL of each crystallization solution from the deepwell block to the corresponding bottom reservoir of the MRC plate. 3. Transfer 0.4 μL of the concentrated protein solution to one of the small top wells in each compartment of the MRC plate. 4. Aspirate 0.4 μL of each bottom reservoir solution and add it to the already placed protein drop in the adjacent top well (see Note 14). 5. When all wells have been processed, remove the MRC plate from the pipetting station and immediately cover with a transparent adhesive sealing sheet or tape (see Note 15). Remove the deep-well block with the crystallization solutions from the pipetting station, cover with an adhesive foil to prevent evaporation, and store at 4  C until next use. If using multichannel pipettes: 1. With a 10–100 μL multichannel pipette, transfer 50 μL of each solution from a crystallization screen in a deep-well block to the corresponding bottom reservoir of the MRC plate. 2. Aliquot at least 13 μL (for use with an 8-channel 0.5–10 μL pipette) or 9 μL (for use with a 12-channel pipette) of the concentrated protein solution into each microtube of an 8- or 12-microtube strip. Using a 0.5–10 μL multichannel pipette, column by column or row by row transfer 1 μL of the protein solution to one of the small top wells in each compartment of the MRC plate. 3. Column by column or row by row, aspirate 1 μL of each bottom reservoir solution and add it to the already placed protein drop in the adjacent top well. Do not repeatedly aspirate to mix as this will likely introduce bubbles. 4. Cover the crystallization plate with a transparent adhesive sealing sheet or tape (see Note 15). Cover the deep-well block containing the crystallization screen with an adhesive foil to prevent evaporation and store at 4  C until next use.

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Immediately after the crystallization plate has been set up, examine the crystallization plate under microscope and note wells with clear drops, precipitation, or crystalline formations. Incubate the plate in a vibration-free environment at a constant temperature (see Note 16). Examine the plate daily for the next couple of days and then once a week or until suitable crystals have grown (see Note 17). In case of small or thin crystals, optimize the conditions by varying the amounts of the constituent components of the mixture in a systematic manner (see Note 18). When crystals of satisfactory quality are obtained, using a microloop mounted on a CryoCap, pick a crystal from the drop, quickly sweep through a cryoprotectant solution, and place in a cryovial submerged in liquid nitrogen (see Note 19). The crystal can then be stored in liquid nitrogen for long periods of time (at least for several months) until shipment to a synchrotron facility in a dry shipper for data collection. Samples can be stored and transported either in CryoCanes and CryoSleeves or directly in baskets compatible with the system used at the planned data collection site. 3.5

Data Collection

The experimental part of the structure determination project is concluded by collection of X-ray diffraction data from the crystals. Data collection is carried out at specialized beamlines at synchrotron light sources, large national- or international-scale research facilities purpose-built for providing high-intensity X-rays for various research applications (see Note 20). Beamline scientists and other highly qualified staff will provide the necessary training, technical assistance, and professional advice of how to collect data at each particular station, but it is still at the users’ discretion to know their samples and what they want to do with them to decide on the best strategy for data collection. When the crystals are brought to the synchrotron, the initial business usually is to screen them for the best diffraction, i.e., collect two diffraction images 90 apart along the rotation axis and visually examine them to estimate the maximum resolution and detect crystal defects such as spot smearing or multiple lattices. The diffraction pattern will also tell whether crystals grown at different conditions are actually different crystal forms and if so which of these is the most perspective, and, if several cryoprotecting substances had been tested when freezing the crystals, help to identity the most suitable one. The best crystals are then subjected to the collection of a complete dataset for structure determination (see Note 21). The optimal data collection strategy will differ depending on the purpose for which the data are being collected [5]. Experimental phasing using selenomethionine-labeled protein crystals relies on measuring very small (usually 3–5%) differences in intensity between sets of reflections called Friedel pairs; hence high accuracy of the measured intensities is of topmost importance. Such

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level of accuracy is best achieved by increasing redundancy, i.e., by measuring any reflection several times and then calculating the average value which is bound to be more accurate than the noisier measurement of any individual reflection. In practice this means collecting more data images at a lower X-ray dose (i.e., exposure time) to limit radiation damage. To collect the weak signal in the first place, an essential parameter for a SAD data collection experiment is a precise setting of the X-ray wavelength to one where the anomalous signal actually exists and is preferably the strongest possible. An obvious requirement is a beamline with a tunable X-ray wavelength which must of course be checked in advance before requesting beamtime at a particular experimental station. At the beamline, when a suitable crystal has been mounted and awaits ready for data collection, an X-ray fluorescence scan is first performed around the theoretical selenium absorption edge to experimentally determine the wavelength where anomalous scattering is greatest (“the peak”). The peak wavelength is then used for data collection. The minimum amount of data that needs to be collected depends on the symmetry of the crystal and its orientation with respect to the beam, but it is always recommended, as much as radiation damage and available time allow, to collect the maximum amount of data, i.e., to do a full 360 scan or even more for low-symmetry triclinic or monoclinic space groups. However, since usually it cannot be known in advance what radiation dose the crystal will be able to sustain, it is always useful to follow recommendations provided by diffraction image processing software (e.g., the “Strategy” tool in iMosflm) for choosing the best rotation angle for starting data collection to maximize the amount of useful data that are collected from the crystal. If the beamline supports it, a helical scan data collection protocol is highly recommended in which the crystal is continuously shifted during data collection to constantly expose a fresh part to the beam, significantly reducing radiation damage and improving data quality. The method is particularly useful for long and thin needle-shaped crystals but will improve data quality for any crystal that is considerably larger than the beam size. Diffraction at both ends of the helical path should however be tested before starting data collection, especially if the crystal is of irregular shape and does not appear monolithic, as it might occur that the diffraction is not uniformly good at all positions within the crystal. If the crystal still diffracts strongly after a full dataset at the peak wavelength has been collected, data collection at other (inflection point, high-energy remote and low-energy remote) wavelengths from the same crystal can be pursued to provide better data for MAD phasing. If, on the other hand, the crystal suffers from excessive radiation damage, which is evident by rapid weakening of spot intensities as the collection of diffraction images progresses,

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it might be necessary to collect more data from a different spot in the crystal (if the crystal is large enough and helical scan is not available), or collect data from more crystals and then scale them together during data processing to extract enough signal for solving the structure. It is also recommended to do “live” data processing while collecting data to get an early estimate of the anomalous signal present in the data that will help to make a decision whether more data needs to be collected. If the structure is intended to be solved using molecular replacement, high accuracy of the measured intensities is of secondary importance while completeness of data, particularly at low resolution, is more significant. The optimal data collection strategy can be determined using “strategy” options provided in software such as iMosflm; however, as the correct space group of the crystal is usually unknown at this stage, it is recommended that 180 worth of diffraction images are collected from the starting position recommended by the software, which will ensure a complete dataset even if the space group later turns out to be of lower symmetry than the automatically selected one. Also, preliminary processing of collected data on-site is recommended to avoid unpleasant surprises when processing the data later at home. Solving of the structure both by experimental phasing and by molecular replacement does not require a particularly high resolution to be effective, but refinement and model building greatly benefit from having the highest resolution possible. Collection of a high-resolution dataset will usually be done using a “native,” i.e., non-selenomethionine-labeled, protein crystal, although there is no reason to expect that a native crystal will necessarily diffract better than a labeled one. In case of molecular replacement, oftentimes only a single high-resolution dataset is collected that is used both for solving the structure and for subsequent high-resolution refinement. Nevertheless, a strategy to collect high-resolution data will use longer exposure times to better measure weak reflections, at some expense of the number of diffraction images that can be collected before the crystal starts suffering from radiation damage. When aiming for high-resolution data, it is also recommended to set the detector distance to a maximum resolution of at least 0.3 Å higher than that of the weakest reflection still visible on the image by eye to avoid missing useful data. It is still necessary to collect a complete dataset, and higher redundancy certainly does not harm as it provides more accurate data. 3.6 Structure Determination

The structure determination stage of the project makes use of many different software programs to index and scale the measured reflections, do either experimental phasing or molecular replacement, and build and refine the atomic model of the protein. Usually at least a couple of equally good alternatives exist for each of these tasks, and indeed the structure can be solved via many different

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paths, sometimes coming down more to personal preferences of the crystallographer than to other factors. However, the most comprehensive software package for X-ray crystallography is the CCP4 suite (http://www.ccp4.ac.uk) which at its core is a collection of well over a hundred individual programs each dedicated for a specific task but all using unified input/output parameter and file formats to easily pass data along to each other. The most convenient method of using the CCP4 programs for most users will be the CCP4i graphical user interface which comes included with the suite. 3.6.1 Data Reduction

The first step in solving the structure from the collected X-ray data is finding, indexing, and integrating the diffraction spots in the collected images; determining the space group of the crystal; and scaling and merging of the integrated reflections, collectively called data reduction. For spot finding, autoindexing, and integration the CCP4 suite includes the program MOSFLM together with a graphical user interface iMosflm. A standard workflow for processing data with iMosflm involves the following steps: 1. Set up a new CCP4 project. From the top-left drop-down menu, select “Data Reduction and Analysis” and from the “Data Processing using Mosflm” subcategory select “Start iMosflm.” 2. In the iMosflm window that opens, click “Add images” in the toolbar and select an X-ray diffraction image from your dataset. 3. In the image viewer window that appears, use the Masking tool to draw a mask around the beam stop to exclude this region from subsequent processing. 4. Switch to the Indexing tab in the left panel of the main window. Two diffraction images are automatically picked and used for spot finding and autoindexing. In the table with possible lattices, a solution with the highest symmetry and reasonably good fit to data is automatically selected. The predicted spot locations of that solution are shown in the image viewer window (see Note 22). 5. If the resolution is higher than 3.5 Å, go to the Cell Refinement tab in the left panel. Press the Process button to refine cell parameters of the selected solution. 6. Select the Integration tab and click “Process” to integrate the data. After data integration is finished, the correct space group of the crystal must be determined which is most easily done with the CCP4 program POINTLESS. To do this: 1. From the “Data Reduction and Analysis” category open the “Import Integrated Data” subcategory in the CCP4i and select the “Find or Match Laue Group” task.

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2. In the window that opens, select the output .mtz file from iMosflm as the MTZ #1 and select Run ! Run Now to execute the task (see Note 23). Finally, the integrated reflections must be scaled, and equivalent reflections merged together. This can be done using the program SCALA from the CCP4 suite: 1. Open the “Data Reduction and Analysis” category and the “Import Integrated Data” subcategory in the CCP4i and select the “Scale and Merge Intensities” task. 2. In the window that opens, select the output file from POINT LESS as the input MTZ file. It is also convenient to select the “Ensure unique data & add FreeR column” option at this stage. 3. Run the task by selecting Run ! Run Now. After the job has finished, double-click the job to examine the results. If necessary, rerun the SCALA job with an appropriate resolution cutoff (see Note 24). 3.6.2 Phasing

The distribution of electron density in the crystal—i.e., the much sought-after structure—and the diffraction pattern recorded in the collected images are interrelated by a mathematical operation called a Fourier transform. If the indices, intensities, and phases for each of the reflections are known, calculation of the electron density map is straightforward. Unfortunately, phase information is not present in the diffraction images, and the structure cannot be calculated without it. Hence, the “phase problem” has to be solved by other means, such as by employing anomalous diffraction from SeMetlabeled crystals, or by using an already determined similar structure via molecular replacement (see Note 25). In case of the Enc34 SSB, the structure was initially solved by SAD using SeMet-ORF6 crystals, and the subsequent ORF6ΔC and ORF6ΔC-ssDNA complex structures were determined using molecular replacement. Experimental phasing techniques will almost certainly include a density modification step which requires knowledge of what is the proportion of the solvent in the crystal. If the cell parameters and the molecular weight of the protein are known, the most probable number of molecules in the crystallographic asymmetric unit and the solvent content of the crystal can be estimated as follows: 1. Prepare a fasta file with the sequence of your protein. 2. Open the “Density Improvement” category in the CCP4i and select the “Cell Content Analysis” task. 3. Select the output file from SCALA as the input MTZ file, choose to use molecular weight estimated from sequence file, select your fasta file, and then choose the task to Run Now.

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4. In the Solvent content analysis panel, note the number of molecules in the asymmetric unit (Nmol/asym) and solvent content (%solvent) with the highest probability (P(tot)). For experimental phasing it is convenient to use the SHELX C/D/E pipeline provided with the CCP4 suite. 1. Select the “Experimental Phasing” category and then the “Automated Search & Phasing” subcategory in the CCP4i and pick the “SHELX C/D/E pipeline” task. 2. In the window that opens, select to Run SHELX to prepare HA data from a SAD experiment and input the previously calculated solvent fraction (not percentage) in the respective box. Unselect the main chain autotracing option for SHELXE. In the “Input files” subcategory, select the output file from SCALA as the MTZ input file, specify that input is in the form of structure factors, and make sure that the HA F(+) and SigF(+), and HA F() and SigF(), columns have been correctly selected. Run the task by selecting Run ! Run Now. 3. When the run has finished, two sets of output files will be produced corresponding to two enantiomeric solutions, the “original” and “inverted.” Select to view any of the two .pdb output files from the job. The file will list the coordinates of the located selenium atoms, followed by their estimated occupancies. A sharp drop in occupancy at a point corresponding to the expected number of methionine residues in the asymmetric unit strongly suggests that the solution is correct. Inspect the SHELX log file which at the very bottom presents a table of map correlation coefficients versus resolution for the original and inverted solutions, one of which should be noticeably higher than the other. Finally, open both output MTZ files in COOT. The correct solution should reveal recognizable structural features of a protein such as α-helices and β-strands (Fig. 5). Molecular replacement (MR) requires a search model of a similar already determined structure. In some cases, the structure to be solved will be of a protein with an already determined structure, for example, a different, better diffracting crystal form of the same protein. In such cases, the model of the already determined structure can be used in MR without any modifications. Alternatively, the unsolved structure might be of the same protein with some minor modifications such as the C-terminally truncated ORF6 protein ORF6ΔC, or an assembly of the known protein with another component of unknown structure such as the ORF6ΔC-ssDNA complex. In this case, it is recommended that the input model is accordingly modified, which in case of ORF6ΔC meant removal of the deleted residues from the prior structure (this

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Fig. 5 Electron density maps of the Enc34 ORF6 protein. Top, the first experimental SeMet-SAD map before auto-building; bottom, a 2Fo-Fc map after several cycles of model building and refinement

can be done either by manually editing the PDB file in a text editor or visually in software such as PyMol or COOT). In many cases, however, the protein with the unsolved structure will have some level of sequence identity with a different protein of a known threedimensional structure. The success of MR relies on the similarity of the known structure to the unknown one, which is, by definition, unknown and hence is but an educated guess. Particularly when the sequence similarity is quite low, several different models often have to be tested until a solution is found. As a general recommendation, the first model to try would be the most similar structural homolog with, guided by sequence alignment, extra parts such as flexible loops removed. If several structures of similar sequence are

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available, these can be tried as further search models in an order of decreasing sequence identity. If these options fail, another possibility is to truncate amino acid side chains in the search models to the Cβ atom, i.e., make a poly(alanine) model. The CCP4 suite includes a program CHAINSAW for this purpose. Yet another option is to use a homology model in which, guided by sequence alignment, either only the matching residues are kept while others are pruned or a complete model is generated using more advanced comparative modeling techniques. The first option is implemented in CHAINSAW while software such as Rosetta (http://robetta. bakerlab.org) can be used for advanced structure prediction. While we have not seen much success with homology models in MR, these are certainly an option if other efforts are unsuccessful. When, one way or another, the search model has been prepared, a powerful and often-used program for carrying out the MR procedure within the CCP4 suite is PHASER. 1. Open the “Molecular Replacement” category and the “Model Generation” subcategory in the CCP4i and select the “Phaser MR” task. 2. In the “Define data” category, select the merged intensities from SCALA as the input MTZ file. In the “Define ensembles (models)” category, choose your prepared model file as PDB #1 and input the requested sequence identity value in the corresponding box. In the “Define composition of the asymmetric unit” category, specify a fasta file with the sequence of the unsolved structure (not the input model) and adjust the number of molecules in the asymmetric unit if necessary (this can be determined using the “Cell Content Analysis” task as outlined above). Finally, in “Search parameters,” select ensemble1 as the search model and specify the expected number of copies to search for. Select Run ! Run Now to start the job. 3. After completion, double-click the job to view the results. In the Results tab, under “MR Result,” inspect the translation function Z-score (TFZ) of the top solution(s). Generally, a TFZ of over six suggests a possible correct solution and over eight a definite solution. At the bottom of the result page, click the COOT button to visually inspect the placed model and electron density. A correct solution should reveal features in the electron density that were not present in the input model such as different side chains and missing ligands, and these should correlate with positive (green) density in the Fo-Fc difference map. Before model building, you can further refine the solution in REFMAC (see in the next section) to get a better map.

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When the phasing efforts have resulted in an interpretable electron density map, the structure is fundamentally “solved,” and ahead lies just the final task to actually interpret the map and build a realistic atomic model of the protein. The initial model can be built automatically by several auto-building tools which can save a lot of manual labor, but as the process advances, increasingly more effort has to be devoted to manually inspect, correct, and complete the model. The model then undergoes refinement in software programs that optimize the modeled atomic coordinates and other parameters to produce a model that is both in good agreement with experimental data and follows the known geometry of bond lengths, angles, and other features of macromolecular structure. The refined structure is then subjected to validation where it is evaluated by a number of criteria such as bond and angle geometry, side-chain conformations, or steric clashes between atoms, to assess how realistic the model is and pinpoint areas which need further improvement. After repeated cycles of model building, refinement, and validation, the final model can then be proudly presented to the rest of the scientific community by depositing it into the Protein Data Bank, the worldwide public repository of macromolecular structures. If the structure was solved by experimental phasing, almost certainly the model will have to be built from scratch. If one wishes so, this can be done manually in the modeling software COOT (see Note 26) by first fitting dummy secondary structure elements (α-helices and β-strands) into the map (Calculate ! Other Modeling Tools ! Place Helix Here and Place Strand Here), then recognizing the protein sequence and modeling side chains (Simple Mutate in toolbar, or Calculate ! Mutate Residue Range) with their correct residue numbers (Edit ! Renumber Residues), and finally joining the separate elements together in a single model (Edit ! Merge Molecules, Change Chain IDs) and modeling the loops between them (Add Residue in toolbar, Calculate ! Fit Loop). Much more often, however, this task is delegated to a piece of software which traces the chain and builds an initial model automatically given the protein’s sequence. Within the CCP4 suite, one of the programs able to do this is BUCCANEER . 1. Open the “Model Building” category in the CCP4i and select the “Buccaneer—autobuild/refine” task. 2. As “Work SEQ in,” specify a fasta file with your protein’s sequence and as “Work MTZ in,” select the MTZ file with the correct solution from SHELX. Select to “Use PHI/FOM instead of HL coefficients” and make sure that the labels for PHI and FOM columns are selected. Select Run ! Run Now to start the job.

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3. BUCCANEER will perform several cycles of model building and refinement. After the job is finished, examine the model in COOT. A new subdirectory in the CCP4 project directory will also be created that contains a refine.mtz file with map coefficients for the refined structure for generation of 2Fo-Fc and Fo-Fc maps in COOT (File ! Auto Open MTZ). Evaluate the generated model, remove incorrectly built parts, and extend and complete the model with modeling tools available in COOT. If the structure was solved using molecular replacement, the input model often serves as the starting material for building the new structure. In cases where the newly solved structure is of the same protein as the input model, the model will likely require only minor modifications such as remodeling of surface-exposed loops and side chains (given of course that the input model was well built). If the structure additionally contains a ligand or substrate bound to the protein, these of course then have to be modeled de novo (see Note 27). In cases where the solved structure is of another, yet rather similar protein, the conserved residues between the two proteins are already likely in the correct place and will need only minor adjustment. The rest of the structure is most conveniently built by systematically moving through the input model and mutating and fitting into electron density the sequence of the new structure, remodeling loops, and building additional secondary structure elements that might be present in the new structure. If the new structure is significantly different from the model (i.e., low sequence identity, or the input model covers only a part of the new structure with significant portions missing), the initial model can also be auto-built with software such as BUCCANEER (run some cycles of refinement in REFMAC first to generate the FOM column required by BUCCANEER). When it is felt that no further improvement can be made to the structure by additional modeling, the model is subjected to refinement in which the atomic coordinates and other parameters are adjusted to achieve the best fit to experimental data. A universally used measure to express the agreement between the model and the X-ray data is the R-factor; the lower the R-factor, the better the model explains the experimental data. An important variety of the R-factor is the “free” R-factor which is calculated from a small subset (usually 5%) of randomly selected reflections that have been set aside immediately after data reduction and are never used in refinement. A correct model should agree well with all reflections, not only those used for refinement, and the small test set of “free” reflections serves as a means of cross-validation for this purpose. If modeling and refinement actually make the model a better representation of the real crystal, both R-work and R-free will decrease in a similar manner and R-free will be only a few

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percent higher than R-work. The opposite is a sign of overfitting the model to noise rather than the true signal; hence R-free is probably the most useful indicator to be monitored during the course of model building and refinement. In practice, refinement of the structure within the CCP4 suite is done using the program REFMAC. 1. Open the “Refinement” section and “Restraint Preparation” subsection in the CCP4i and select the “Run Refmac5” task. 2. As “MTZ in,” select the MTZ file with the scaled and merged intensities from SCALA and as the “PDB in” select your latest model built in COOT. Select Run ! Run Now to start the job. 3. After the job has finished, double-click the job to see the results. The initial and final R-work and R-free are reported near the top of the page. At the bottom of the page, click the COOT button to load the refined model and electron density for examination in COOT. Usually several iterations of refinement and model building are required until the model is finished. As the structure nears completion, increasingly more attention needs to be devoted to validating that the model is consistent with the known principles of macromolecular structure. General indicators of model geometry are root-mean-square deviations (RMSDs) from ideal bond lengths and angles, and for realistic structures these should not exceed approximately 0.015 Å and 2 , respectively. These values are reported by refinement programs and can be controlled by adjusting the X-ray/stereochemistry weighting term in refinement settings; however, excessively high values might indicate serious problems with the model which cannot merely be fixed by tightening the geometric restraints. To examine local model quality and detect possible model errors, a variety of tools exist under the Validation menu in COOT for finding unmodeled parts of density, checking backbone and side-chain conformations, detecting clashes between atoms, and performing other analyses. The structure can also be submitted to external services such as MolProbity (http:// molprobity.biochem.duke.edu) or the PDB validation server (https://validate-rcsb.wwpdb.org) for comprehensive validation. Unless they are strongly justified by electron density, there should be no or very little geometry outliers in a well-refined structure. Still, despite the best efforts, it is understandably not always possible to build a flawless model, particularly at low resolution, but nonetheless, an experimenter should always strive, and has a certain responsibility in front of the rest of the scientific community that will build up on this work, to provide a maximally realistic model when interpreting the available data.

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3.7 Structural Analysis

The tertiary structure of a protein is generally more conserved than its sequence, and as the collection of known protein threedimensional structures continues to grow, more and more insight can be gained by comparative structure-based analyses. A popular tool for comparing protein three-dimensional structures is the Dali server, which features an option to query your newly solved structure against the existing entries in the PDB. The primary output of Dali analysis lists structurally similar proteins, which can be further selected for in-browser visualization as either aligned amino acid sequences or three-dimensional superimpositions. Overall, the Dali analysis will uncover any potentially related protein structures, which can aid in structural and functional classification of the protein of interest. In the case of the Enc34 SSB, which prior to structure determination was regarded as a hypothetical protein having apparent sequence-level homology only to other uncharacterized proteins, Dali analysis conclusively showed its relatedness to the gp2.5 protein from bacteriophage T7 and allowed us to uncover a cluster of four T7-type SSB structures. As a quantitative measure of similarity, the Dali analysis outputs a Z-score for each hit; however, a more often used measure for expressing similarity of two structures is the root-mean-square deviation (RMSD) between corresponding atomic positions in superimposed protein structures. Dali reports RMSDs in its output but these can also be calculated manually in COOT (Calculate ! SSM Superpose and Calculate ! LSQ superpose; look in terminal output for the RMSD value) or using the CCP4 program SUPER POSE. Another Web-based tool for examining a newly solved crystal structure is PDBePISA (PISA for short) which offers extensive analysis on intermolecular interactions within the crystal and calculations of solvent-accessible and buried surface areas. As crystallization conditions occasionally produce the native quaternary protein structure, the PISA service will also evaluate the potential significance of monomer complexation. The PISA significance assessments, however, are only predictions that should be validated by experiments, e.g., gel filtration. Additionally, PISA analysis on interfaces details all hydrogen, disulfide, and covalent bonds and salt bridges between selected molecules, which can be a very helpful guide when manually inspecting the structure in 3D visualization software such as PyMOL. For Enc34 SSB, PISA was used primarily to calculate the intermolecular interface areas and to identify residues involved in monomer-monomer and protein-DNA contacts. Most of the model visualization, structural analysis, and creation of publication-grade images can be done in the open-source software PyMOL, the very basics of which can be found in the PyMOL User’s Manual online. While it is possible to carry out many essential actions through its graphical user interface and the command line, in many cases it is convenient to create, edit, and

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run PyMOL macros, which are plaintext files that list a number of PyMOL commands to be sequentially executed. The most comprehensive resource for all-things PyMOL, including commands and tutorials, is the PyMOL Wiki (https://pymolwiki.org/index.php/ Main_Page). A good place to start with any new structure is to manually check the hydrogen bonding at the boundaries of secondary structure elements in the structure because PyMOL’s automatic secondary structure assignment is not always correct, and create a PyMOL macro for redefining secondary structure elements (command “alter”) that can be reloaded in all relevant visualization steps. Additional macros for defining objects and atom selections and their formatting are also quite useful, especially when preparing figures. In the study of Enc34 SSB, PyMOL was used to visualize all structures, to create structural alignments with related proteins (command “super”), and to examine the SSB-ssDNA, as well as protein-protein interactions (see Note 28). Another useful way to analyze your SSB structure is to calculate the electrostatic surface of the protein to reveal positively and negatively charged areas and hydrophobic patches on the molecular surface. Analysis of such regions can provide useful hints on how the protein interacts with DNA or other proteins and can also provide interesting hypotheses for subsequent experimental exploration. Calculation and representation of the electrostatic surface can be achieved via the “APBS Tools” plug-in in PyMOL, instructions for setting up and using which can be found in PyMOL wiki (https://pymolwiki.org/index.php/APBS and links therein) and will not be repeated here. Lastly, if the structure contains a ligand or substrate bound to the protein, it is often informative, and generally required upon publication, to calculate a type of electron density map of the ligand (ssDNA in case of the Enc34 SSB) called an “omit map,” i.e., an electron density map calculated from a model with the ligand omitted. The resulting 2Fo-Fc and Fo-Fc maps should regardless display electron density corresponding to the ligand, thus validating the interpretation of the structure. An omit map can be created using REFMAC and CCP4i as follows: 1. Open your .pdb file in COOT and change the occupancy of ligand to zero (Edit ! Residue Info). Save the file for later use. 2. Prepare a REFMAC job with the latest .mtz file in the refinement and the saved .pdb from step 1 as the input files. Choose to do either a “restrained refinement” using “no prior phase information” or a “rigid body refinement” with zero cycles. Run the job and inspect the resulting map in COOT which should show significant unmodeled density around the ligand. 3. Start an FFT job in CCP4i, select “simple map,” and select the output .mtz from REFMAC as the input file. For a Fo-Fc map, select DELFWT for F1 and PHDELWT for PHI, and for a

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2Fo-Fc map, select FWT for F1 and PHWT for PHI. Add the extension .ccp4 to the output map so it can be recognized by PyMOL. Select Run ! Run Now to start the job. 4. Visualize the map in PyMOL as described here: https:// pymolwiki.org/index.php/Display_CCP4_Maps

4

Notes 1. The protease is 6
His-tagged and prepared according to the protocol by Tropea et al., 2009 [6]. 2. For initial screening of crystallization conditions, we recommend using the sparse-matrix solution kits JCSG-plus and Structure Screens 1 and 2, as well as grid-matrix screens Clear Strategy and PACT premier (Molecular Dimensions). However, a number of further screens are offered from several companies for continued trials with similar outcomes. 3. Other frequently used options are the Strep-tag which employs binding of an eight-residue peptide to an engineered version of streptavidin [7], or genetic fusion with the maltose-binding protein which can be purified by selective binding to dextrin [8]. 4. Unstructured regions can be identified using secondary structure prediction or other software tools such as JPred (http:// www.compbio.dundee.ac.uk/jpred) or PSIPRED (http:// bioinf.cs.ucl.ac.uk/psipred). 5. The StuI site can also be utilized to excise and replace the DNA insert with another gene of interest to easily create a new expression construct. 6. For successful phasing, as a rule of thumb, the protein must contain at least one selenomethionine residue per every 100 amino acids. If there are not enough methionine residues in the protein, additional ones can be introduced by sitedirected mutagenesis of amino acids such as leucine or isoleucine [9]. Alternatively, the protein can be additionally labeled with selenocysteine [10]. 7. By a series of small-scale protein production experiments, we found that decreased growth temperatures correlated with increased SeMet-ORF6 solubility. In all cases protein production was induced with 1 mM IPTG. 8. A recommended standard buffer for crystallization is 20 mM Tris–HCl (pH 7.5–8.0). Generally, the crystallization buffer should contain as little additional ingredients (salts, additives) as possible to avoid unnecessary interference with components in crystallization solutions, unless these are strictly necessary

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for stability of the protein. Phosphate-based buffers (e.g., PBS) are not recommended as magnesium and calcium ions often present in crystallization solutions will readily form phosphate crystals, greatly increasing the rate of false-positive crystallization hits. The full-length ORF6 protein was quite stable in a 20 mM Tris buffer and had a shelf life of a couple of weeks at 4  C in the absence of NaCl, but the deletion of the C-terminus apparently rendered it significantly less stable and at least 100 mM NaCl was required in all buffer solutions to keep the protein from precipitating. 9. In case of the selenomethionine-substituted protein, the TN buffer must be supplemented with 5 mM DTT; otherwise the protein might form soluble aggregates (in case of the Enc34 SSB this was verified by a series of gel filtration experiments) which impede further purification. All other buffers used in subsequent purification should contain 1 mM DTT (not noted hereafter). Prepare the DTT-supplemented buffers right before use. 10. For sample volumes above ~2.2 mL, one 5 mL column will not suffice to separate proteins from imidazole. Depending on sample volume you can stack up to four such columns, or alternatively dialyze against the TN buffer overnight. 11. Elute bound proteins with 3 mL of buffer B and check for the presence of uncleaved protein on SDS-PAGE. A significant amount of uncleaved protein suggests insufficient amount or low activity of TEV protease, in which case the amount of TEV protease and/or the incubation time should be increased. 12. Concentration of a purified protein preparation is most easily determined by measuring absorbance at 280 nm (A280) and calculating an extinction coefficient from the sequence from which the molarity and concentration in mg/mL can be calculated. Online services such as ProtParam (https://web.expasy. org/cgi-bin/protparam/protparam) can be used for such calculations. 13. Single-stranded DNA oligonucleotides of desired sequence and length can be purchased from a variety of commercial nucleic acid synthesis services. Make sure to order a number of small-scale (e.g., ~40 nmol) aliquots of desalted lyophilized oligonucleotide. Calculate (or refer to the datasheet provided with the oligonucleotide) the molar amount of the DNA and add an appropriate volume of a 10 mL/mg SSB solution so that the DNA is in a 1.5- to 2-fold molar excess over the protein. 14. Due to the small liquid volumes, it is important to recognize that the drops will quickly evaporate if left uncovered for more than a few minutes. This is not a serious issue if using a

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pipetting robot with a 96-tip setup where the bottom solution can be added at once to all 96 wells. If using an 8-tip, 4-tip, or similar setup, a method to cover the finished drops should be devised, such as a sliding cover or a similar contraption. If this is not possible, consider using larger volumes for the drops (e.g., 1 μL). If using non-disposable tips, make sure to include appropriate steps for washing the tips to avoid crosscontamination of crystallization solutions. 15. Make sure that the sheet is firmly pressed down and well adhered to the plate and that no wrinkles or bubbles between the sealing sheet and the flat top surface of the plate are present. 16. Crystallization trials are usually done at or near room temperature in a temperature-controlled room or thermostat. While different incubation temperatures can be tested (e.g., 4, 15, or 30  C), in our experience this has not resulted in any significant improvements. If it is not possible to provide temperaturecontrolled conditions, care must be taken at least not to leave the plates at a location subject to direct sunlight or near devices producing vibration and heat (e.g., a benchtop centrifuge). 17. Immediately after setting up the crystallization plate, clear drops can be noted as the most perspective for developing subsequent crystal growth, although quite often crystals can start growing also in wells with precipitation. Large crystals that appear immediately after setting up the drops are most likely of salt but regardless should not be automatically discarded. A very fine crystalline precipitate likely suggests that the protein is prone to crystallization under these conditions, but the composition of the mixture is suboptimal, with likely too high concentration of the precipitant. When crystals form under several conditions, try to spot any similarities and patterns in crystal shapes and chemical compositions of the drops, which will help to recognize the best starting conditions for subsequent optimization trials. It is also possible that a protein crystallizes in several different forms; in such cases optimization of all crystal forms should be pursued as it is impossible to tell which of those would diffract the best. Do not judge a crystal only from its looks, as more than once the ugliest crystals have yielded the best data while the beautiful ones have proven completely useless. 18. For example, if the crystallization conditions contain a PEG precipitant, NaCl, and a Tris–HCl buffer at pH 8.0, prepare several 1D or 2D grids of various PEG vs. NaCl concentrations around the original values while leaving the buffer unchanged, or explore buffers at different pH while leaving everything else the same. The effect of different chain-length polymer

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precipitants (e.g., PEG 4000 vs. PEG 3000 or PEG 6000) or different cations or anions (e.g., NaCl vs. KCl or NaBr) can also be tested to see if they improve the crystals. 19. To alleviate radiation damage, X-ray diffraction data from crystals are collected at a cryogenic (~100 K) temperature. To avoid formation of ice crystals that will damage the protein crystal, before cooling the crystal is submerged in a crystallization solution supplemented with a cryoprotecting agent. The most frequently used cryoprotectant in protein crystallography is glycerol at a final concentration of around 30%; however, occasionally the cryoprotecting substances can also damage the crystal, and alternatives such as ethylene glycol, sucrose, or 2-methyl-2,4-pentanediol (MPD) can be tried. Several crystallization conditions already contain significant amounts of cryoprotecting substances (e.g., low-molecular-weight PEGs, MPD, very high concentrations of salt); in such cases, the crystal can be frozen directly from the drop. 20. Users have to apply for beamtime at synchrotron facilities to get an allocated time slot for their experiments. Most often the facilities’ issue calls for project proposals once or twice a year, yet others operate on a rolling-schedule basis when proposals can be submitted at any time. Data collection should regardless be planned well in advance as it may take up to 6 months to get to data collection from the moment of submitting a proposal. 21. The state-of-the-art beamlines are currently capable of operating at such speed that it might be feasible to collect a full dataset from any crystal of acceptable quality rather than screen all crystals first and then remount and collect data from the best one(s). The downside of such a strategy is the generation of huge amounts of data, which is however becoming increasingly less of a problem due to modern data storage, transfer, and computing capabilities. 22. The predicted and actual spot positions should closely match. If this is not the case, try selecting the topmost (P1) solution to see if this sufficiently improves the fit. If it does not, or in case autoindexing fails altogether, try manually picking different images for autoindexing, avoiding frames with very weak diffraction or serious diffraction defects such as smeared and weakly separated spots. 23. POINTLESS will output the data in the most probable automatically determined space group. However, the log file from the task should always be examined where near the bottom the probabilities of different space groups will be listed. In case of high uncertainties, it might be necessary to try other space group options in the experimental phasing or molecular replacement steps. Also, some sets of space groups remain

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undistinguishable from the diffraction pattern (e.g., space groups P 61 and P 65) and the correct one needs to be determined empirically by testing both options in subsequent phasing steps. If the determined crystal system is different than that used for data integration, POINTLESS will recommend to rerun data integration in the new space group; this advice should be followed. 24. In the Results viewer, inspect the “Analysis against resolution” and “Correlations within dataset” tables. As a rule of thumb, the mean I/σI (Mn(I/sd)) should be at least 1.0 and the mean half-set correlation (CC(1/2)) at least 0.3 in the highest resolution bin. If the data are to be used for experimental phasing (i.e., from a selenomethionine-labeled crystal), the anomalous correlation coefficient (CC_anom) should be at least 0.3 and the RMS correlation ratio (RCR_anom) at least 1.5 to a resolution of at least 3 Å to suggest a successful structure solution. 25. Structure solution using molecular replacement (MR) requires that a model of sufficiently similar structure is already available. Similarity in three-dimensional structure correlates with sequence similarity, and over 35–40% sequence identity usually implies that the proteins are sufficiently similar for MR to work. In the range of 20–35% sequence identity, MR might work in some cases but often requires extensive efforts using different MR protocols and starting models. Below 20% sequence identity, MR will almost certainly fail. It is therefore recommended that in case the closest homolog of known structure shares less than some 35% of sequence identity with the protein of interest, experimental phasing is pursued as the primary approach for structure determination. 26. COOT is the gold standard tool for building macromolecular models at an atomic resolution. COOT is a very comprehensive and feature-rich software, and detailed descriptions of all aspects of model building are beyond the scope of this chapter. New users should familiarize themselves with COOT using the documentation and tutorials available online (https://www2. mrc-lmb.cam.ac.uk/personal/pemsley/coot/). 27. In case of building DNA, which is of relevance to SSBs, some useful tools in COOT include the “Build Nucleic Acid” and “Ideal DNA/RNA” features found under Calculate ! Other Modeling Tools. Alternatively, a single nucleotide can be first placed (File ! Get Monomer or File ! Search Monomer Library) and fitted into density, after which the DNA strand can be extended using the Add Residue option in the toolbar. 28. Stacking interactions were identified visually, but the hydrogen bonds and salt bridges were taken from PISA and drawn using the Measurement wizard in PyMOL.

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References 1. Cernooka E, Rumnieks J, Tars K, Kazaks A (2017) Structural basis for DNA recognition of a single-stranded DNA-binding protein from Enterobacter phage Enc34. Sci Rep 7:15529 2. Marintcheva B, Marintchev A, Wagner G, Richardson CC (2008) Acidic C-terminal tail of the ssDNA-binding protein of bacteriophage T7 and ssDNA compete for the same binding surface. Proc Natl Acad Sci U S A 105:1855–1860 3. Studier FW, Moffatt BA (1986) Use of bacteriophage T7 RNA polymerase to direct selective high-level expression of cloned genes. J Mol Biol 189:113–130 4. Green MR, Sambrook J (2012) Molecular cloning: a laboratory manual. Cold Spring Harbor, New York 5. Dauter Z (2010) Carrying out an optimal experiment. Acta Crystallogr D Biol Crystallogr 66:389–392

6. Tropea JE, Cherry S, Waugh DS (2009) Expression and purification of soluble His(6)tagged TEV protease. Methods Mol Biol 498:297–307 7. Schmidt TG, Skerra A (2007) The Strep-tag system for one-step purification and highaffinity detection or capturing of proteins. Nat Protoc 2:1528–1535 8. Lebendiker M, Danieli T (2011) Purification of proteins fused to maltose-binding protein. Methods Mol Biol 681:281–293 9. Leahy DJ, Erickson HP, Aukhil I, Joshi P, Hendrickson WA (1994) Crystallization of a fragment of human fibronectin: introduction of methionine by site-directed mutagenesis to allow phasing via selenomethionine. Proteins 19:48–54 10. Strub MP, Hoh F, Sanchez JF, Strub JM, Bo¨ck A, Aumelas A, Dumas C (2003) Selenomethionine and selenocysteine double labeling strategy for crystallographic phasing. Structure 11:1359–1367

INDEX A Antibacterial drugs ........................................................ 118 Archaea ............................................................... 10, 23–45 Atomic force microscopy (AFM) .................93, 241–256, 258, 260 ATPase ....................................................... 68, 75–77, 314

Forces...............................................................93–99, 102, 104–109, 113, 178, 185, 241, 242, 251, 255, 259, 273, 274, 276, 277, 280, 283–287, 292, 296–300

G Genome maintenance ............................. 4, 9, 13, 95, 108

B Bacteriophage Phi29.............................................. 13, 333 Biosensor .............................................................. 217, 219

C Cancer........................................................................7, 193 Construct design ......................................... 345, 347, 349 ConSurf ........................................................68, 70–72, 78

H Helix-destabilization activity ........................................ 334 High-throughput screening (HTS) .................... 117–132 hSSB1 ................................................................6, 8, 9, 25, 229–231, 235–239 hSSB2 .................................................... 8, 9, 25, 229–239

I

D

Immunofluorescence .................................................... 209

DNA binding proteins ............................................... 7, 9, 13, 23, 34, 170, 265, 323–331, 337, 338, 340 DNA-protein complex nucleoprotein complexes .................................. 274 protein-DNA interactions .................94, 223, 226 protein-DNA organization ............................... 274 DNA curtains ....................................................... 193–206 DNA helicase..........................................68, 81, 136, 265, 313–321, 323, 329 DNA polymerase γ................................................................................ 314 DNA repair .............................7–9, 25, 33, 136, 193, 135 DNA replication mitochondrial ................................................. 274, 304 Drosophila ........................................... 241–260, 303–310 dsDNA unwinding assays ............................................................... 314, 318

K

E

O

Electron microscopy (EM) ..........................179, 265–271

OB-fold............................................................1, 2, 4, 6, 8, 11–13, 24, 26, 28, 29, 31, 33–37, 39, 40, 170, 171, 181 Optical tweezers ................................... 98, 106, 273–287, 289, 290, 292–298

F Fluorescent in situ hybridization (FISH) .......... 209, 210, 212–214

Kinetics real-time.................................................... 50, 289–300

L Leishmania ........................................................... 169–190

M Magnetic tweezers............................................ 93–98, 108 Mitochondrial single-stranded DNA-binding protein (mtSSB)..................................... 6, 9, 265–271, 304–311, 313–321

N Nuclear Magnetic Resonance (NMR) ............. 31–33, 35, 40, 129, 229–239

Marcos T. Oliveira (ed.), Single Stranded DNA Binding Proteins, Methods in Molecular Biology, vol. 2281, https://doi.org/10.1007/978-1-0716-1290-3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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376 Index P

Phylogenetics........................................... 3, 26–29, 70, 72 PriA ..............................................3, 4, 68, 71, 78, 81, 82, 84, 85, 118, 119, 121, 122, 129, 130 Primase ........................3, 10, 39, 68, 314, 323–325, 329 Protein dynamics ..................................................6, 37, 49, 50, 169–190, 230, 243, 274 production ..................... 81, 344, 347, 349, 350, 367 purifications ................................................. 82, 83, 87, 129, 152, 159, 232, 235–238, 250, 344, 347, 348, 351, 352, 369 Protein-protein interactions ........................ 6, 24, 25, 81, 118, 152, 217, 367

Single-stranded DNA (ssDNA) binding protein (SSB) SsbA ..................................................................... 68 intermediates ...................................98, 313, 323, 344 Srs2 .................................... 195, 196, 198, 203, 204, 206 Stalled fork rescue ........................................................... 81 Stimulation ............................................... 68, 75–77, 109, 112, 265, 266, 268, 270, 313–321, 334 Structural analysis ...................8, 31, 40, 43, 170, 180, 243, 366 biology ..................................................................... 344 determination .......................................................... 366 Supramolecular complexes .......................................49–64 Surface plasmon resonance (SPR).......................... 49, 68, 71–75, 129, 217–227

R

T

Rad51 .....................................................7, 135, 141, 142, 193–196, 198, 201–205 RecA..........................................4, 40, 135, 141, 142, 193 RecG ..........................................................................81–85 Recombination homologous ............................. 7, 135, 136, 193, 194 RecQ helicase ..........................................................93–113 Replication Protein A (RPA) ........................ 5–11, 23–41, 43, 142, 146, 151–167, 169–171, 177, 181, 194, 196, 197, 200, 201, 203, 206, 210, 213, 220, 229 RNA interference (RNAi) ................................... 303–310

Telomere end-binding proteins (TEBP) ............ 170, 171 Telomeres telomeric DNA...........................................7, 212, 220 Tethered particle motion (TPM) ........................ 135–148 Trypanosoma cruzi .............................................. 171, 182, 209–215, 217–227 Trypanosomatids .................................170, 171, 181, 182 Twinkle P66........................................ 314, 315, 317, 318, 320

S Schneider cells .....................................303, 305–307, 310 Single-molecule manipulation............................................................ 273 microscopy................................................................. 56

V Ver ...................................... 243, 244, 246, 254, 256, 258 Volume analysis ............................................................. 256

X X-ray crystallography ........................................31, 33, 40, 41, 43, 344, 347, 350, 358