Helicases: Methods and Protocols (Methods in Molecular Biology, Vol 587)

METHODS

IN

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

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

For other titles published in this series, go to www.springer.com/series/7651

Helicases Methods and Protocols

Edited by

Mohamed M. Abdelhaleem University of Toronto, Toronto, ON, Canada

Editor Mohamed M. Abdelhaleem Hospital for Sick Children Department of Paediatric Laboratory Medicine 555 University Ave. Toronto ON M5G 1X8 Canada [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-60327-354-1 e-ISBN 978-1-60327-355-8 DOI 10.1007/978-1-60327-355-8 Library of Congress Control Number: 2009936373 # Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer ScienceþBusiness Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper springer.com

Preface The objective of this book is to provide the scientific community with the current methods used to study helicases, the enzymes that utilize the energy derived from nucleoside triphosphate (NTP) hydrolysis to unwind the double-stranded helical structure of nucleic acids. The book starts with an overview chapter that provides a brief introduction of helicases, with few examples of their role in nucleic acid metabolism. This chapter is intended for readers new to the field. The chapters that follow are written by leading international scientists who contributed significantly to our current understanding of helicases. In these chapters, the reader finds methods for the production and purification of helicases from different species as well as detailed studies of helicase activities, including NTP binding and hydrolysis, nucleic acid binding and unwinding, and translocation along nucleic acid substrates. Helicase activities are generally measured by methods that rely on radiometric, enzymatic, and fluorescence-based techniques. As described in Chapter 2, fluorescence probes have high sensitivity and rapid response that permit real time analysis. The ability to use fluorescence probes to investigate helicase activity at the single molecule level has provided significant insights into the mechanism of helicase function. A general guide to such assays is given in Chapter 3. Another advance in helicase studies is the use of pre-steady state kinetic techniques, as described for the translocase activity along single-stranded nucleic acids (Chapter 4) and for the DNA unwinding and polymerization activities of bacteriophage T7 molecule (Chapter 5). In addition to nucleic acid unwinding, helicases are involved in the disruption of protein–nucleic acid interactions. Chapter 6 has four examples of protein displacement by helicases involved in various steps of nucleic acid metabolism. As helicases occur in vivo as part of molecular complexes that include nucleic acid and protein, characterization of their protein and nucleic acid interactions provides insights into their in vivo roles. Chapter 7 describes a protocol to investigate protein–protein interactions in vivo using tandem affinity purification. Chapter 8 describes the use of chromatin immunoprecipitation (ChIP) to determine nucleic acid targets of DNA helicases. The following three chapters describe methods to study DNA helicases involved in replication (Chapter 9), transcription termination (Chapter 10), and recombination (Chapter 11). The RecQ family of DNA helicases is involved in DNA repair and recombination and has been the subject of intense research because of their roles in maintaining genome stability. Three chapters are devoted to the study of RecQ helicases from different species. A protocol for the production and characterization of mutants of the helicase core of human Bloom syndrome gene (BLM) is described in Chapter 12. This protocol can be adapted to study other helicases. Chapter 13 describes the role of Drosophila Blm helicase in double-stranded gap repair. Protocols for the expression and characterization of RecQ helicases from the plant model Arabidopsis thaliana are described in Chapter 14. v

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The following chapters describe methods to study hepatitis C virus (HCV) nonstructural protein 3 (NS3), a potential therapeutic target for the liver disease caused by HCV. Chapter 15 describes a fluorescence-based high-throughput assay to test inhibitors of NS3 helicase activity. In addition to the C-terminal helicase domain, NS3 protein has an N terminal serine protease. A method to simultaneously monitor the helicase and protease activities of HCV NS3 is described in Chapter 16. This method could be used to identify dual NS3 inhibitors. In Chapter 17, computational techniques to study NS3 are described. RNA helicase members of the DExD/H-box family of proteins are involved in all aspects of cellular RNA metabolism. Chapter 18 provides a protocol for the quantitative evaluation of the unwinding activity of the largest family of RNA helicase, the DEAD-box proteins. Examples of the methods used to study the versatile roles played by RNA helicases include the role of Ddx5 in transcription (Chapter 19), the role of DDX3 in HIV infection (Chapter 20), and the activities of DHX9 (RNA helicase A and nuclear DNA helicase II) (Chapter 21) and its drosophila homolog maleless protein (Chapter 22). Chapter 23 describes cloning, expression and activities of the human RNA helicase, Upf1, which is involved in non-sense mediated decay of mRNA. Helicases are involved in regulating the mitochondrial genome. Chapter 24 provides protocols to study the activities of the mitochondrial degradosome, including the activity of its helicase component, Suv3p. Chapter 25 describes the helicase and antitelomere activities of Pif1p, a functionally versatile helicase involved in regulating both the mitochondrial and nuclear genomes. The last two chapters illustrate two applications of helicase research in agriculture and medicine. Chapter 26 describes a method to confer salinity stress tolerance to plants by overexpression of a DNA helicase. Chapter 27 describes the potential targeting of helicases to inhibit the growth of malaria parasites. This book has only been made possible by the contributions from the authors. I would like to thank all of them for their cooperation and timely submissions. I am grateful to the series editor, John Walker, for his helpful comments and editorial expertise. Finally, I would like to acknowledge the support of my family. Mohamed M. Abdelhaleem

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

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Helicases: An Overview. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mohamed Abdelhaleem

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Fluorescent Biosensors to Investigate Helicase Activity . . . . . . . . . . . . . . . . . . . . . Martin R. Webb

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Single-Molecule FRET Analysis of Helicase Functions . . . . . . . . . . . . . . . . . . . . . . Eli Rothenberg and Taekjip Ha

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Kinetics of Motor Protein Translocation on Single-Stranded DNA . . . . . . . . . . . . Christopher J. Fischer, Lake Wooten, Eric J. Tomko, and Timothy M. Lohman

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Experimental and Computational Analysis of DNA Unwinding and Polymerization Kinetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Manjula Pandey, Mikhail K. Levin, and Smita S. Patel

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Protein Displacement by Helicases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Laxmi Yeruva and Kevin D. Raney

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In Vivo Investigation of Protein–Protein Interactions for Helicases Using Tandem Affinity Purification. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Matthew Jessulat, Terry Buist, Md Alamgir, Mohsen Hooshyar, Jianhua Xu, Hiroyuki Aoki, M. Clelia Ganoza, Gareth Butland, and Ashkan Golshani

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Mapping Genomic Targets of DNA Helicases by Chromatin Immunoprecipitation in Saccharomyces cerevisiae . . . . . . . . . . . . . . . . . . . . . . . . . . 113 Jennifer Cobb and Haico van Attikum

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Methods to Study How Replication Fork Helicases Unwind DNA . . . . . . . . . . . . 127 Daniel L. Kaplan and Irina Bruck

10. Simple Enzymatic Assays for the In Vitro Motor Activity of Transcription Termination Factor Rho from Escherichia coli. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 137 Marc Boudvillain, Ce´line Walmacq, Annie Schwartz, and Fre´de´rique Jacquinot 11. Single-Molecule Studies of RecBCD. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155 Thomas T. Perkins and Hung-Wen Li 12. Mutational Analysis of Bloom Helicase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 173 Xu Guang Xi 13. In Vivo Analysis of Drosophila BLM Helicase Function During DNA Double-Strand Gap Repair . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 185 Mitch McVey 14. Purification and Characterization of RecQ Helicases of Plants . . . . . . . . . . . . . . . . 195 Daniela Kobbe, Manfred Focke, and Holger Puchta

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15. Fluorometric Assay of Hepatitis C Virus NS3 Helicase Activity . . . . . . . . . . . . . . . 211 Mariusz Krawczyk, Anna Stankiewicz-Drogon´, Anne-Lise Haenni, and Anna Boguszewska-Chachulska 16. A Method to Simultaneously Monitor Hepatitis C Virus NS3 Helicase and Protease Activities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 223 David N. Frick, Olya Ginzburg, and Angela M.I. Lam 17. Computer Modeling of Helicases Using Elastic Network Model . . . . . . . . . . . . . . 235 Wenjun Zheng 18. Duplex Unwinding with DEAD-Box Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . 245 Eckhard Jankowsky and Andrea Putnam 19. Analysis of the RNA Helicase p68 (Ddx5) as a Transcriptional Regulator . . . . . . . 265 Samantha M. Nicol and Frances V. Fuller-Pace 20. A Method to Study the Role of DDX3 RNA Helicase in HIV-1 . . . . . . . . . . . . . . 281 Chia-Yen Chen, Venkat R.K. Yedavalli, and Kuan-Teh Jeang 21. Molecular Characterization of Nuclear DNA Helicase II (RNA Helicase A) . . . . . 291 Suisheng Zhang and Frank Grosse 22. Regulation of Inter- and Intramolecular Interaction of RNA, DNA, and Proteins by MLE. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 303 Hyangyee Oh, Andrew M. Parrott, Yongkyu Park, and Chee-Gun Lee 23. Biochemical Characterization of Human Upf1 Helicase . . . . . . . . . . . . . . . . . . . . . 327 Zhihong Cheng, Gaku Morisawa, and Haiwei Song 24. Assays of the Helicase, ATPase, and Exoribonuclease Activities of the Yeast Mitochondrial Degradosome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 339 Michal Malecki, Piotr P. Stepien, and Pawel Golik 25. Characterization of the Helicase Activity and Anti-telomerase Properties of Yeast Pif1p In Vitro . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 359 Jean-Baptiste Boule´ and Virginia A. Zakian 26. A Method to Confer Salinity Stress Tolerance to Plants by Helicase Overexpression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 377 Narendra Tuteja 27. A Method to Inhibit the Growth of Plasmodium falciparum by Double-Stranded RNA-Mediated Gene Silencing of Helicases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 389 Renu Tuteja Subject Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 401

Contributors MOHAMED ABDELHALEEM • Department of Paediatric Laboratory Medicine, The Hospital for Sick Children, University of Toronto, Toronto, ON, Canada MD ALAMGIR • Department of Biology and Ottawa Institute of Systems Biology, Carleton University, Ottawa, ON, Canada HIROYUKI AOKI • Banting and Best Department of Medical Research, University of Toronto, Toronto, ON, Canada ANNA BOGUSZEWSKA-CHACHULSKA • Genomed, Warsaw, Poland MARC BOUDVILLAIN • Centre de Biophysique Moleculaire (UPR4301), CNRS, Orleans cedex 2, France JEAN-BAPTISTE BOULE • Department of Molecular Biology, Princeton University, Princeton, NJ, USA IRINA BRUCK • Department of Biological Sciences, Vanderbilt University Nashville, TN, USA TERRY BUIST • Department of Biology and Ottawa Institute of Systems Biology, Carleton University, Ottawa, ON, Canada GARETH BUTLAND • Life Science Division, Lawrence Berkeley National Laboratory, Berkeley, CA, USA CHIA-YEN CHEN • Molecular Virology Section, Laboratory of Molecular, Microbiology, the NIAID, NIH, Bethesda, MD, USA ZHIHONG CHENG • Cancer and Developmental Cell Biology Division, Institute of Molecular and Cell Biology, A*STAR (Agency for Science, Technology and Research), Singapore, Singapore JENNIFER COBB • Department of Biochemistry and Molecular Biology, Southern Alberta Cancer Research Institute, University of Calgary, Calgary, AB, Canada CHRISTOPHER J. FISCHER • Department of Physics and Astronomy, University of Kansas, Lawrence, KS, USA MANFRED FOCKE • Botanik II, Universita¨t Karlsruhe (TH), Karlsruhe, Germany FRANCES V. FULLER-PACE • Centre for Oncology & Molecular Medicine, University of Dundee Ninewells Hospital & Medical School, Dundee, UK DAVID N. FRICK • Department of Biochemistry & Molecular Biology New York Medical College Valhalla, NY, USA M. CLELIA GANOZA • Banting and Best Department of Medical Research, University of Toronto, Toronto, ON, Canada OLYA GINZBURG • Department of Biochemistry & Molecular Biology, New York Medical College, Valhalla, NY, USA PAWEL GOLIK • Faculty of Biology, Institute of Genetics and Biotechnology, University of Warsaw, Warsaw, Poland ASHKAN GOLSHANI • Department of Biology and Ottawa Institute of Systems Biology, Carleton University, Ottawa, ON, Canada

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FRANK GROSSE • Leibniz Institute for Age Research, Fritz Lipmann Institute (FLI), Jena, Germany TAEKJIP HA • Department of Physics, Center for Biophysics and Computational Biology, University of Illinois at Urbana-Champaign, Urbana, IL, USA ANNE-LISE HAENNI • Institut Jacques Monod, Paris, France MOHSEN HOOSHYAR • Department of Biology and Ottawa Institute of Systems Biology, Carleton University, Ottawa, ON, Canada ECKHARD JANKOWSKY • Department of Biochemistry & Center for RNA Molecular Biology, School of Medicine, Case Western Reserve University, Cleveland, OH, USA FREDERIQUE JACQUINOT • Centre de Biophysique Moleculaire (UPR4301), CNRS, Orleans cedex 2, France KUAN-TEH JEANG • Molecular Virology Section, Laboratory of Molecular, Microbiology, the NIAID, NIH, Bethesda, MD, USA MATTHEW JESSULAT • Department of Biology and Ottawa Institute of Systems Biology, Carleton University, Ottawa, ON, Canada DANIEL L. KAPLAN • Vanderbilt University Department of Biological Sciences Nashville, TN, USA DANIELA KOBBE • Botanik II, Universita¨t Karlsruhe (TH), Karlsruhe, Germany MARIUSZ KRAWCZYK • Institute of Biochemistry and Biophysics, Polish Academy of Sciences, Warsaw, Poland ANGELA M. I. LAM • Department of Biochemistry & Molecular Biology, New York Medical College, Valhalla, NY, USA CHEE-GUN LEE • UMDNJ-New Jersey Medical School, Department of Biochemistry and Molecular Biology, Newark, NJ, USA HUNG-WEN LI • Department of Chemistry, National Taiwan University, Taipei 10617, Taiwan TIMOTHY M. LOHMAN • Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA MICHAL MALECKI • Institute of Genetics and Biotechnology, Faculty of Biology, University of Warsaw, Warsaw, Poland MIKHAIL K. LEVIN • Department of Biostatistics & Bioinformatics, Duke University Medical Center, Durham, NC, USA MITCH MCVEY • Department of Biology, Tufts University, Medford, MA GAKU MORISAWA • Cancer and Developmental Cell Biology Division, Institute of Molecular and Cell Biology, A*STAR (Agency for Science, Technology and Research), Singapore, Singapore SAMANTHA M. NICOL • Centre for Oncology & Molecular Medicine, University of Dundee Ninewells Hospital & Medical School. Dundee, UK HYANGYEE OH • HHMI, Waksman Institute, Rutgers University, Piscataway, NJ, USA MANJULA PANDEY • Department of Biochemistry, University of Medicine and Dentistry of New Jersey-Robert Wood Johnson Medical School, Piscataway, NJ, USA SMITA S. PATEL • Department of Biochemistry, University of Medicine and Dentistry of New Jersey-Robert Wood Johnson Medical School, Piscataway, NJ, USA YONGKYU PARK • UMDNJ-New Jersey Medical School, Department of Cell Biology and Molecular Medicine, Newark New Jersey, USA

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ANDREW M. PARROTT • UMDNJ-New Jersey Medical School, Department of Biochemistry and Molecular Biology, Newark, NJ, USA THOMAS T. PERKINS • MCD Biology, JILA, NIST & CU, University of Colorado at Boulder, Boulder, CO, USA HOLGER PUCHTA • Botanik II, Universita¨t Karlsruhe (TH), Karlsruhe, Germany ANDREA PUTNAM • Department of Biochemistry & Center for RNA Molecular Biology, School of Medicine, Case Western Reserve University, Cleveland, OH, USA KEVIN D. RANEY • Department of Biochemistry and Molecular Biology, University of Arkansas for Medical Sciences, Little Rock, AR, USA ELI ROTHENBERG • Department of Physics, NSF Center for the Physics of Living Cells, University of Illinois at Urbana–Champaign, Urbana, IL, USA ANNIE SCHWARTZ • Centre de Biophysique Moleculaire (UPR4301), CNRS, Orleans cedex 2, France HAIWEI SONG • Cancer and Developmental Cell Biology Division, Institute of Molecular and Cell Biology, A*STAR (Agency for Science, Technology and Research), Singapore, Singapore ANNA STANKIEWICZ-DROGON´ • Institute of Biochemistry and Biophysics, Polish Academy of Sciences, Warsaw, Poland PIOTR P. STEPIEN • Institute of Genetics and Biotechnology, Faculty of Biology, University of Warsaw, Warsaw, Poland ERIC J. TOMKO • Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA NARENDRA TUTEJA • Plant Molecular Biology Group, International Centre for Genetic Engineering and Biotechnology, New Delhi, India RENU TUTEJA • Malaria Group, International Centre for Genetic Engineering and Biotechnology, New Delhi, India HAICO VAN ATTIKUM • Department of Toxicogenetics, Leiden University Medical Center, Leiden, The Netherlands MARTIN R. WEBB • MRC National Institute for Medical Research, The Ridgeway, Mill Hill, London, UK CELINE WALMACQ • National Cancer Institute, NIH, Frederick, MD LAKE WOOTEN • Department of Physics and Astronomy, University of Kansas, Lawrence, KS, USA XU GUANG XI • Institut CURIE Recherche, Orsay, France JIANHUA XU • Banting and Best Department of Medical Research, University of Toronto, Toronto, ON, Canada VENKAT R.K. • YEDAVALLI, Molecular Virology Section, Laboratory of Molecular, Microbiology, the NIAID, NIH, Bethesda, MD, USA LAXMI YERUVA • Department of Biochemistry and Molecular Biology, University of Arkansas for Medical Sciences, Little Rock, AR, USA VIRGINIA A. ZAKIAN • Department of Molecular Biology, Princeton University, Princeton, NJ, USA SUISHENG ZHANG • Department of Biochemistry, National University of Ireland, Galway, Ireland WENJUN ZHENG • Physics Department, University at Buffalo, Buffalo, NY, USA

Chapter 1 Helicases: An Overview Mohamed Abdelhaleem Abstract Helicases are essential enzymes involved in all aspects of nucleic acid metabolism including DNA replication, repair, recombination, transcription, ribosome biogenesis and RNA processing, translation, and decay. They occur in vivo as part of molecular complexes that include the components required for each specific step of nucleic acid metabolism. The role of the helicases is to utilize the energy derived from nucleoside triphosphate hydrolysis to translocate along nucleic acid strands, unwind/separate the helical structure of double-stranded nucleic acid, and, in some cases, disrupt protein–nucleic acid interactions. Because of their essential function, helicases are ubiquitous and evolutionary conserved proteins. This chapter briefly highlights helicase structure and activities and provides examples of the helicases involved in nucleic acid metabolism. Key words: Helicases, DNA replication, DNA repair, DNA recombination, transcription, ribosome biogenesis, RNA splicing, translation, RNA degradation.

1. Helicase Structure and Activities

Helicases are characterized by the presence of conserved motifs in the form of short amino acid sequences. Based on variations of the number of motifs, their amino acid sequence, and spacing, helicases are grouped into superfamilies, including three large (SF1–SF3) and two small (SF4 and SF5) ones (1). SF1 and SF2 have at least seven conserved motifs (I, Ia, II, III, IV, V, and VI) and are monomers, whereas SF3–SF5 members assemble into hexamers (2). The crystal structures of several representative helicases were resolved (3–10) and revealed common features. Monomeric helicases (SF1/SF2) have a core that consists of two domains with a linker region. Hexameric helicases (SF3–SF5) form a core that

M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_1, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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includes six individual domains arranged in a ring. The domains are termed RecA-like because of the similarity to the ATP-binding core of RecA recombination protein. The conserved helicase motifs include those involved in NTP binding and hydrolysis, which are similar to the Walker A and B boxes of ATPase. The other conserved motifs are involved in coupling of the NTP hydrolytic state to protein conformational changes and in nucleic acid binding (2, 11–13). Helicase core structure allows for cycles of nucleic acid binding and release driven by NTP binding and hydrolysis (11). By utilizing the energy derived from NTP hydrolysis, helicases function as nucleic acid motor or translocases. Models have been proposed to describe helicase coupling of NTP hydrolysis with directional translocation and unwinding (11, 14, 15). Another group of nucleic acid motors include members of the ATPases associated with various cellular activities (AAA+) which have been proposed as SF6 of helicases/translocases (2). These molecules contain the AAA+ fold and include hexameric motor proteins such as mini chromosome maintenance (MCM) protein complexes involved in replication initiation (16). Helicase activities are described in terms of rate, processivity, directionality, and step size. Rate is the number of unwound/ translocated base pairs, or the number of NTP molecules hydrolyzed, per unit time. Processivity is the number of base pairs unwound/translocated before the helicase dissociates from the nucleic acid. Directionality is the bias exhibited by a processive helicase in its movement along nucleic acids, either 30 to 50 or 50 to 30 direction. Finally, step size has been described as either mechanical (the average distance moved) or kinetic (the average number of unwound/translocated) during each catalytic cycle (2, 11, 17, 18). Depending on the type of their nucleic acid targets, helicases are generally classified as DNA or RNA helicases. Some helicases can unwind both targets (19, 20), whereas others preferentially unwind RNA–DNA duplexes (21). DNA helicases are involved in replication, repair and recombination. RNA helicases are involved in all aspects of RNA metabolism. The majority of RNA helicases belongs to DExD/H-box proteins which have the conserved motifs and domain structure of SF2 helicases (13, 22, 23). DExD/H-box proteins derive this name from the single letter code of the four amino acids of motif II (which is equivalent to Walker B motif of ATPases). They are classified into several subgroups including DEAD-box (DDX) and DEAH-box (DHX) families, which are distinguished by consistent sequence differences that extend beyond motif II (23, 24). DEAD-box members constitute the largest subgroup and demonstrate significant functional differences compared to other helicases (13, 23, 25, 26).

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Viral DExD/H RNA helicases (e.g., NPH-II) are able to translocate in a directional and processive manner on their targets and function as molecular motors for RNA (27). RNA unwinding is one of several activities that have been demonstrated thus far for DExD/H proteins. Some DExD/H proteins can function as RNPases (enzymes that disrupt protein–RNA interactions) (26, 28), whereas others have RNA annealing activity (29). The RNA unwinding and annealing and RNPase activities are consistent with the role of DExD/H-box proteins as the major players in remodeling of the ribonucleoprotein complexes (13, 23, 30). Structural differences in the amino acid sequences of the conserved motifs and the RecA-like domain as well as differences in the mode of interaction of the helicase core with nucleic acid targets allow a particular helicase to be best suited for the nucleic acid processing step in which it is involved (2, 11–13). However, the unwinding activity of the helicase catalytic core is not sequence specific. Therefore, helicases require mechanisms to recognize, and to be loaded onto, their targets. In addition, their activities are required to be optimally and tightly regulated to prevent potentially deleterious consequences. Moreover, unwinding is only one of multiple steps that take place during nucleic acid processing. Helicase-mediated unwinding is required to be coupled with subsequent steps (31). These various requirements (target recognition and loading, regulation of activities, and coupling of unwinding) are met, at least in part, through the presence of other domains within the helicase itself and/or through protein–protein interactions between helicases and other proteins that assemble in a coordinated manner to form the in vivo molecular complexes involved in nucleic acid processing (11, 18, 31, 32).

2. Examples of Helicases Involved in Nucleic Acid Metabolism 2.1. DNA Replication

The Escherichia coli hexameric helicase DnaB (SF4 member with 50 to 30 directionality) is a component of the replisome, the molecular complex responsible for the duplication of the genetic material (33). Assembly of the replisome is initiated by initiator proteins, which recognize the origin of replication and recruit other replisome components to the DNA in a coordinated manner, including loading of the helicase into the DNA. Replication proceeds with the synthesis of one daughter strand (the leading strand) by DNA polymerase in the same direction as the unwinding of the replication fork (50 to 30 ). Synthesis of the other daughter strand (the lagging strand) occurs in the opposite direction and is initiated by a special RNA polymerase (primase), carried out by DnaG protein.

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The helicase and primase activities are closely associated and co-regulated (34). The two activities are present within the same molecule in T7 bacteriophage gene product 4 (T7gp4) (35, 36). In archaea and eukaryotes, the minichromosome maintenance (Mcm) protein members of SF6 helicase/translocase with AAA+fold are involved in DNA replication (16, 37, 38). In archaea, a single Mcm protein forms ring-shaped homo-hexamers or doublehexamers. In eukaryotes, six different members (Mcm2–Mcm7) form a heterohexameric complex. The Mcm2–Mcm7 complex is involved both in the initiation and the elongation steps of eukaryotic DNA replication. Before the initiation of replication, other replication factors, including the Cdc45 and GINS complexes, are recruited. During elongation, the Mcm complex, together with Cdc45 and GINS, unwinds double-stranded DNA to produce single-stranded templates for DNA synthesis (39–41). DNA unwinding activity has been characterized for a subcomplex composed of Mcm4, 6, 7 proteins. Recently, the entire Mcm2–Mcm7 complex has been shown to have DNA helicase activity in vitro (42). 2.2. DNA Repair

UvrD is an E. coli hexameric helicase (SF1 member with 30 to 50 directionality) involved in DNA repair. Base–base mismatches can occur as errors of DNA polymerases. The mismatch repair (MMR) process in E. coli involves recognition of the error by a MutS homodimer, which recruits a homodimer of MutL. MutS-MutL activates MutH, which incises the strand. UvrD helicase unwinds the ends of the nicked error-containing strand, followed by its exonuclease-mediated digestion by an exonuclease. The resulting gap is filled by RNA polymerase III followed by sealing the remaining nick by DNA ligase (43, 44). The UvrABC endonuclease (which consists of UvrA, UvrB, and UvrC proteins) is the central enzyme for nucleotide excision repair (NER) process in E. coli (45). UvrB is a SF2 DNA helicase, whereas UvrC is an endonuclease. The main function of UvrA is to deliver UvrB to the damage site. UvrB also assists in the location of DNA damage via a b-hairpin that protrudes from its N-core domain and inserts between the two strands of the DNA duplex. UvrA2B heterotrimer complex recognizes damaged DNA, and a bend is formed in the DNA backbone in an ATP-dependent manner. This is followed by the release of UvrA and formation of a stable UvrB-DNA complex, which is recognized by the endonuclease UvrC which cleaves the damaged strand at two points. UvrD helicase unwinds the damaged fragment followed by exonuclease cleavage (31, 45). XPD (ERCC2) (SF2 member with 50 to 30 directionality) is a component of the transcription factor TFIIH complex and is involved in transcription-coupled NER. Mutations in the human XPD gene result in three different disorders: xeroderma

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pigmentosum (XP), cockayne syndrome (CS), and trichothiodystrophy (TTD) (46). Recent structural data provided the basis for the different disease phenotypes produced by XPD mutations (47–49) 2.3. DNA Recombination

The E. coli RecBCD complex is involved in several processes including homologous recombination. The N-terminus of RecB and the RecD subunits have the conversed motifs of SF1 helicases. The C-terminus of RecB has the motifs of nucleases. The RecC subunit contains a region implicated in the recognition of Chi (crossover hotspot instigator), an eight base pair cis-acting DNA sequence. In addition to being one of the fastest, the RecBCD complex is unique in that it has two DNA helicases with opposite directionality, which result in a net movement in the same direction because of the anti-parallel nature of the DNA duplex. The two helicases also vary in their speed. The fast RecD helicase (50 to 30 directionality) leads the complex initially. Upon encountering Chi, the slower RecB helicase (30 to 50 directionality) becomes the leading helicase (50–52). RecQ helicases (SF2 members with 30 to 50 directionality) play important roles in maintaining genome stability including regulation of DNA recombination. These helicases participate in several DNA repair pathways through their catalytic activities and interaction with DNA repair proteins (53–55). The RecQ helicase family has been the subject of much interest as mutations in three human members result in cancer predisposition syndromes. Bloom, Werner, and Rothmund-Thomson syndromes result from mutations of the BLM, WRN, and RECQ4 helicases, respectively. BLM plays a key role in regulating homologous recombination. There is an increase in both sister and non-sister chromatid exchange in cells that lack BLM helicase. Werner syndrome is characterized by premature ageing. WRN helicase has both helicase and exonuclease activities and plays an important role in maintaining telomere stability (54, 56).

2.4. Transcription

Rho (SF5 helicase from E. coli) is a hexameric helicase required for transcription termination. Rho binds a specific region in the nascent RNA transcript which is devoid of secondary structures and rich in cytosine residues. Binding activates the RNA-dependent ATPase activity of Rho, which drives the movement of Rho hexamer along the RNA transcript in a 50 to 30 direction (31, 57). In addition to their role in RNA metabolism, some members of the DExD/H-proteins are also implicated in transcription regulation. Examples include RNA helicase A (Dhx9) (58–60) and p68 (Ddx5) (61–63). The role of some of these DExD/ H-proteins is to couple transcription to subsequent steps of RNA processing (61, 64).

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2.5. Ribosome Biogenesis

The eukaryotic ribosome consists of four ribosomal (r) RNAs and numerous ribosomal proteins. Three (28S, 18S, and 5.8S) rRNAs result from a primary transcript (pre-rRNA) produced by RNA polymerase I in the nucleolus. The fourth (5S rRNA) is transcribed by RNA polymerase III in the nucleoplasm. The pre-rRNA contains three coding regions and spacer fragments, the latter are removed by endo- and exonucleolytic processing reactions. The pre-rRNA is modified by methylation of the 20 -hydroxyl group of specific riboses and conversion of specific uridine residues to pseudouridine. The position of cleavage sites in pre-rRNA and the specific sites of 20 -O-methylation and pseudouridine formation are determined by small nucleolar RNAs (snoRNAs), which hybridize transiently to pre-rRNA molecules (65). Several DExD/H-box proteins are involved in ribosome biogenesis. Their RNA unwinding activity is required for snoRNA/ pre-rRNA dynamic interactions. They are also involved in regulating RNA–protein interactions within the RNP complexes of the ribosomes (23, 66, 67). Mutational analyses suggest that different DExD/H-box proteins have distinct functions in ribosome biogenesis (68, 69).

2.6. RNA Splicing

The spliceosome is a ribonucleoprotein complex that assembles in an ordered manner to catalyze the splicing of intervening introns during the processing of mRNAs from primary RNA transcripts. The well-studied yeast spliceosome contains five small nuclear (sn) RNAs (U1, U2, U4, U5, and U6) that interact with numerous proteins to form small nuclear ribonucleoprotein particles (snRNPs). Catalysis proceeds by two transesterification reactions. Once the exons are ligated, the mature mRNA is released for export and the spliceosome is disassembled and recycled for a new round of activity (70). NTP hydrolysis is not required for the transesterification reactions. Instead, it is has been suggested to drive the extensive spliceosomal conformational changes that take place during splicing. These conformational changes involve dsRNA unwinding and RNA–protein interactions (70). Several DExD/H box proteins are components of the splicesome, each with a role in a specific step during splicing. Prp2 (71) and Prp16 (72) are involved in the first and second catalytic reactions, respectively. Prp22 is involved in mRNA release (73), whereas Prp43 has a role during the release of lariat intron from the spliceosme (74).

2.7. Translation

The DEAD-box protein, eIF4A, is one of the earliest RNA helicases to be structuraly and biochemically characterized. EIF4A is involved in translation initiation as part of the cap-binding complex. The likely role of eIF4A is to unwind RNA secondary structure in the 50 UTR of mRNA to facilitate ribosome binding (75).

Helicases: An Overview

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Two other DEAD-box proteins have been shown to play roles in translation, including the yeast Ded1 (76, 77) and the drosophilia vasa (Ddx4) (78). 2.8. RNA Degradation

In E. coli, the DEAD-box RNA helicase RhlB is a component of the high molecular weight complex degradosome (79). In eukaryotes, the DExH-box protein Mtr4 is required for most of the nuclear activities of the exosome (80). The cytoplasmic RNA helicase, Ski2, is required for the activities of the yeast cytoplasmic exosome (81). Other helicases involved in RNA degradation include RHAU, a DExH-box protein implicated in the turnover of AU rich elements (AREs) (82), and Upf1, a key component of the nonsense-mediated mRNA decay (NMD) machinery, which is responsible for degrading mRNAs that contain premature termination codons (10, 83).

2.9. Mitochondrial Genome

Twinkle is a mitochondrial (mt) DNA helicase with 50 to 30 directionality (84), which was originally identified as the gene mutated in patients with the autosomal dominant disease progressive external ophthalmoplegia (85). The structural similarity of Twinkle to phage T7 gene 4 primase/helicase and other hexameric ring helicases is consistent with its role as the mitochondrial replicative helicase essential for mtDNA maintenance and copy number regulation (86). It has been suggested that Twinkle serves as the primase as well as the helicase for mtDNA replication in most eukaryotes with the exception of Metazoa (87). Pif1p is another mitochondrial DNA helicase with 50 to 30 directionality (SF1 member), which is involved in mtDNA repair and recombination (88–90). A nuclear isoform of Pif1p has several roles in maintaining genome stability including regulation of telomerase activity (91). There are species-dependent variations in the size and organization of the mitochondrial genome that result in differences in RNA processing requirements such as splicing and editing (92). The mammalian mitochondrial genes are arranged in a compact form with no introns, unlike the yeast. Several yeast DEAD-box proteins are involved in splicing mtRNA introns, including Cyt-19 (93), Mss116p (94), and Mrh4 (95). Another DEAD-box protein, mHel61, plays a role in editing mtRNAs in Trypanosoma brucei (96). Suv3p, a DExD/H helicase (Ski2p subfamily), is a component of the mitochondrial degradosome required for mtRNA degradation (97). Suv3p is essential for mammalian development (98) and maintenance of proper function of human mitochondria (99). Other DExH/D-box proteins have been localized to the human mitochondria including DDX28 (100), DHX30 (101), and DHX32 (102). Their exact roles in regulating the mitochondrial genome are yet to be determined.

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64. Clark E. L., Fuller-Pace F. V., Elliott D. J., and Robson C. N. (2008) Coupling transcription to RNA processing via the p68 DEAD box RNA helicase androgen receptor co-activator in prostate cancer. Biochem. Soc. Trans. 36, 546–547. 65. Granneman S. and Baserga S. J. (2004) Ribosome biogenesis: of knobs and RNA processing. Exp. Cell Res. 296, 43–50. 66. Cordin O., Banroques J., Tanner N. K., and Linder P. (2006) The DEAD-box protein family of RNA helicases. Gene 367, 17–37. 67. De La Cruz J., Kressler D., and Linder P. (1999) Unwinding RNA in Saccharomyces cerevisiae: DEAD-box proteins and related families. Trends Biochem. Sci. 24, 192–198. 68. Granneman S., Bernstein K. A., Bleichert F., and Baserga S. J. (2006) Comprehensive mutational analysis of yeast DEXD/H box RNA helicases required for small ribosomal subunit synthesis. Mol. Cell Biol. 26, 1183–1194. 69. Bernstein K. A., Granneman S., Lee A. V., Manickam S., and Baserga S. J. (2006) Comprehensive mutational analysis of yeast DEXD/H box RNA helicases involved in large ribosomal subunit biogenesis. Mol. Cell Biol. 26, 1195–1208. 70. Staley J. P. and Guthrie C. (1998) Mechanical devices of the spliceosome: motors, clocks, springs, and things. Cell 92, 315–326. 71. Kim S. H. and Lin R. J. (1996) Spliceosome activation by PRP2 ATPase prior to the first transesterification reaction of pre-mRNA splicing. Mol. Cell Biol. 16, 6810–6819. 72. Schwer B. and Guthrie C. (1991) PRP16 is an RNA-dependent ATPase that interacts transiently with the spliceosome. Nature 349, 494–499. 73. Schwer B. (2008) A conformational rearrangement in the spliceosome sets the stage for Prp22-dependent mRNA release. Mol. Cell. 30, 743–754. 74. Martin A., Schneider S., and Schwer B. (2002) Prp43 is an essential RNA-dependent ATPase required for release of lariat-intron from the spliceosome. J. Biol. Chem. 277, 17743–17750. 75. Rogers G. W., Jr., Komar A. A., and Merrick W. C. (2002) eIF4A: the godfather of the DEAD box helicases. Prog. Nucleic Acid Res. Mol. Biol. 72, 307–331. 76. Chuang R. Y., Weaver P. L., Liu Z., and Chang T. H. (1997) Requirement of the DEAD-Box protein ded1p for messenger RNA translation. Science 275, 1468–1471.

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86. Tyynismaa H., Sembongi H., BokoriBrown M., Granycome C., Ashley N., Poulton J., Jalanko A., Spelbrink J. N., Holt I. J., and Suomalainen A. (2004) Twinkle helicase is essential for mtDNA maintenance and regulates mtDNA copy number. Hum. Mol. Genet. 13, 3219–3227. 87. Shutt T. E. and Gray M. W. (2006) Twinkle, the mitochondrial replicative DNA helicase, is widespread in the eukaryotic radiation and may also be the mitochondrial DNA primase in most eukaryotes. J. Mol. Evol. 62, 588–599. 88. Cheng X., Dunaway S., and Ivessa A. S. (2007) The role of Pif1p, a DNA helicase in Saccharomyces cerevisiae, in maintaining mitochondrial DNA. Mitochondrion 7, 211–222. 89. Pinter S. F., Aubert S. D., and Zakian V. A. (2008) The Schizosaccharomyces pombe Pfh1p DNA helicase is essential for the maintenance of nuclear and mitochondrial DNA. Mol. Cell Biol. 28, 6594–6608. 90. Foury F. and Kolodynski J. (1983) pif mutation blocks recombination between mitochondrial rho+ and rho– genomes having tandemly arrayed repeat units in Saccharomyces cerevisiae. Proc. Natl. Acad. Sci. USA 80, 5345–5349. 91. Boule J. B. and Zakian V. A. (2006) Roles of Pif1-like helicases in the maintenance of genomic stability. Nucleic Acids Res. 34, 4147–4153. 92. Gagliardi D., Stepien P. P., Temperley R. J., Lightowlers R. N., and ChrzanowskaLightowlers Z. M. (2004) Messenger RNA stability in mitochondria: different means to an end. Trends Genet. 20, 260–267. 93. Mohr S., Stryker J. M., and Lambowitz A. M. (2002) A DEAD-box protein functions as an ATP-dependent RNA chaperone in group I intron splicing. Cell 109, 769–779. 94. Huang H. R., Rowe C. E., Mohr S., Jiang Y., Lambowitz A. M., and Perlman P. S. (2005) The splicing of yeast mitochondrial group I and group II introns requires a DEAD-box protein with RNA chaperone function. Proc. Natl. Acad. Sci. USA 102, 163–168. 95. Schmidt U., Lehmann K., and Stahl U. (2002) A novel mitochondrial DEAD box protein (Mrh4) required for maintenance of mtDNA in Saccharomyces cerevisiae. FEMS Yeast Res. 2, 267–276. 96. Missel A., Souza A. E., Norskau G., and Goringer H. U. (1997) Disruption of a gene encoding a novel mitochondrial

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DEAD-box protein in Trypanosoma brucei affects edited mRNAs. Mol. Cell Biol. 17, 4895–4903. 97. Margossian S. P., Li H., Zassenhaus H. P., and Butow R. A. (1996) The DExH box protein Suv3p is a component of a yeast mitochondrial 30 -to-50 exoribonuclease that suppresses group I intron toxicity. Cell 84, 199–209. 98. Pereira M., Mason P., Szczesny R. J., Maddukuri L., Dziwura S., Jedrzejczak R., Paul E., Wojcik A., Dybczynska L., Tudek B., Bartnik E., Klysik J., Bohr V. A., and Stepien P. P. (2007) Interaction of human SUV3 RNA/DNA helicase with BLM helicase; loss of the SUV3 gene results in mouse embryonic lethality. Mech. Ageing Dev. 128, 609–617. 99. Khidr L., Wu G., Davila A., Procaccio V., Wallace D., and Lee W. H. (2008) Role of

SUV3 helicase in maintaining mitochondrial homeostasis in human cells. J. Biol. Chem. 283, 27064–27073. 100. Valgardsdottir R., Brede G., Eide L. G., Frengen E., and Prydz H. (2001) Cloning and characterization of MDDX28, a putative dead-box helicase with mitochondrial and nuclear localization. J. Biol. Chem. 276, 32056–32063. 101. Wang Y. and Bogenhagen D. F. (2006) Human mitochondrial DNA nucleoids are linked to protein folding machinery and metabolic enzymes at the mitochondrial inner membrane. J. Biol. Chem. 281, 25791–25802. 102. Alli Z., Ackerley C., Chen Y., Al-Saud B., and Abdelhaleem M. (2006) Nuclear and mitochondrial localization of the putative RNA helicase DHX32. Exp. Mol. Pathol. 81, 245–248.

Chapter 2 Fluorescent Biosensors to Investigate Helicase Activity Martin R. Webb Abstract ATP-driven translocation of helicases along DNA can be assayed in several ways. Reagentless biosensors, based on fluorophore–protein adducts, provide convenient ways for real-time assays of both the separation of dsDNA and the hydrolysis of ATP. Single-stranded DNA can be assayed using a modified singlestranded DNA-binding protein (SSB), and phosphate production during ATP hydrolysis can be measured by a modified phosphate-binding protein. Advantages and limitations of these approaches are compared with those of other types of measurements. Key words: Helicase, assay, fluorescence, kinetics, ATPase, phosphate.

1. Introduction Different helicases translocate along DNA with a wide range of rates, typically 10–1000 bases per second and may move for short distances of a few bases (low processivity) or up to several thousand bases depending on the kinetics of translocation versus dissociation (1). These kinetics are presumably tuned to the biological role of the helicase. They generally use the energy of ATP hydrolysis to drive this movement and separation of duplex DNA. To study these processes in real time, methods are required that have sufficient sensitivity and time resolution to quantify strand separation and ATP hydrolysis typically on the time scale of milliseconds to seconds. Fluorescence probes often have high sensitivity and rapid response and various types of fluorescence instrumentation are available so that the signals can be measured rapidly and continuously. This chapter describes the use of fluorescent biosensors that have been developed for these types of applications and compares M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_2, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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them with other types of measurements that may give similar or complementary information. On the whole, the chapter is limited to bulk measurements in solution and methods that are based on fluorescence. 1.1. Translocation Assays

Potentially standard gel-based assays might be used to measure duplex separation catalyzed by helicases, but with difficulty to get the time resolution required for real-time measurements. In any case, by their discontinuous nature, such measurements are likely to give single time points after the reaction is quenched. Fluorescence probes have proved useful for continuous measurement of the conversion of double-stranded DNA (dsDNA) to singlestranded DNA (ssDNA), although they have the potential for disrupting the natural system being studied. Protein–DNA interactions may be modified by the presence of a dye moiety. Other molecules, added as probes, may bind to the helicase or DNA and so modify the translocation. For translocation along short lengths of DNA, several strategies have been used with fluorescent labels attached to oligonucleotides, to give a signal to monitor translocation with single-base resolution (2–7). If the end of the DNA is labeled, then no signal change is observed until the helicase approaches near to the fluorescent label. The signal change then occurs when the helicase reaches the end label. Even if the fluorescent label interferes with the translocation in some way, the time elapsed before the signal change provides a measure of the time taken to translocate the length of DNA. This is sometimes called an ‘‘all-or-none’’ measurement. Ideally this measurement should be done with different lengths of the DNA track. There should be a linear dependence of the time taken on the length and this allows the translocation speed to be determined. Use of several lengths enables at least partial elimination of end effects in the translocation, such as a lag before movement starts or a fluorescence change due to environmental effects when the helicase complexes are close to the fluorophore. Several laboratories have used fluorescence resonance energy transfer (FRET) (for example, (4)) or a fluorophore– quencher pair, such as Cy3 and Dabcyl groups (for example, (8)). These are at the same end of a length of dsDNA so that they can interact fully and have little or no fluorescence while the duplex is intact. Use of a quencher in the substrate DNA has the advantage of giving a fluorescence increase during the helicase assay, a property that is useful for measuring small extents of reaction and limiting interference by photobleaching. However, these types of approach are limited to < 100 base lengths for bulk measurements: desynchronization of helicases tends to produce smaller signals as the length increases. An advantage is that measurements of short lengths of translocation with end-labeling fluorophores can give single-base resolution in both bulk and

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single-molecule assays. A precise and powerful analysis for these types of all-or-none assays has been described based on kinetic modeling (9). For translocation over long lengths of DNA (hundreds to thousands of bases), different approaches are required. Strand separation can be measured using probes that discriminate between ssDNA and dsDNA. A number of dyes bind tightly to dsDNA, but weakly, or not at all, to ssDNA and give a fluorescence change, usually an increase, on binding (10, 11). If such an interchelating dye is bound along dsDNA, the helicase will displace the dye molecules as it translocates: in bulk solution assays, there is a continuous, gradual increase in fluorescence that gives a measure of the extent of translocation. However, the presence of the dye may disrupt the helicase action or even, in extreme cases, prevent translocation. In addition, this assay has the disadvantage of producing a decrease in fluorescence, which means that the measurement may be made against a high background and possibly against a simultaneous decrease due to photobleaching. Triplex displacement assays show promise for translocation assays. Here, particular sections of duplex DNA sequence bind a third length of DNA. Helicases can potentially translocate through this triplex section and in doing so release the third DNA molecule. If a fluorescently labeled oligonucleotide is used for this, it can give a change in fluorescence on release, which provides an ‘‘all-or-none’’ measure of translocation time to that point (12, 13). As for end labeling of oligonucleotides, measurements of translocation times for different lengths is advantageous, although, in practice, such end effects are likely to be a much smaller fraction of the total translocation time when long lengths of dsDNA are used. An approach that measures the product ssDNA formation has advantages over ones that measure substrate depletion. Singlestranded DNA-binding protein (SSB) has been used to achieve this, either using the intrinsic tryptophan fluorescence (14) or by use of an extrinsic fluorescence label attached to SSB (15). SSB from Escherichia coli exists as a tetramer and binds a length of up to 70 nucleotides of ssDNA (16, 17). Under some circumstances there may be a different binding mode, whereby approximately half that length binds to the tetramer. This ‘‘35 base’’ binding mode is favored by low ionic strength and high ratio of SSB to DNA. An internal tryptophan has provided a continuous signal for assaying helicase unwinding of duplex DNA. As DNA binds to the SSB, there is a decrease in tryptophan fluorescence. Because of the relatively low-intensity decrease and the potential for other components of the assay solution to interfere with the fluorescence, this assay has relatively low sensitivity. To circumvent some of these problems, a single cysteine mutant of SSB was labeled with a coumarin fluorophore so that it gives a sixfold increase in fluorescence on binding ssDNA (15). The aim was to produce a probe

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for helicase assays with high sensitivity and time resolution. Although binding is complex (18) with multiple modes as described above and potentially cooperative effects as multiple SSB tetramers bind along ssDNA, this biosensor gives an approximately linear response to the amount of ssDNA. The fluorescence response is also linear during measurement of dsDNA unwinding by a helicase. Examples of such assays have been published that demonstrate the high sensitivity of the coumarin-labeled SSB (DCC-SSB) (15). This chapter describes the use of this biosensor to measure helicase-driven separation of dsDNA. 1.2. ATPase Activity Assays

There are several different ways to measure the ATPase activity of a helicase, generally by monitoring one of the products, ADP and inorganic phosphate (Pi). Relative advantages of the various assays depend in part on whether a steady-state measurement is required, which is likely to be over a longer time course, or whether ATP hydrolysis is measured in real time to correlate directly with translocation. Use of radiolabeling provides a relatively simple assay, which requires no additional components to be present. However, such measurements are discontinuous and each assay point requires separation of ATP hydrolysis products. There are a number of coupled-enzyme assays that measure product ADP (for example, (19, 20)) or inorganic phosphate, Pi (for example, (21–23)). Some can provide a fluorescence signal or often are based on an absorbance change. The latter sacrifices sensitivity, but has the advantage in that the signal response is linear and readily quantified, based on an extinction coefficient (e.g., of NADH). Fluorescence assays usually have to be calibrated for each different set of conditions. However, coupled-enzyme assays inevitably required addition of multiple components and there needs to be care to ensure that these additions do not affect the assay and that the observed rate is that of the helicase, not of the coupled enzyme. A typical pitfall is to assume that the coupled enzymes are operating at maximal velocity, although pH and buffer conditions are, in practice, producing a suboptimal rate. While these types of assays may be very useful for steady-state measurements, they may not be fast enough for the high activity of helicases during single-turnover measurements with respect to DNA. For the latter, fluorescence-based reagentless biosensors may have advantages, including the need for only one added component (the biosensor molecule). There is, therefore, only one coupled rate that needs to be checked and compared with that of the helicase. There have been several such biosensors described for ADP (for example, (24, 25)) and Pi (for example, (26, 27)), although some are aimed particularly at high-throughput assays and may not have a high rate of response. This chapter describes the use of the biosensors based on the phosphate-binding protein. This type has been widely used in assays of ATPase activity and mechanism of helicase (28) and

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other motor proteins such as myosins (29) and kinesins (30). The method is particularly suited to stopped-flow measurements, for example, to measure single-turnover kinetics, the details of the ATPase cycle, or the precise relationship between translocation and the number of ATP molecules hydrolyzed. To produce the original phosphate biosensor, a single cysteine mutant of the bacterial phosphate-binding protein was labeled covalently with a single coumarin to give a signal increase (approximately an order of magnitude) on Pi binding. More recently a version of this with two rhodamines was developed with greater sensitivity and photostability (27). The bases for these fluorescence changes, the kinetic mechanism of binding Pi to this protein, have been described (31, 32). The binding is tight (50 nM) so that for most conditions, the biosensor must be in excess over the maximum amount of Pi likely to be encountered in the assay or at least during the time period of the measurement. Because of this and the ubiquity of Pi contamination, it is important to assess and minimize any such contamination. This contamination has been discussed and an enzymic method of reducing it has been described (33, 34). The ‘‘phosphate mop’’ consists of purine nucleoside phosphorylase and 7-methylguanosine, which converts Pi to ribose-1-phosphate and so to a chemical species that is silent with respect to the phosphate biosensor. 1.3. Described Methods

The methods described include examples of applications of these biosensors to measure the enzymic activity of helicases, either in the steady state or as rapid reaction measurements using stoppedflow. Some general points about experimental design are given below. 1.Check the labeled protein. The quality of the fluorescent protein can be simply checked with a titration of the ligand, measuring fluorescence. Examples are given in Section 3. The response should be linear and give fluorescence increases similar to those published. In the case of the Pi sensors, this can also assess the Pi contamination in the preparation and buffers. Addition of Pi mop components rather than Pi will give a small decrease in fluorescence, whose size depends on the degree of contamination. The breakpoint of the titration reflects the concentration of active biosensor. 2.Optical considerations. These are mainly outside the scope of this chapter, but it should be realized that optics plays a large part in determining the quality of the signal obtained. Thus the coumarin-based sensors described here are particularly well suited to excitation via a strong Hg line at 436 nm. There is an Hg line at 546 nm that can be used for rhodamine excitation. Where there is a choice between a xenon lamp and a xenon–mercury lamp, for example, with a stopped-flow instrument, much higher excitation intensity can be obtained with the latter.

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The possibility of inner filter effects must also be considered: is the absorbance of the fluorophore significant (say, > 0.1 for the pathlength being used)? Changing the excitation wavelength may be possible. For the coumarin-based sensors the extinction coefficient does not change between apo and bound forms, so to some extent inner filter effects will cause the fluorescence change to decrease, but remain linear. The calibration can be altered accordingly. Note that the rhodamine extinction coefficient does change, so the interpretation may be complex, but in this case, there may be significant scope to change the wavelength. 3.Rate of response. Depending on the type of measurement, the rate of response may also be a significant consideration: this typically depends on the concentration of the binding protein (the biosensor), since the concentration of free ssDNA or Pi is likely to be very low during the assay. The aim is to ligate the product rapidly with the biosensor. The publications describing these sensors have binding measurements for some conditions that are a good guide. Nevertheless significant differences in conditions can lead to large changes in rate constants.

2. Materials 2.1. Assay Components

1. Fluorescent reagents are available as follows: MDCC (N-[2(1-maleimidyl)ethyl]-7-diethylaminocoumarin-3-carboxamide) and IDCC (N-[2-(iodoacetamido)ethyl]-7-diethylaminocoumarin-3-carboxamide) (Invitrogen, USA, or Synchem, Germany); 6-IATR (6-iodoacetamidotetramethylrhodamine) (Chemos, Germany). 2. SSB (G26C): prepared as previously described (15). 3. MDCC-PBP: prepared as previously described (26, 31, 34) (Invitrogen). Rhodamine-PBP was prepared as described and stored as a concentrated solution (1 mM) in small aliquots at –80C (27). 4. Components for the phosphate mop (lyophilized, ‘‘bacterial’’ purine nucleoside phosphorylase and 7-methylguanosine) (Sigma-Aldrich). The additional ‘‘supermop’’ components were glucose-1,6-bisphosphate and manganese chloride (Sigma-Aldrich) and phosphodeoxyribomutase from E. coli, prepared as previously described (33) (see Note 1).

2.2. Other Proteins and Plasmids

1. Wild-type phosphate-binding protein from E. coli was prepared as described (26). This is the phosphate complex. Phosphate was partially removed by treatment with the ‘‘supermop’’ components. A solution (500 ml) in 10 mM PIPES, pH 7.0, of 100 mM protein, 200 mM

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7-methylguanosine, 0.2 unit ml–1 purine nucleoside phosphorylase, 5 mM MnCl2, 1 mM glucose-1,6-bisphosphate, and 1.5 mg ml–1 phosphodeoxyribomutase was incubated at 20C for 15 min. It was desalted on a PD10 column (GE Heathcare) and pre-equilibrated in the same buffer. This resulted in a solution of 0.5 ml of 80 mM phosphate-binding protein from the peak fraction (see Note 2). 2. RepD from Staphylococcus aureus and PcrA from Bacillus stearothermophilus were prepared as described (35, 36). For steady-state assays, as described, use a fresh 5-nM stock containing 5 mM BSA as carrier. 3. Plasmid pCERoriD, containing the oriD sequence, was prepared as described (37). 2.3. Other Biochemicals

1. 100 mM sodium 2-mercaptoethane-sulfate (MESNA): prepare fresh. 2. 1 M dithiothreitol (DTT). 3. Phosphate (Pi) standard solution (VWR, Aristar, 1000 ppm as ‘‘P,’’ which refers to PO43–). 4. ATP (Sigma) (SigmaUltra grade), which has low Pi contamination (see Note 3). 5. dT20 and dT70 (Eurogentec) and HPLC-purified grade.

2.4. Buffers and Solutions

1. Buffer for DCC-SSB labeling: 20 mM Tris–HCl, pH 7.5, 1.0 mM EDTA, 500 mM NaCl, 20% (v/v) glycerol. 2. Buffer for DCC-SSB purification: 20 mM Tris–HCl, pH 8.3, 1.0 mM EDTA, 500 mM NaCl, 20% (v/v) glycerol. 3. Buffers for testing pH and ionic strength variations in DCCSSB signal: 25 mM Tris–HCl, pH 8.0, or 25 mM PIPES, pH 7.0, each containing 20 or 200 mM NaCl. 4. Buffer for helicase activity assay: 50 mM Tris–HCl, pH 7.5, 200 mM KCl, 10 mM MgCl2, 1 mM EDTA, and 10% ethanediol. 5. Buffer for rhodamine-PBP characterization: 10 mM PIPES, pH 7.0. 6. Buffer for steady-state ATPase assay: 50 mM Tris–HCl, pH 7.5, 150 mM NaCl, 3 mM MgCl2.

3. Methods 3.1. Label SSB (G26C) with IDCC

1. Add 5 mmol DTT to 10 mg SSB (530 nmol) and incubate for 20 min at room temperature to ensure all cysteines are fully reduced.

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2. Degas the labeling buffer just before use, by bubbling nitrogen through for 5 min. 3. Equilibrate a PD10 column (GE Healthcare) in the degassed buffer: pass 25 ml through. 4. Pass the DTT-treated SSB through the PD10 column, eluting with degassed buffer. Collect 15  0.5 ml fractions and measure their absorbance at 280 nm (dilute a portion  20 and use the buffer as blank). Collect protein fraction(s). 5. In a sealable tube, make the protein 100 mM by diluting, if necessary, with labeling buffer and add 200 mM IDCC. 6. Play nitrogen over the solution just before sealing the tube. 7. Incubate 2 h at 22C with end-over-end stirring, protected from light. 8. Add 1 mM MESNA and leave for 30 min with end-over-end stirring, protected from light, to react with remaining coumarin. 9. Pass the solution through a membrane filter (0.2 mm pores, polyethersulfone from Whatman). 10. Pre-equilibrate a P4 gel filtration column (BioRad; 1  20 cm) in the DCC-SSB purification buffer. 11. Pass the labeled protein through this column, collecting 0.5 ml fractions and pooling the first colored peak. 12. Concentrate the protein to 50–100 mM monomers using a Centricon YM10 membrane concentrator. 13. Measure the absorbance spectrum and calculate the protein concentration at 430 nm, where the extinction coefficient of the coumarin is 44,800 M–1 cm–1. 14. Store the protein at –80C after quick freezing.

3.2. Check the Labeled Protein – Calibration of DCC-SSB

1. This section describes calibrating DCC-SSB at different pH and ionic strength conditions (see Note 4). 2. Place 100 ml of 200 nM DCC-SSB tetramers in the appropriate buffer in a 3  3 mm fluorescence cuvette. 3. Measure fluorescence at 20C in a Cary Eclipse fluorimeter with excitation at 430 nm and emission at 470 nm (see Note 5). 4. Titrate in aliquots of dT70 over the range to 50 nM, as shown in Fig. 2.1. Correct the data for any dilution. 5. Finally, add a twofold molar excess of the oligonucleotide to obtain an end point; in the examples shown, this gives the 100% level for the signal (see Note 6).

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Fig. 2.1. Calibration of DCC-SSB with DNA at different pH and ionic strength conditions. These data are a single set of measurements. Measurements were done at 25 mM Tris, pH 8.0, 200 mM NaCl (circles); 25 mM Tris–HCl, 20 mM NaCl (diamonds); 25 mM PIPES, pH 7.0, 200 mM NaCl (triangles); 25 mM PIPES, pH 7.0, 20 mM NaCl (squares). All solutions were at 20C and contained 250 nM DCC-SSB (tetramer concentration) and 5 mM BSA as carrier. Fluorimeter settings are as described in the text. One hundred percent fluorescence represents the signal with excess DNA as described in the text.

3.3. Check the Labeled Protein – Titration of Rhodamine-PBP with Phosphate

1. Place a solution (200 ml in 10 mM PIPES, pH 7.0) of 2.5 mM rhodamine-PBP in a 3  3 mm fluorescence cuvette. 2. Measure fluorescence at 20C in a Cary Eclipse fluorimeter with excitation at 555 nm and emission at 578 nm (see Note 5). 3. Add aliquots of Pi standard, suitably diluted, to produce a titration curve as in Fig. 2.2. Correct the data for any dilution (see Note 7). 4. The intercept of the linear fits to the fluorescence rise and to the horizontal portion gives a measure of the active, Pi-free protein.

Fig. 2.2. Titration of rhodamine-PBP with inorganic phosphate. The upper plot (circles) is for rhodamine-PBP alone. The lines are linear fits to the rise and horizontal portions. The lower plot (triangles) is offset by –10% for clarity and is for equimolar mixture of wildtype PBP and rhodamine-PBP. The wild-type protein was treated with phosphate mop as described in the text to remove bound Pi partially. The curvature shows that wild-type protein binds Pi more tightly than the labeled protein. The line is a best fit to a model, in which there is tight binding, as previously described (15): the ratio of dissociation constants is 4 based on this, with the wild type binding Pi tighter.

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5. To demonstrate the effect of unlabeled phosphate-binding protein, repeat the titration in the presence of 2.5 mM wildtype protein, after partial removal of bound Pi by treatment with the phosphate ‘‘supermop’’ (see Note 8). 3.4. Real-Time Helicase Activity Assay Using Coumarin-Labeled SSB (DCC-SSB)

1. Prepare 1 ml of a solution in the helicase assay buffer: 2 nM pCERoriD circular plasmid, pre-incubated for 10 min with 4 nM RepD, 400 nM tetramer DCC-SSB, 50 nM PcrA (Note 9). 2. Prepare 1 ml of a 2-mM ATP solution in the helicase assay buffer. 3. After equilibration at 37C, mix these two solutions rapidly, using a stopped-flow apparatus (Hi-Tech, TgK Scientific, UK), equipped with xenon–mercury lamp. Follow the fluorescence with time, exciting at 436 nm and using a 455-nm cut-off filter on the emission. A sample trace is shown in Fig. 2.3 (see Note 10). 4. An identical solution, but in the absence of RepD, can be used as a zero activity control. PcrA has negligible helicase activity under these conditions. This provides a check on the stability of the fluorescent sensor for the experimental (solution and optical) conditions (see Note 11).

Fig. 2.3. Real-time helicase assay using fluorescent SSB. The assay is for unwinding a 3094-bp pCERoriD plasmid by PcrA/RepD at 37C in the presence of DCC-SSB, with details described in the text. The fluorescence of the DCCSSB was followed with time. The lower line is a control in the absence of RepD, for which there should be little or no strand separation.

3.5. Steady-State ATPase Activity Assay Using Rhodamine-PBP

1. Prepare 1 ml of assay solution in the ATPase assay buffer: 0.5 mM dT20, 20 mM ATP, 5 mM rhodamine-PBP (see Note 12). 2. Prepare a 5-nM solution PcrA in ATPase activity buffer, containing 5 mM BSA as carrier. 3. Place 200 ml assay solution in a 3  3 mm fluorescence cuvette and equilibrate to 20C in a fluorimeter (Cary Eclipse – see Note 5).

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4. Follow fluorescence with time, exciting at 555 nm and measuring emission at 578 nm. 5. When the fluorescence is constant, add PcrA to give a concentration of 50 pM. Following a period (200 s) of constant increase in fluorescence, add a second aliquot of PcrA to give a total of 100 pM to check that the rate doubles. A sample trace is shown in Fig. 2.4. 6. Repeat the fluorescence measurement, but obtain a control in the absence of PcrA (see Note 11). 7. Do linear fits of each section of the time course and the control, to obtain the rate of the reaction at each enzyme concentration. 8. Calibrate the fluorescence signal, using a further 200 ml aliquot of the assay solution, but add 2  0.5 mM aliquots of Pi standard, measuring the fluorescence change on each addition. This should be repeated with a fresh aliquot of assay solution. Average the four
fluorescence values to convert the (arbitrary) fluorescence scale to nanomolar phosphate.

Fig. 2.4. Steady-state ATPase assay of PcrA with dT20, measured using the rhodaminePBP phosphate sensor. The incubation solution at 20C contained the following: 450 mM Tris–HCl, pH 7.5, 150 mM NaCl, 3 mM MgCl2, 20 mM ATP, 0.5 mM dT20, and 5 mM rhodamine-PBP. The reaction was initiated at zero time by addition of 50 pM PcrA from a 5-nM stock containing 5 mM BSA as carrier. A similar addition at 260 s illustrates the doubling of rate. A control with no PcrA is shown to check the stability of the fluorescence signal.

4. Notes

1. Store the 7-methylguanosine as a 20 mM solution in water at –20C. Store the phosphorylase as 1000 unit ml–1 in the lyophilized buffer in small aliquots at –80C: repeated freezing and thawing deactivates this enzyme. It is important that the protein solutions are snap frozen. Some other types of this

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phosphorylase are available commercially but may not be suitable: either they do not accept 7-methylguanosine as a substrate with high activity or they come with phosphate buffer. 2. Because wild-type phosphate-binding protein binds Pi much tighter than the labeled protein, removal of Pi needs the extra components of glucose-1,6-bisphosphate, MnCl2, and phosphodeoxyribomutase. Normal mopping of protein solutions, if required at all, requires a similar treatment but only with 7-methylguanosine and purine nucleoside phosphorylase. Labeled phosphate-binding protein should have < 10% Pi as prepared. 3. To prevent hydrolysis during storage, keep the solid nucleotide desiccated (over Drierite) in a sealed container at –20C. To avoid condensation, warm to room temperature before opening. Concentrated stock solutions can be stored at –80C with minimal decomposition if the pH is adjusted to 4–7. Avoid many repeats of thawing and freezing. 4. As in almost all cases with fluorescence measurements, the signal must be calibrated at the conditions and concentrations being used. To obtain a relatively linear response, generally for these types of biosensors, there needs to be a compromise with signal sensitivity. Usually the response is closest to linear at low extents of saturation. In the case of SSB, the mode of binding, as discussed above, as well as general factors, such as temperature and ionic strength, affects the fluorescence response. For measuring ssDNA, therefore, we proposed a rule of thumb that the ratio of SSB subunit to nucleotides of ssDNA is 5, based on the maximum ssDNA to be measured (15). This keeps the response fairly linear for a variety of conditions. This method illustrates the variation in fluorescence as pH and ionic strength vary. However, for each condition the response is approximately linear. It is important to test the quality of the labeled protein, using the ‘‘standard’’ buffer, but do a calibration with the conditions that are being used in the helicase assay. 5. The fluorimeter is a standard instrument, equipped with xenon lamp, and any equivalent one would be appropriate. Because any fluorescence intensity can vary with temperature, it is important to have temperature control. In addition, fluorescence measurements may be instrument dependent so that calibrations should be done, if possible, on the same instrument as the actual assay. 6. In the example shown, the 100% signal was, in absolute terms, very similar for the four conditions (within 10%). The main variation is in the starting fluorescence (free DCC-SSB) and whether there is significant ‘‘35-base’’ binding mode at low ionic strength, as described in the text above.

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7. The rhodamine-PBP has a linear response until almost saturated (7). However, part of the capacity of the PBP may be taken (‘‘used up’’) by contaminating Pi. This must be dealt with in subsequent assays, either by minimizing contamination or by having extra PBP present or by having ‘‘Pi mop’’ present at low enough activity not to affect the measurement. In general, the measurement should be tried first in the absence of mop. Ways of dealing with phosphate contamination have been discussed (34). 8. For each sensor, the labeling does cause a decrease in affinity for the ligand and so at low concentrations of ligand, any unlabeled protein will preferentially bind the ligand. So the fluorescence change is smaller at this stage of the titration. This is illustrated in Fig. 2.2, where a mixture of labeled and unlabeled proteins is used. The same effect will be observed in the titration of labeled protein alone: lack of linearity of the fluorescence rise may reflect incomplete labeling. Mass spectrometry of the protein would act as a check on this. 9. The procedure described for a single turnover of dsDNA strand separation can readily be adapted for other helicases, either using a similar concentration of DNA in terms of bases or by modifying the concentration of DCC-SSB sensor appropriately. The buffer appropriate for the helicase can be used, but with appropriate calibration of the fluorescence signal, as described. Slower helicases or multi-turnover assays can be done in a cuvette using a standard fluorimeter. 10. Several suppliers produce stopped-flow equipment suitable for these types of assay. The technical aspects of stoppedflow fluorimetry have been described in several reviews (for example, (38)), as has the additional information that can be obtained from single-turnover measurements as opposed to steady-state ones (39). 11. The control fluorescence measurements in the absence of enzyme activity are important to test the signal stability under the same experimental conditions and the same timescale as the assay itself. The fluorescence signal may change due to protein stability, solution cloudiness (light scatter), or photobleaching, for example. Note that photobleaching of the Pi-free rhodamine-PBP results in an increase in fluorescence: as one of the stacked (and hence quenched) rhodamine pair photobleaches, the other then exhibits full monomer fluorescence (27). 12. This illustrates a sensitive assay for ATPase activity that can be adapted to other enzymes with appropriate solution and temperature conditions. In any case, it is important to use ATP solutions that are low in phosphate contamination.

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Acknowledgments This work was supported by the Medical Research Council, UK. I would like to thank all those who collaborated on developing and implementing these biosensors at NIMR or elsewhere and are coauthors in the referenced papers. References 1. Singleton M.R., Dillingham M.S., and Wigley D.B.(2007) Structure and mechanism of helicases and nucleic acid translocases.Annu. Rev. Biochem. 76, 23–50. 2. Dillingham M. S., Wigley D. B., Webb M. R. (2002) Direct measurement of single stranded DNA translocation by PcrA helicase using the fluorescent base analogue 2aminopurine.Biochemistry 41, 643–651. 3. Ha T., Rasnik I., Cheng W., Babcock H. P., Gauss G. H., Lohman T. M., et al. (2002) Initiation and re-initiation of DNA unwinding by the Escherichia coli Rep helicase.Nature 419, 638–641. 4. Bjornson K. P., Amaratunga M., Moore K. J., Lohman T. M. (1994) Single-turnover kinetics of helicase-catalyzed DNA unwinding monitored continuously by fluorescence energy transfer. Biochemistry 33, 14306–14316. 5. Myong S., Bruno M. M., Pyle A. M.,and Ha T. (2007) Spring-loaded mechanism of DNA unwinding by hepatitis C virus NS3 helicase. Science 31, 7513–516. 6. Martinez-Senac M. M. and Webb M. R. (2005) Mechanism of translocation and kinetics of DNA unwinding by the helicase RecG. Biochemistry 44, 16967–16976. 7. Lucius A. L., Wong C. J., and Lohman T. M. (2004) Fluorescence stopped-flow studies of single turnover kinetics of E. coli RecBCD helicase-catalyzed DNA unwinding.J. Mol. Biol. 339, 731–750. 8. Boguszewska-Chachulska A. M., Krawczyk M., Stankiewicz A., Gozdek A., Haenni A., and Strokovskaya L. (2004) Direct fluorometric measurement of hepatitis C virus helicase activity.FEBS Lett. 56, 7253–258. 9. Lucius A. L., Maluf N. K., Fischer C. J., and Lohman T. M. (2003) General methods for analysis of sequential ‘‘n-step’’ kinetic mechanisms: application to single turnover kinetics of helicase-catalyzed DNA unwinding. Biophys. J. 85, 2224–2239. 10. Eggleston A. K., Rahim N. A., and Kowalczykowski S. C. (1996) A helicase assay based

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on the displacement of fluorescent, nucleic acid-binding ligands.Nucleic Acids Res. 24, 1179–1186. Rye H. S., Quesada M. A., Peck K., Mathies R. A., and Giazer A. N. (1991) High-sensitivity two-color detection of double-stranded DNA with a confocal fluorescence gel scanner using ethidium homodimer and thiazole orange. Nucleic Acids Res. 19, 327–333. McClelland S. E., Dryden D. T. F., and Szczelkun M. D. (2005) Continuous assays forDNA translocationusingfluorescenttriplex dissociation: application to Type I restriction endonucleases. J. Mol. Biol. 348, 895–915. Firman K. and Szczelkun M. D. (2000) Measuring motion on DNA by the type I restriction endonuclease EcoR124I using triplex displacement. EMBO J. 19, 2094–2102. Roman L. J. and Kowalczykowski S. C. (1989) Characterization of the helicase activity of the Escherichia coli RecBCD enzyme using a novel helicase assay. Biochemistry 28, 2863–2873. Dillingham M. S., Tibbles K. L., Hunter J. L., Bell J. C., Kowalczykowski S. C., and Webb M. R. (2008) A fluorescent singlestranded DNA binding protein as a probe for sensitive, real time assays of helicase activity. Biophys. J. 95, 3330–3339. Lohman T. M. and Ferrari M. E. (1994) Escherichia Coli single-stranded DNA-binding protein: multiple DNA-binding modes and cooperativities. Annu. Rev. Biochem. 63, 527–570. Raghunathan S., Kozlov A. G., Lohman T. M., and Waksman G. (2000) Structure of the DNA binding domain of E. coli SSB bound to ssDNA.Nat. Struct. Biol. 7, 648–652. Kuznetsov S. V., Kozlov A. G., Lohman T. M., and Ansari A. (2006) Microsecond dynamics of protein-DNA interactions: direct observation of the wrapping/unwrapping kinetics of single-stranded DNA around the E. coli SSB tetramer. J. Mol. Biol. 359, 55–65.

Fluorescent Biosensors to Investigate Helicase Activity 19. Charter N. W., Kauffman L., Singh R., and Eglen R. M. (2006) A generic, homogenous method for measuring kinase and inhibitor activity via adenosine 5’-diphosphate accumulation. J. Biomol. Screen 11, 390–399. 20. Lowry O. H. and Passonneau J. V. (1972) A flexible system of enzymatic analysis. New York Academic Press. 21. Webb M. R. (1992) A continuous spectrophotometric assay for inorganic phosphate and for measuring phosphate release kinetics in biological systems. Proc. Natl. Acad. Sci. USA 89, 4884–4887. 22. Banik U. and Roy S. (1990) A continuous fluorimetric assay for ATPase activity. Biochem. J. 266, 611–614. 23. De Groot H. and Noll T. (1985) Enzymic determination of inorganic phosphates, organic phosphates and phosphate-liberating enzymes by use of nucleoside phosphorylase-xanthine oxidase (dehydrogenase)coupled reactions. Biochem. J. 229, 255–260. 24. Srinivasan J., Cload S. T., Hamaguchi N., Kurz J., Keene S., Kurz M., et al. (2004) ADP-specific sensors enable universal assay of protein kinase activity. Chem. Biol. 11, 499–508. 25. Brune M., Corrie J. E. T., and Webb M. R. (2001) A fluorescent sensor of the phosphorylation state of nucleoside diphosphate kinase and its use to monitor nucleoside diphosphate concentrations in real time. Biochemistry 40, 5087–5094. 26. Brune M., Hunter J. L., Corrie J. E. T., Webb M. R. (1994) Direct, real-time measurement of rapid inorganic phosphate release using a novel fluorescent probe and its application to actomyosin subfragment 1 ATPase. Biochemistry 33, 8262–8271. 27. Okoh M. P., Hunter J. L., Corrie J. E. T., and Webb M. R. (2006) A biosensor for inorganic phosphate using a rhodaminelabeled phosphate binding protein. Biochemistry 45, 14764–14771. 28. Dillingham M. S., Wigley D. B., and Webb M. R. (2000) Demonstration of unidirectional single-stranded DNA translocation by PcrA helicase: measurement of step size and translocation speed. Biochemistry 39, 205–212. 29. White H. D., Belknap B., and Webb M. R. (1997) Kinetics of nucleoside triphosphate cleavage and phosphate release steps by associated rabbit skeletal actomyosin, measured using a novel fluorescent probe for phosphate. Biochemistry 36, 11828–11836.

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30. Gilbert S. P., Webb M. R., Brune M., and Johnson K. A. (1995) Pathway of processive ATP hydrolysis by kinesin. Nature 373, 671–676. 31. Brune M., Hunter J. L., Howell S. A., Martin S. R., Hazlett T. L., Corrie J. E. T., et al. (1998) Mechanism of inorganic phosphate interaction with phosphate binding protein from Escherichia coli. Biochemistry 37, 10370–10380. 32. Hirshberg M., Henrick K., Haire L. L., Vasisht N., Brune M., Corrie J. E. T., et al. (1998) The crystal structure of phosphate binding protein labeled with a coumarin fluorophore, a probe for inorganic phosphate. Biochemistry 37, 10381–10385. 33. Nixon A. E., Hunter J. L., Bonifacio G., Eccleston J. F., and Webb M. R. (1998) Purine nucleoside phosphorylase: its use in a spectroscopic assay for inorganic phosphate and to remove inorganic phosphate with the aid of phosphodeoxyribomutase. Anal. Biochem. 265, 299–307. 34. Webb M. R. (2003) A fluorescent sensor to assay inorganic phosphate. In: Johnson K.A., ed. Kinetic analysis of macromolecules: a practical approach. Oxford, UK: Oxford University Press, 131–152. 35. Thomas C. D., Balson D. F., and Shaw W. V. (1990) In vitro studies of the initiation of Staphylococcal plasmid replication. Specificity of RepD for its origin (oriD) and characterization of the RepD-ori tyrosyl ester intermediate. J. Biol. Chem. 265, 5519–5530. 36. Bird L. E., Brannigan J. A., Subramanya H. S., and Wigley D. B. (1998) Characterisation of Bacillus stearothermophilus PcrA helicase: evidence against an active rolling mechanism. Nucleic Acids Res. 26, 2686–2693. 37. Soultanas P., Dillingham M. S., Papadopoulos F., Phillips S. E., Thomas C. D., and Wigley D. B. (1999) Plasmid replication initiator protein RepD increases the processivity of PcrA DNA helicase. Nucleic Acids Res. 27, 1421–1428. 38. Eccleston J. F., Hutchinson J. P., and White H. D. (2001) Stopped-flow techniques In: Harding S. E. and Chowdry B.Z., eds.Protein ligand interactions: structure and spectroscopy. A practical approach series. Oxford, UK: Oxford University Press, 201–237. 39. Johnson K. A. (2003) Introduction to kinetic analysis of enzyme systems. In: Johnson K.A., ed. Kinetic analysis of macromolecules: a practical approach. Oxford: Oxford University Press, 1–18.

Chapter 3 Single-Molecule FRET Analysis of Helicase Functions Eli Rothenberg and Taekjip Ha Abstract In recent years, advancements in single-biomolecule probing techniques have provided critical information on and greater insight into the nature of biomolecules. Of significance is the application of single-molecule fluorescence resonance energy transfer (smFRET) to probe isolated events and changes at the nanometer scales. In particular, the study of helicases using smFRET has supplied much information regarding the nature and dynamics of these enzymes and provided a toolbox for further investigations. In this chapter we provide a general guide for the construction and execution of single-molecule FRET assays for the study of helicase properties and functionalities. Key words: Helicase, DNA, single molecule, FRET, binding, translocation, unwinding.

1. Introduction Single-molecule techniques enable us to uncover specific features otherwise masked by the averaging of the ensemble measurements. Of particular advantage is the single-molecule technique based on FRET (1), which utilizes the energy-transfer interaction between a pair of reporter molecules (FRET pair), donor and acceptor, thus allowing us to probe distance changes on a range of a few nanometers. This approach has proven to be extremely beneficial in probing different functional aspects of various helicases (2–6). Our studies of DNA helicases’ functions have revealed many important characteristics, providing a toolbox for assaying such enzymes. Investigating DNA-binding properties of helicases revealed the binding orientation and mechanism of bacterial Rep helicase (3) and archaeal MCM helicase substrate specificity (5). Studies on the translocation kinetics of single M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_3, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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Rep (4), PcrA, and UvrD (Jeehae Park, ‘‘Reeling in DNA One Base at a Time: Repetitive DNA Looping Coupled with PcrA Helicase Translocation,’’ unpublished work) helicases led to the discovery of unique repetitive shuttling behaviors, revealing specific features of these helicases’ translocation mechanisms. Investigations on the unwinding of dsDNA by the bacterial Rep helicase showed unwinding re-initiation (2), while viral NS3 (6) and human Bloom helicases (Jaya and Yodh, ‘‘Single-Molecule Study of Bloom Syndrome Helicase Reveals Repetitive Unwinding via Strand-Switching,’’ unpublished work) revealed a repetitive unwinding behavior that disclosed the unwinding mechanisms of these helicases. Further studies of unwinding dynamics of the T7 phage helicase revealed a cooperative unwinding processivity (Manjula Pandey, ‘‘Kinetic Coupling of DNA Primase-Helicase and DNA Polymerase Coordinates DNA Replication,’’ unpublished work). The general guidelines for approaching and designing such assays are provided herein. The design of a suitable smFRET experiment for helicase functionality depends on a number of factors. Initially, the functional aspects that are intended to be probed in the assay need to be defined. These could be divided into three general categories: 1. Binding of the helicase to the DNA substrate. 2. Translocation of the helicase along ssDNA. 3. Unwinding of dsDNA. The above functionalities are closely intertwined and the outcome of one assay would most likely contribute to the interpretation of another. Section 3 will provide different assays for probing these functionalities. When approaching an assay, one should first classify what type of reaction occurs, and construct the experimental probing scheme accordingly. For instance, monitoring single-turnover short-lived reactions, such as a complete unwinding reaction of dsDNA, would require flow-type experiments. In this type of experiment the initiation and progression of the reaction are continuously monitored. In other cases, if the reaction is longer lived, or cyclic, where re-initiation occurs, then data collection may be carried out at different time intervals following the initiation of the reaction. In this chapter we will provide general protocols for assaying the functions of helicases using smFRET. First the materials and general instrumentation used will be described. We note that a comprehensive guide for the construction of a home-built smFRET setup is not provided here and to the inquisitive reader we recommend Ref. (7) for an excellent tutorial on the topic.

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2. Materials 2.1. Reagents

We list the commonly required materials for these experiments. Other necessary reagents that are not listed here are of life sciences grade and may be obtained commercially. 1. Aminopropylsilane (United Chemical Technologies) (store at –20C). 2. Glucose oxidase (Sigma). 3. Catalase (Roche). 4. Oligonucleotides (IDT technology). 5. Coverslips: VWR 240  40 mm, CAT. No. 48393 230 (vwr.com). 6. Standard 300  100  1 mm microscope slides (Fisher). 7. mPEG-SC: MW 5000, 1 g, lot#101-68 (Laysan Bio Incorporated). 8. Biotin-PEG-SC: MW 5000 (Laysan Bio Incorporated). 9. Neutravidin: 31000, 10 mg (http://www.piercenet.com). Prepare a concentration of 5 g/mL in T50 buffer. Store in 4C. 10. T50 Buffer: 10 mM Tris, pH 8, 50 mM NaCl. 11. Drill bits: Kingsley North Inc.: 1-0500-100 (www. kingsleynorth.com). 12. Permanent double-sided tape (Scotch/3 M). 13. Epoxy, 5-min (http://www.devcon.com/). 14. Tubing for flow experiments. ETT-28 (http://www. weicowire.com/).

2.2. Preparation of Gloxy

1. Put 100 mL of T50 buffer in a 250-mL tube. Add 20 mL of catalase. 2. Add 10 mg of glucose oxidase. Centrifuge at 10,000 rpm for 1 min. 3. Recover the yellow supernatant. Store in 4C; it is good for 3 weeks.

2.3. Instrumentation

As mentioned above, for a thorough guide on the assembly of homebuilt TIRF-FRET microscope please review Ref. (7).

2.4. Software and Analysis

When analyzing smFRET data, it is best to begin by looking at representative smFRET histograms. These histograms are constructed by averaging the initial 8–10 data points from many smFRET trajectories in the smFRET histograms. For substrates having a well-defined donor–acceptor distance, a second peak will appear corresponding to that distance, as shown in Fig. 3.1 for a DNA substrate having a separation of seven nucleotides between the donor and acceptor. The donor-only peak serves as the zero FRET mark of the histogram.

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Fig. 3.1. smFRET histogram for the substrates imaged in Fig. 3.5b. Two picks are shown in the histogram. The donor-only peak appears at approximately zero FRET efficiency corresponding to substrates having no emitting acceptor, while the highly excited acceptor peak appears at FRET efficiency of about 0.85.

For more in-depth analysis the specific features of single donor/acceptor trajectories should be monitored closely. For instance, Fig. 3.2 shows two trajectories exhibiting changes: one

Fig. 3.2. Simulated single-molecule trajectories. a,–b. Trajectory of donor (grey) and acceptor (black) and their corresponding FRET efficiency trajectory, showing several well-defined and long-lived FRET values. c. Rapidly and periodically changing FRET efficiency similar to observed repetitive translocation and unwinding.

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showing discrete changes with substantial dwell times, while the other showing rapid periodical changes. Analysis of these types of trajectories can provide useful measurements such as dwell times, frequency and magnitude of FRET changes, and intensity of donor and acceptor.

3. Methods 3.1. Coverslips and Slides Preparations and Assembly 3.1.1. Cleaning

Here we provide a basic glass cleaning protocol, though several protocols may be suitable for single-molecule experiments. 1. Rinse and fill the glass container with MilliQ-H2O. Fill with acetone. Sonicate for 15 min. 2. Fill with 1 M KOH. Sonicate for 20 min. Rinse slides thoroughly with MilliQ-H2O. 3. Burn the slides with propane torch on each side for 30 s. Burn the coverslips for 1 s on one side. Place back in the container (see Note 1).

3.1.2. Aminosilanization and PEGylation

1. Pour 150 mL of MeOH into the flask. Add 7.5 mL of acetic acid (glacial) with a glass pipette. Add 1.5 mL of aminopropylsilane by a glass pipette into the flask and mix well. 2. Pour the mixture quickly into both slide and coverslip containers. Incubate for 10 min on bench, sonicate for 1 min, and then incubate on bench for an additional 10 min. After completion of aminosilanization, rinse coverslips and slides with MeOH and water and dry them with N2(g). Place the coverslips inside tip boxes. Place slides inside tip boxes (see Note 2). 3. Measure 1–2 of biotin-PEG (for five slides) and place inside a 1.5 mL tube. 4. Measure 40 mg of mPEG and put it in the same tube. 5. Add 320 mL of the PEGylation buffer (10 mL MilliQ-H2O + 84 mg sodium bicarbonate) and mix gently with pipette. Centrifuge for 1 min at 10,000 rpm. 6. Drop 70 mL of it on each slide. Gently place a coverslip on the top of each slide (be careful not to create bubbles). Place boxes in a dark, well-leveled location. Incubate for 2–3h. 7. After the incubation period, disassemble the slides, rinse them thoroughly with MilliQ-H2O, and dry completely with N2(g). Store and assemble the slides according to your application. Always store them in the dark (see Note 3).

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3.1.3. Assembly of Slides and Coverslips into Flow Chambers

The number of channels in the flow chamber depends on the desired experiments and applications. The holes should be drilled in the glass prior to cleaning, using a Dremel and diamond-coated drill bits. For a single experiment or flow experiments, a single diagonal channel would suffice, as shown in Fig. 3.3, while for running several experiments sequentially or simultaneously several channels may be used (Fig. 3.4). 1. Place the slide on a stable surface with the PEG-coated surface facing up. 2. Place two strips of double-sided tape on the slide bordering a single diagonal channel between the drilled flow holes (see Fig. 3.3 and Note 5). 3. For assembly of multiple channels, place parallel strips of double-sided tape as spacers between channels (refer to Fig. 3.4). 4. Gently place the coverslip on top of the slide. Cut doublesided tape around the coverslip and carefully peel off the unsandwiched portions. 5. Reinforce double-sided adhesion by carefully pressing tape regions through the coverslip (using a pipette tip). Seal open regions of the channels with 5-min epoxy (see Note 4).

Fig. 3.3. Assembly of single-channel diagonal flow chamber (see text for protocol). a. After drilling the holes and cleaning, the double-sided tape and coverslip is placed. b. The channel is filled with buffer and sealed with epoxy. c. The flow apparatus is assembled and the chamber is placed on the microscope.

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Fig. 3.4. Assembly of multichannel flow chamber (see text for protocol). a. After drilling the holes and cleaning, small strips of double-sided tape are placed as spacers and the coverslip is placed on top. b. The channel is filled with buffer and (c) sealed with epoxy. d. Buffer is added with a pipette directly to the channel and imaged with the microscope.

6. For flow channel assembly, cut two pieces of tubing, approximately 20–30 each. Carefully insert a syringe needle into the end of one of the tubes. 7. Insert the two tubes directly into the drilled holes in the glass. 8. Verify flow through the channel. Insert the loose end of one tube into a tube containing buffer. Pull buffer through the channel using the syringe. Seal the tubing-holes connections using 5-min epoxy (refer to Fig. 3.3). 3.2. Sample Preparation 3.2.1. Nonspecific Binding

Checking for surface integrity and nonspecific binding is a prerequisite for the reliability of surface-tethered experiments. Please refer to Fig. 3.5 for an example of nonspecific labeled protein binding and specific binding of DNA substrates. 1. Image surface. 2. Add 1 nM of Cy3-labeled DNA and image the surface. If image does not contain fluorescent spots resulting from nonspecifically bound DNA, use T50 buffer to wash off channels with 8 channel volumes (see Note 6). 3. To determine the level of nonspecific binding by protein, add protein labeled with Cy3 or Cy5 at similar concentrations as intended to be used in the experiments (at least 1 nM) and

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Fig. 3.5. Nonspecific and specific binding in TIRF-FRET imaging. a. Nonspecific binding of Cy3 (donor) labeled protein. The left-donor channel is saturated. b. Specific binding with optimal coverage of a DNA substrate exhibiting high FRET. The right-acceptor channel shows a number of spots corresponding to the fluorescence of Cy5 molecules.

image the surface. Figure 3.5a shows an example of high levels of nonspecifically bound Cy3-labeled protein. If no fluorescent spots resulting from nonspecific binding are observed, use at least 8 channel volumes of T50 buffer to wash off each channel. 3.2.2. Surface Tethering of DNA Constructs

1. In a 1.5 mL tube, add 960 mL of T50 buffer and 40 mL of Neutravidin stock, giving a final concentration of 0.2 mg/ mL. Add 100 mL of Neutravidin solution to each channel. Incubate for 1 min. 2. Wash off Neutravidin with 400 mL T50 buffer (4–6 channel volumes). 3. Add 100 mL of 30 pM biotinylated DNA (see Note 7). Wait for 5 min and wash off with 400 mL T50 buffer.

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4. Image surface and obtain the density of fluorescent spots. If the number of spots is sufficient, the channel is now ready for the experiments. Refer to Fig. 3.1 for an example of desired density of spots for a high FRET substrate (see Note 8). 3.2.3. Helicase Activity Assays

As discussed earlier, the three main activity assays would be helicase–DNA binding, ssDNA translocation, and dsDNA unwinding. The experimental components for which concentrations should be optimized for kinetic analysis include helicase, hydrolyzable nucleotides and analogues, and salt (see Note 9). The various helicase functionality assays differ mainly in the choice of FRET pair and substrates that will be used, which consequently will define the measured functionality. Besides the various substrates and FRET pairs, the manner by which the experiments are executed is similar. Hence, we will provide some choices for substrates to be used in each type of assay, followed by a general assay that can be used to probe each of the pairs/substrates. The total volume of the imaging/reaction buffer to be added to the channel is typically 100 mL (see Note 10). In a flow-type experiment, data recording is started several seconds prior to the flow of imaging/reaction buffer. For recording data, initially take three long (1000 frames each) consecutive movies followed by ten short movies (30–50 frames each). The longer movies will provide information on single-molecule dynamics over time while the shorter movies will provide data on multiple molecules, which can be used to construct statistically significant fluorescence intensity and FRET histograms.

3.2.4. Imaging Buffer

All single-molecule reactions will be carried out in imaging buffer with an oxygen scavenger system (gloxy/catalase) to reduce the photobleaching rate and BME for reduced blinking. Buffer is made immediately before addition to the channel. 1. In a 250 mL tube prepare 98 mL of reaction buffer already containing the desired reactants (helicase, ATP, etc.) and 0.4% beta-D-glucose. Add 1 mL of Gloxy. Add 1 mL of BME (see Note 11). Mix by pipetting up and down several times (being careful to avoid forming bubbles) and add to the channel. 2. Capture data.

3.3. Binding Assay with Labeled DNA and Protein 3.3.1. Binding Assays

To verify binding and gain structural and organizational information on DNA-bound helicase, the FRET between a labeled helicase and the DNA substrate is monitored. The choice of which will serve as donor and which as acceptor would depend on the binding stoichiometry and labeling efficiency of the helicase. Figure 3.6 shows typical substrates for binding assays. The fork substrate in Fig. 3.6a may be used to monitor the distance

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between the tails, which may be drawn together or restricted due to binding of a helicase (Fig. 3.6a (ii)). In the substrate shown in Fig. 3.6b, the distance between the end and the junction is monitored and may display a change upon binding. An example of such a change is shown in Fig. 3.6b (iii), where binding of ssoMCM helicase to such a substrate with a tail of 40 nt shows a shift in the peak in the smFRET histograms. Figure 3.6c shows a scheme for binding assays that involves measuring FRET between the labeled DNA and the labeled helicase. In this assay no FRET will be detected unless the helicase binds the DNA substrate. By using various DNA substrates or a helicase labeled in different locations, the resultant smFRET histograms may provide domain orientation and structural information, as illustrated in Fig. 3.6c (i)–(iii).

Fig. 3.6. DNA substrates for binding assays. a. (i ),(ii ) End-labeled fork substrate for monitoring the change between the single-stranded ends. b. (i ),(ii ) End- and junctionlabeled substrate for monitoring distance change between junction and tail, and helicase loading on the tail. (iii) FRET change induced by loading of helicase on a substrate as shown on the left, having a tail of 40 nt. Blue histogram represents substrate when no protein is present. Red histogram shows a distinct shift to lower FRET in the histogram after helicase was added, representing helicase-induced stretching of the DNA. c. Substrate for probing binding of labeled helicase.

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Translocation can be monitored in three principal assays, illustrated in Fig. 3.7. First, the distortion of the ssDNA track resulting from the translocation can be probed. This is accomplished by monitoring the FRET change along the track itself using a FRET pair on ends of the track, as illustrated in Fig. 3.7a. Since translocation involves a relative motion of the helicase to the DNA substrate, measuring the FRET change between the moving helicase and a fixed point on the DNA track may provide information on translocation speed and step size. Figure 3.7b shows the scheme for such assays. In this assay, no FRET will be observed prior to binding of the helicase. Finally, probing helicase’s conformational changes associated with translocation and hydrolysis can be done by using a helicase labeled with both donor and acceptor and an unlabeled DNA (see Notes 7 and 8), as illustrated in Fig. 3.7c.

Fig. 3.7. DNA substrates for translocation assays. a. (i )–(iii ) End- and junction-labeled substrate to monitor track distortion and end recapture. b. (i )–(iii ) End-labeled substrate to probe translocation of labeled helicase along the track. c. (i )–(iii ) Unlabeled substrates to probe conformational changes of helicase labeled with donor and acceptor.

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Lastly, an additional assay that we wish to include in conjunction with the binding assays is a helicase binding-stability/dissociation assay (Section 3.3.5). This assay is complementary to the binding assays (Sections 3.3.1 and 3.3.4) and may be done in the same flow chamber immediately following binding experiments. Here, after binding is verified, the channel is washed with buffer to remove excess helicase in solution and then the changes in FRET associated with dissociation are monitored. 3.3.3. Unwinding Assays

The principal role of helicase is the unwinding of dsDNA by harvesting energy from nucleotide hydrolysis, resulting in separation of dsDNA into two strands of ssDNA. Generally, unwinding is best approached by monitoring FRET changes on dually labeled DNA in the vicinity of the duplex region of the DNA substrate, and unlabeled helicase, as illustrated in Fig. 3.8. The substrates for unwinding may be partial duplexes, having a free tail for the loading of the helicase (Fig. 3.8a (iii),(iv)), or forked having both tails (Fig. 3.8a (i),(ii)). The substrate may be orientated relative to the surface through tethering, either via the blunt end of the duplex (Fig. 3.8a (i)–(iii)) or in reverse orientation via the ssDNA tail such that the duplex is away from the surface (Fig. 3.8a (ii)–(iv)). The

Fig. 3.8. DNA substrates for unwinding assays. a. (i )–(iv ) Partial duplex substrates for unwinding, with either single tails (iii ),(iv ) or forked (i ),(ii ) structures with the substrate being either directly tethered to the surface through its duplex blunt end or tethered via a ssDNA tail, away from the surface (i.e., reverse orientation). b. (i )–(iii ) Scheme for unwinding where donor strand disengages from substrate after unwinding.

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biotin surface-tethered strand should be labeled with Cy5 such that if full unwinding occurs, as illustrated in Fig. 3.8b (i)–(iii), the Cy3 donor-labeled strand would be released from the substrate and the fluorescence signal will diminish. 3.3.4. Helicase General Activity Assay

This assay is to be customized according to the desired functionality and probing scheme, as specified in Sections 3.3.1, 3.3.2, and 3.3.3 (also see Note 12). 1. Check nonspecific binding properties of the surface (Section 3.2.1) – if a labeled protein is to be used, measure nonspecific binding of labeled protein. 2. Attach substrate of interest (depending on desired activity, see Sections 3.3.1, 3.3.2, and 3.3.3). 3. Place all reagents (helicase, ATP, gloxy, etc.) in a covered ice bucket. 4. Measure FRET of substrate alone as a control – add imaging buffer and take ten short movies (20–40 frames) and three long ones (1000 frames). 5. Prepare reaction buffer (see Note 12), add and take ten short movies (20–40 frames) and three long ones (1000 frames). 6. If the studied reaction can be re-initiated, change the concentration of one of the reaction variables and repeat step 5.

3.3.5. Helicase Dissociation Assay

1. For binding assays, following each of the above additions, wash channel using 400 mL T50 buffer. Add imaging buffer and image. Take two long and ten short movies. 2. Repeat, after each concentration, addition in the above assays. Further suggestion regarding assays and analysis are provided in Notes 13 and 14.

4. Notes 1. Heated coverslips tend to bend; therefore, pass the coverslips quickly through the flame. 2. The layer of PEG solution in between the glass tends to dry; incubate in vapor-saturated environment (empty pipette tip box with a flooded floor). 3. Can be stored in –20C for extended periods. Prior to assembly, let it thaw and reach room temperature. 4. Prior to sealing sides with epoxy, carefully add 100 mL of T50 to the channels. This is done so epoxy will not fill the channel. Apply epoxy to the sides using a pipette tip and let it dry.

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5. Wash with two additions of 200 mL for 50 mL channel volume. When adding solution to a channel, pipette carefully so no bubbles are formed; place a folded Kimwipe on the exit hole to absorb flow-through. 6. For some substrates adjusting the buffer pH to the range of 8–8.5 may lower nonspecific binding, particularly for ssDNA. 7. The concentration of substrate to be added varies and would depend on labeling efficiency of the substrates and the actual amount of biotin and Neutravidin on the PEG surface; typically the concentration range of 30–200 pM yields a sufficient number of surface-bound fluorescence spots. 8. If the initial number of spots is low (due to reasons given in Note 7), gradually increase the concentration of substrate until the desired number of fluorescent spots is reached. 9. T50 buffer with 10 mM of added Mg is typically required for helicase–DNA binding (magnesium acetate or MgCl2). Alternatively enzymatic activity buffer, such as NEBuffer 4 (New England Biolabs), may be used. 10. In the case of narrow channels or if protein is scarce, smaller quantities can be used as long as the concentration of reactants remains the same. 11. BME does not provide a good blinking suppressor in the case of a FRET pair along rigid dsDNA regions. One can use buffer containing TROLOX for blinking suppression (8). 12. For each of the functionalities that are being probed, there exist matrixes of variable quantities. For binding assays, salt and helicase concentration should be titrated. For translocation and unwinding assay, salt, helicase, and hydrolyzable nucleotide concentration should be titrated. Accordingly, for each of these variables a separate control is required. For instance, in translocation and unwinding assays, prior to adding the helicase to the reaction buffer, a control containing imaging buffer plus hydrolyzable nucleotides should be performed in order to ensure that the substrate FRET is not affected. 13. Careful and methodological data analysis is the principal part in quantifying and interpreting the resulting smFRET data. For specific IDL and Matlab code requests, the readers are encouraged to contact the authors for their availability. 14. We note that the assays provided here must be optimized depending on the helicases that are being investigated. Optimizing the assays and constructing appropriate DNA substrates should depend on general helicase characteristics such as dissociation constants and effective concentration, helicase oligomeric forms, DNA footprint, directionality,

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and processivity. We recommend that prior to expediting smFRET experiments, one should perform fluorimeter activity assays in the bulk, with the same (or similar) DNA constructs as are intended for the smFRET measurements.

Acknowledgments The authors would like to thank Jaya Yodh for carefully reading this chapter and providing helpful comments. The authors thank Jaya Yodh, Salman Sayed, Jeehae Park, and Hamza Balci for providing data, comments, and manuscripts in preparation. E.R. is a fellow of the NSF Center for the Physics of Live Cells at the University of Illinois, and he acknowledges its support. References 1. Ha T., Enderle T., Ogletree D. F., Chemla D. S., Selvin P. R., and Weiss S. (1996) Probing the interaction between two single molecules: Fluorescence resonance energy transfer between a single donor and a single acceptor. Proc. Natl. Acad. Sci. USA 93, 6264–6268. 2. Ha T., Rasnik I., Cheng W., Babcock H. P., Gauss G. H., Lohman T. M., and Chu S. (2002) Initiation and re-initiation of DNA unwinding by the Escherichia coli Rep helicase. Nature 419, 638–641. 3. Rasnik I., Myong S., Cheng W., Lohman T. M., and Ha T. (2004) DNA-binding orientation and domain conformation of the E-coli Rep helicase monomer bound to a partial duplex junction: Single-molecule studies of fluorescently labeled enzymes. J. Mol. Biol. 336, 395–408. 4. Myong S., Rasnik I., Joo C., Lohman T. M., and Ha T. (2005) Repetitive shuttling of a

5.

6.

7.

8.

motor protein on DNA. Nature 437, 1321–1325. Rothenberg E., Trakselis M. A., Bell S. D., and Ha T. (2007) MCM fork substrate specificity involves dynamic interaction with the 50 tail. J. Biol. Chem. 282, 34229–34234. Myong S., Bruno M. M., Pyle A. M., and Ha T. (2007) Spring-loaded mechanism of DNA unwinding by hepatitis C virus NS3 helicase. Science 317, 513–516. C. Joo and T. Ha (2008) Ch 2. ‘‘SingleMolecule FRET with Total Internal Reflection Microscopy’’ In, ‘‘Single Molecule Techniques: A Laboratory Manual’’, Eds. Pauls R. Selvin and Taekjip Ha. Cold Spring Harbor Laboratory Press, New York, ISBN 978-087969775-4, pp. 507. Rasnik I., McKinney S. A., and Ha T. (2006) Nonblinking and longlasting singlemolecule fluorescence imaging. Nat. Methods 3, 891–893.

Chapter 4 Kinetics of Motor Protein Translocation on Single-Stranded DNA Christopher J. Fischer, Lake Wooten, Eric J. Tomko, and Timothy M. Lohman Abstract The translocation of nucleic acid motor proteins along DNA or RNA can be studied in ensemble experiments by monitoring either the kinetics of the arrival of the protein at a specific site on the nucleic acid filament (generally one end of the filament) or the kinetics of ATP hydrolysis by the motor protein during translocation. The pre-steady state kinetic data collected in ensemble experiments can be analyzed by simultaneous global non-linear least squares (NLLS) analysis using a simple sequential ‘‘n-step’’ mechanism to obtain estimates of the rate-limiting step(s) in the translocation cycle, the average ‘‘kinetic step-size,’’ and the efficiency of coupling ATP binding and hydrolysis to translocation. Key words: Translocase, kinetics, ATPase, helicase, motor protein.

1. Introduction The ability to translocate processively and with biased directionality along a nucleic acid filament is central to the biological function of several enzymes including polymerases (1), helicases (2–4), chromatin remodelers (5, 6), some nucleases (7, 8), and some restriction enzymes (9–11). These ‘‘molecular motors’’ all use the chemical potential energy obtained through the binding and hydrolysis of nucleoside triphosphates (NTP or dNTP) to perform the mechanical work of directional translocation along the filament. An understanding of the translocation mechanisms of these motor proteins requires quantitative kinetic information to obtain the rate constants, processivities, kinetic step-sizes, and ATP coupling stoichiometries associated with translocation. M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_4, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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Here, we describe the use and analysis of pre-steady state ensemble kinetic approaches (3, 6, 10–17) to probe the translocation mechanisms of processive translocases along nucleic acids using a simple sequential ‘‘n-step’’ kinetic model. The application of this methodology provides an accurate determination of macroscopic kinetic parameters such as the rate of net forward motion of the translocase along the nucleic acid and the net efficiency at which the hydrolysis of ATP is coupled to this net forward motion. However, the estimates of microscopic kinetic parameters, such as the kinetic step-size of translocation, can be inflated under some circumstances if non-uniform motion occurs during translocation.

2. Materials 2.1. Matching Experimental Conditions to Model Assumptions

The kinetic model and associated equations used here assume that no more than one translocase is bound to each nucleic acid. Thus, the application of these equations to the analysis of kinetic data requires that the experiments are performed under conditions that favor this binding distribution; this is generally achieved by performing the experiments under conditions where the concentration of the nucleic acid is in excess of the concentration of the translocase. This model also assumes that any translocase that is initially free in solution at the start of the reaction or that dissociates from the nucleic acid during translocation is prevented from rebinding to the nucleic acid. This is accomplished experimentally by including a protein trap. When selecting such a trap it is best to use one that does not stimulate the ATPase activity of the translocase (18). This will allow for more straightforward and simple analysis of the ATPase activity of the translocase that is associated with translocation.

3. Methods 3.1. Mathematical Model for Translocation

The sequential ‘‘n-step’’ kinetic mechanism shown in Scheme 4.1 has been used to model helicase translocation and its coupling to ATP hydrolysis (13, 18). In this mechanism (13), depicted in Fig. 4.1, a translocase with an occluded site size of b nucleotides and a contact size of d nucleotides binds with polarity to a nucleic acid filament, L nucleotides long. The contact size, d, represents the number of consecutive nucleotides required to satisfy all contacts with the translocase and is thus less than or equal to the

Kinetics of Motor Protein Translocation

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Scheme 4.1

occluded site size. In this discussion we will assume that translocation along the nucleic acid is directionally biased from 30 to 50 , but the results are equally applicable to a translocase that exhibits the opposite directional bias.

Fig. 4.1. Kinetic model for ATP-dependent protein translocation along a nucleic acid filament. Panel A: A cartoon depicting the binding of a translocase with a contact size d and occluded site size b to a nucleic acid filament of length L. As shown in this cartoon, the contact size, d, is always less than or equal to the occluded site size, b. Panel B: Cartoon showing the model used to describe enzyme translocation along a nucleic acid filament. The line segments represent the nucleic acid and the triangles represent the translocase. The translocase binds randomly, but with polarity, to the nucleic acid and upon binding and hydrolysis of ATP proceeds to translocate toward the 50 end of the filament in discrete steps with rate constant kt. The rate constant of dissociation during translocation is kd. Upon reaching the 50 end of the filament, the translocase dissociates with a rate constant kend. Dissociated translocases bind to a protein trap, T, and are thereby prevented from rebinding the nucleic acid.

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The translocase is initially bound i translocation steps away from the 50 end, with concentration Ii. The number of translocation steps, i, is constrained (1
Kinetics of Motor Protein Translocation



f50 ðtÞ ¼ A L

1



 n  1 kt kend þ s kt þ kd þ s

49

[1]

For the case where all the proteins are initially bound at random positions along the nucleic acid, the equation for the timedependent accumulation of protein at the 50 end of the nucleic acid is given by equation [2]. A L 1 1 þ n r     n  1 kt  r kt  1þ 1 s þ kend s þ kd s þ kt þ kd

f50 ðtÞ ¼

[2]

The scalar A in equations [1] and [2] allows for conversion of the concentration of protein bound at the 50 end of the nucleic acid into a signal that can be measured experimentally (e.g., a spectroscopic change) (13, 14, 18). Similarly, equations [3] and [4] are expressions for the timedependent production of ADP or Pi, due to ATP hydrolysis by the translocases (13). In equations [3] and [4], I(0) is the concentration of translocase initially bound to the nucleic acid (at time t ¼ 0). Equation [3] describes the ATP production occurring when all proteins are initially bound at the same position (e.g., the 30 end of the nucleic acid) and equation [4] describes the ATP production occurring when all the proteins are initially bound at random positions along the nucleic acid.   n  2 0 kt ck  1  t kt þkd þs 1 ADPðtÞ ¼I ð0Þ  L 1 4 @ s kd þ s [3]  n !# ka kt þ kend þ s kt þ kd þ s

" ADPðtÞ ¼

I ð0Þ L 1 1þnr 0 kt @1 þ þ

   n  c  kt  r  nðkd þ sÞ þ kt kt þkktd þs  1

1 s

ðkd þ sÞ2

 

kt r 1

kt kt þkd þs

kd þs

n 1

!#

[4]

A

kend þ s

The maximum number of translocation steps, n, for a nucleic acid of a length, L, is related to the translocation kinetic step-size, m, and the translocase contact size by equation [5].

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Fig. 4.2. Stopped-flow assays for monitoring the pre-steady state kinetics of enzyme translocation along nucleic acid filaments. Panel A: A translocase is pre-bound to a nucleic acid filament labeled at the 50 end with a fluorescent dye, then rapidly mixed with ATP, Mg2+, and heparin (protein trap) to initiate translocation. When the translocase nears the 50 end of the nucleic acid the fluorescence of the dye is either quenched or enhanced. Example time courses are shown for three different lengths of nucleic acid. Panel B: A translocase is pre-bound to a nucleic acid filament and then rapidly mixed with

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Ld [5] m Equation [5] can be re-expressed as equation [6], which allows for the determination of m from the experimentally determined dependence of L on n. n¼

L ¼ mn þ d

[6]

3.2. Monitoring the Kinetics of the Arrival of the Translocase at a Specific Site on the Nucleic Acid

The first experimental method we consider is a stopped-flow fluorescence approach first introduced by Dillingham et al. (20) and subsequently modified by Fischer et al. (6) and is depicted in Fig. 4.2A. The method utilizes a series of oligodeoxthymidylates of varying lengths (L (dT)L) that have a fluorophore attached covalently to either the 30 or 50 end of the nucleic acid. The fluorophore is chosen such that a fluorescence intensity change occurs when the translocase is bound at that end possessing the fluorophore. In this way, one can monitor the change in concentration of translocases bound at the end of the nucleic acid resulting from arrival of translocases due to translocation from other sites on the nucleic acid and dissociation of translocases from the end. Quantitative analysis of a series of these time courses performed as a function of L using equation [1] or [2] allows one to estimate the microscopic kinetic parameters associated with translocation of the enzyme along the nucleic acid. A subsequent estimate of the kinetic step size, m, is then obtained from the analysis of the dependence of n on L through equation [6]. We have found that Cy3 and fluorescein have generally yielded good signal changes for the translocases that we have studied (14, 18, 21, 22). The directionality of translocation can be determined by comparing the time courses observed when the fluorophore is attached to the 30 versus the 50 end of the nucleic acid (13, 14, 20). Specifically, characteristic changes in the fluorescence time course as a function of increasing nucleic acid length (e.g., an increase in both the time required to reach peak fluorescence and the breadth of the fluorescence peak as shown in Fig. 4.2A) will occur if the translocation direction is biased toward the fluorophore, but not when it is biased away from the fluorophore.

3.3. Monitoring the Kinetics of ATP Hydrolysis by the Translocase During Translocation

Enzyme translocation along nucleic acids can also be monitored by measuring the amount of ATP hydrolyzed by the enzyme during translocation. This approach requires transient pre-steady state kinetic experiments rather than steady-state ATPase experiments since steady state rates of ATP hydrolysis will generally be

Fig. 4.2. (continued) ATP, Mg2+, heparin, and an excess concentration of fluorescently labeled phosphate-binding protein (PBP-MDCC) to initiate translocation. As the translocase moves along the filament, ATP is hydrolyzed into ADP and inorganic phosphate (Pi). PBP-MDCC rapidly binds the Pi resulting in an increase in the PBP-MDCC fluorescence. Example time courses are shown for three different lengths of nucleic acid.

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limited by other kinetic processes that are slower than protein translocation (e.g., dissociation and/or rebinding of protein to another nucleic acid molecule). The pre-steady state rate and extent of ATP hydrolysis by the translocating protein can be monitored, for example, by directly measuring the conversion of ATP to ADP using a radioactive assay (23, 24) or by monitoring the release of inorganic phosphate using a fluorescently labeled phosphate- binding protein (18, 19) as depicted in Fig. 4.2B. Analysis of a series of time courses of ATP hydrolysis during translocation performed as a function of DNA length, L, can be analyzed using equation [3] or [4] to determine estimates of the microscopic parameters c and ka and, thus, the macroscopic ATP coupling stoichiometry c/m. In this analysis, the values of the microscopic parameters obtained from the analysis of translocation time courses using Method 3.2 (kt, kd, kend, m, and r) are used as fixed constraints in the application of equations [3] or [4] (18).

4. Notes

1. Significant correlation exists between the parameters in equations [1], [2], [3], and [4]. For this reason, it is best to independently determine as many parameters as possible so that they can be constrained in the NLLS analysis using these equations. For example, the rate of dissociation during translocation (kd in Scheme 4.1) can be determined independently by monitoring the dissociation of enzyme during translocation along an infinitely long nucleic acid (13, 14, 18) 2. It is worth noting that the presence of the fluorophore might affect the rate of translocation and/or the rate of dissociation near the fluorophore (14, 18). It is also possible that variations in the electrostatics of the nucleic acid molecule near its ends may contribute to differences in kt and kd at binding positions near the ends. Thus, for an enzyme that translocates in a 30 to 50 direction, the values for kt and kd obtained from fitting experimental time courses obtained with nucleic acid labeled with a fluorophore at the 30 end may not equal the values of kt and kd that apply in the absence of the fluorophore or to interior regions of the nucleic acid. Similarly, the value of kend obtained from fitting experimental time courses obtained

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with nucleic acid labeled with a fluorophore at the 50 end may not equal the value of kend that applies in the absence of the fluorophore (13, 18). 3. The model (Scheme 4.1) used to obtain equations [1], [2], [3], and [4] assumes translocation occurs via a uniform repetition of irreversible rate-limiting steps, and ignores any nonuniformity in the translocation process. To further evaluate the efficacy of this methodology when such non-uniformity exists, we used a Monte Carlo computer simulation to generate translocation time courses for models that contained non-uniform motion as well as heterogeneity in the rates. We considered backward motion, random pausing, simple heterogeneity in the microscopic translocation rate constant (kt) or step-size (m) for each individual kinetic step, persistent heterogeneity (25) in the microscopic translocation rate constant (kt) or the kinetic step size (m), and repetitive shuttling of the translocase on the nucleic acid (26, 27). We note that the existence of persistent heterogeneity for the entire translocation process, commonly referred to as ‘‘static disorder’’ (28), could result only if each enzyme were chemically or conformationally different over the time period of the experiment. Yet, there are clear examples of such static disorder in processive nucleic acid enzymes, such as RNA polymerase (29) and some helicases (25, 30, 31). These simulated time courses were then analyzed using the simple uniform sequential n-step mechanism (equations [1], [2], [3], [4], [5] and [6]) to obtain estimates of kt c, and m that were then compared to the input values of these kinetic parameters. Our analysis of these simulated data sets demonstrated that the values of the macroscopic translocation rate (mkt) and the coupling stoichiometry (c/m) obtained using these equations reliably reflect the actual input values used for the simulations regardless of the presence of any non-uniform motion of the translocase. The macroscopic translocation rate is well constrained in the NLLS analysis since it is determined from the dependence on nucleic acid length of the mean arrival time of proteins at the end of the nucleic acid. Similarly, the resulting estimate of c/m provides an accurate estimate of the total ATP consumption associated with the net forward motion of the translocase along the nucleic acid. Intuitively when backward motion or random pausing is introduced, the coupling stoichiometry will increase since some ATP hydrolysis results in backward motion or pausing rather than in forward motion. The individual estimates of m and kt are largely constrained by both the standard deviation of the distribution of arrival times of proteins at the end of the nucleic acid and the dependence of this distribution on nucleic acid length. Importantly, the

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inclusion of simple heterogeneity in kt or m did not affect the estimates of mkt, c/m, m, r, or ka obtained from the analysis of the simulated time courses using equations [1], [2], [3], [4], [5], and [6]. This result is significant since we believe that this simulation of simple heterogeneity is a reasonable model of the stochastic fluctuations in kt and m that are expected for a chemically and structurally homogenous population of translocases. However, any perturbation that increases the standard deviation of the distribution of arrival times at any particular point along the nucleic acid will also increase the estimate of m obtained from the fit. Clearly, introducing backward motion, static disorder, repetitive shuttling, or random pausing will spread out the distribution of arrival times at the 50 end of the nucleic acid and will furthermore increase the dependence of the standard deviation on the length of the nucleic acid. As such, the introduction of any of these perturbations will, in fact, result in an overestimate of m. The exact amount of the overestimate will naturally depend upon the type of perturbation and its magnitude, but also on the magnitudes of the other rate constants associated with translocation since all of these variables are highly correlated in the sequential ‘‘n-step’’ model. 4. Use of the uniform sequential ‘‘n-step’’ model to analyze time courses obtained from experiments in which the translocase initiates at random positions along the nucleic acid requires inclusion of the r parameter in equations [2] and [4]. If the translocase has equal affinity for all potential binding sites on the nucleic acid then r must have a value between 1 and m, depending upon the specific details of the translocation mechanism near the end of the nucleic acid (see (13) for more details). Thus, estimated values of r for which r > m may indicate a failure of the simple model to correctly describe the translocation process, regardless of the quality of the fits. In other words, r can serve as an indicator of potential nonuniformity in the translocation mechanism. It is worth noting that increases in the estimate of r were associated with each of the perturbations we considered with the exception of simple heterogeneity in kt or m. Furthermore, these increases in the estimate of r were especially large when static disorder or repetitive shuttling was incorporated into the simulations. Hence, a value of r>m can be an indicator of non-uniformity or static disorder in the translocation kinetics. 5. The results of analysis of computer simulated data using equations [1], [2], [3], and [4] also indicate that the presence of non-uniform motion can also increase the estimate of the rate of futile ATP hydrolysis at the end of the nucleic acid (ka in Scheme 4.1). Hence the observation of futile hydrolysis

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at the end of the nucleic acid is also an indication that nonuniform motion may be occurring. In such a case, one should perform additional experiments or additional computer simulations to independently determine whether futile hydrolysis at the end of the nucleic acid actually occurs.

Acknowledgments The authors thank Dr. Carsten Timm, Dr. Siyuan Han, Dr. Matthew Antonik, Dr. Karl Maluf, Dr. Aaron Lucius, and Dr. Nathan Baker for useful discussions concerning this manuscript. This research was supported, in part, by startup funding from the University of Kansas (to C.J.F.), an University of Kansas Undergraduate Research Award (to L.W.), and by NIH grants GM045948 (to T.M.L) and P20 RR17708 from the Institutional Development Award (IDeA) Program of the National Center for Research Resources (to C.J.F.). References 1. Kornberg R. D. (2007) The molecular basis of eukaryotic transcription. Proc. Natl. Acad. Sci. U.S.A. 104, 12955–12961. 2. Lohman T. M. and Bjornson K. P. (1996) Mechanisms of helicase-catalyzed DNA unwinding. Annu. Rev. Biochem. 65, 169–214. 3. Lohman T. M., Hsieh J., Maluf N. K., Cheng W., Lucius A. L., Fischer C. J., Brendza K. M., Korolev S., and Waksman G. (2003) DNA helicases, motors that move along nucleic acids: lessons from the SF1 helicase superfamily. In The Enzymes (Hackney D. D. and Tamanoi F., Eds.) Third ed., Academic Press, New York. 4. Matson S. W. and Kaiser-Rogers K. A. (1990) DNA helicases. Annu. Rev. Biochem.. 59, 289–329. 5. Becker P. B. (2005) Nucleosome remodelers on track. Nat. Struct. Mol. Biol. 12, 732–733. 6. Fischer C. J., Saha A., and Cairns B. R. (2007) Kinetic model for the ATPdependent translocation of Saccharomyces cerevisiae RSC along double-stranded DNA. Biochemistry 46, 12416–12426. 7. Kovall R. A. and Matthews B. W. (1998) Structural, functional, and evolutionary relationships between lambda-exonuclease

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and the type II restriction endonucleases. Proc. Natl. Acad. Sci. U.S.A. 95, 7893–7897. Kovall R. A. and Matthews B. W. (1999) Type II restriction endonucleases: structural, functional and evolutionary relationships. Curr. Opin. Chem. Boil. 3, 578–583. Szczelkun M. D. (2002) Kinetic models of translocation, head-on collision, and DNA cleavage by type I restriction endonucleases. Biochemistry 41, 2067–2074. Firman K. and Szczelkun M. D. (2000) Measuring motion on DNA by the type I restriction endonuclease EcoR124I using triplex displacement. EMBO J. 19, 2094–2102. McClelland S. E., Dryden D. T., and Szczelkun M. D. (2005) Continuous assays for DNA translocation using fluorescent triplex dissociation: application to type I restriction endonucleases. J. Mol. Biol. 348, 895–915. Ali J. A. and Lohman T. M. (1997) Kinetic measurement of the step size of DNA unwinding by Escherichia coli UvrD helicase. Science 275, 377–380. Fischer C. J. and Lohman T. M. (2004) ATP-dependent translocation of proteins along single-stranded DNA: models and

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Fischer et al. methods of analysis of pre-steady state kinetics. J. Mol. Biol. 344, 1265–1286. Fischer C. J., Maluf N. K., and Lohman T. M. (2004) Mechanism of ATP-dependent translocation of E.coli UvrD monomers along single-stranded DNA. J. Mol. Biol. 344, 1287–1309. Lucius A. L., Jason Wong C., and Lohman T. M. (2004) Fluorescence stopped-flow studies of single turnover kinetics of E.coli RecBCD helicase-catalyzed DNA unwinding. J. Mol. Biol. 339, 731–750. Lucius A. L., Maluf N. K., Fischer C. J., and Lohman T. M. (2003) General methods for analysis of sequential ‘‘n-step’’ kinetic mechanisms: application to single turnover kinetics of helicase-catalyzed DNA unwinding. Biophys. J. 85, 2224–2239. Lucius A. L., Vindigni A., Gregorian R., Ali J. A., Taylor A. F., Smith G. R., and Lohman T. M. (2002) DNA unwinding step-size of E. coli RecBCD helicase determined from single turnover chemical quenched-flow kinetic studies. J. Mol. Biol. 324, 409–428. Tomko E. J., Fischer C. J., Niedziela-Majka A., and Lohman T. M. (2007) A Nonuniform Stepping Mechanism for E. coli UvrD Monomer Translocation along SingleStranded DNA. Mol. Cell 26, 335–347. Dillingham M. S., Wigley D. B., and Webb M. R. (2000) Demonstration of unidirectional single-stranded DNA translocation by PcrA helicase: measurement of step size and translocation speed. Biochemistry 39, 205–212. Dillingham M. S., Wigley D. B., and Webb M. R. (2002) Direct measurement of singlestranded DNA translocation by PcrA helicase using the fluorescent base analogue 2aminopurine. Biochemistry 41, 643–651. Brendza K. M., Cheng W., Fischer C. J., Chesnik M. A., Niedziela-Majka A., and Lohman T. M. (2005) Autoinhibition of Escherichia coli Rep monomer helicase activity by its 2B subdomain. Proc. Natl. Acad. Sci. U.S.A. 102, 10076–10081. Niedziela-Majka A., Chesnik M. A., Tomko E. J., and Lohman T. M. (2007) Bacillus

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stearothermophilus PcrA monomer is a single-stranded DNA translocase but not a processive helicase in vitro. J. Biol. Chem. 282, 27076–27085. Hsieh J., Moore K. J., and Lohman T. M. (1999) A two-site kinetic mechanism for ATP binding and hydrolysis by E. coli Rep helicase dimer bound to a single-stranded oligodeoxynucleotide. J. Mol. Biol. 288, 255–274. Wong I., Moore K. J., Bjornson K. P., Hsieh J., and Lohman T. M. (1996) ATPase activity of Escherichia coli Rep helicase is dramatically dependent on DNA ligation and protein oligomeric states. Biochemistry 35, 5726–5734. Dessinges M. N., Lionnet T., Xi X. G., Bensimon D., and Croquette V. (2004) Single-molecule assay reveals strand switching and enhanced processivity of UvrD. Proc. Natl. Acad. Sci. U.S.A. 101, 6439–6444. Myong S., Bruno M. M., Pyle A. M., and Ha T. (2007) Spring-loaded mechanism of DNA unwinding by hepatitis C virus NS3 helicase. Science 317, 513–516. Myong S., Rasnik I., Joo C., Lohman T. M., and Ha T. (2005) Repetitive shuttling of a motor protein on DNA. Nature 437, 1321–1325. Lu H. P., Xun L., and Xie X. S. (1998) Single-molecule enzymatic dynamics. Science 282, 1877–1882. Neuman K. C., Abbondanzieri E. A., Landick R., Gelles J., and Block S. M. (2003) Ubiquitous transcriptional pausing is independent of RNA polymerase backtracking. Cell 115, 437–447. Bianco P. R., Brewer L. R., Corzett M., Balhorn R., Yeh Y., Kowalczykowski S. C., and Baskin R. J. (2001) Processive translocation and DNA unwinding by individual RecBCD enzyme molecules. Nature 409, 374–378. Perkins T. T., Li H. W., Dalal R. V., Gelles J., and Block S. M. (2004) Forward and reverse motion of single RecBCD molecules on DNA. Biophys. J. 86, 1640–1648.

Chapter 5 Experimental and Computational Analysis of DNA Unwinding and Polymerization Kinetics Manjula Pandey, Mikhail K. Levin, and Smita S. Patel Abstract DNA unwinding and polymerization are complex processes involving many intermediate species in the reactions. Our understanding of these processes is limited because the rates of the reactions or the existence of intermediate species is not apparent without specially designed experimental techniques and data analysis procedures. In this chapter we describe how pre-steady state and single-turnover measurements analyzed by model-based methods can be used for estimating the elementary rate constants. Using the hexameric helicase and the DNA polymerase from bacteriophage T7 as model systems, we provide stepwise procedures for measuring the kinetics of the reactions they catalyze based on radioactivity and fluorescence. We also describe analysis of the experimental measurements using publicly available models and software gfit (http://gfit.sf.net). Key words: Hexameric helicase, replication, DNA unwinding, T7 bacteriophage, DNA polymerase, DNA synthesis, strand displacement, primer extension, gfit, global regression analysis.

1. Introduction Helicases are motor proteins that use the chemical energy of NTP hydrolysis to separate the strands of double-stranded nucleic acids (1–4). Often helicases work in association with other proteins, such as the DNA polymerase or single-strand binding proteins to perform their function with a greater efficiency. Characterization of the nucleic acid unwinding activity involves measurement of the rate and the processivity of the helicase-catalyzed unwinding reaction. These unwinding parameters can be used as basic handles to understand the mechanism of unwinding and the role of the helicase in a particular biological process. Measurement of the M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_5, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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unwinding rate of helicases as a function of DNA duplex stability provided insights into the active or passive nature of the helicasecatalyzed reaction (5). Measurement of the unwinding rate of phage T7 hexameric helicase in the presence of the DNA polymerase provided insights into the synergistic action of the two motor proteins in DNA replication (6). In this chapter, we describe assays to measure DNA unwinding catalyzed by the helicase, and DNA polymerization catalyzed by the helicase and polymerase proteins using the hexameric ringshaped T7 gp4 helicase as our model system. We outline procedures to fit the kinetic data to specific models that provide kinetic parameters such as the rate of DNA unwinding and the rate of nucleotide incorporation during DNA polymerization. The product of bacteriophage T7 gene gp4 is a ring-shaped protein (7–9) that has both DNA unwinding and primase activities (10). T7 gp4 can move along single-stranded (ss) (11) or doublestranded (ds) (12) DNA and separate DNA strands using the energy of dTTP hydrolysis. The protein can also synthesize short (4- to 5-mer) RNA primers using ATP and CTP at priming sites on the lagging DNA strand. T7 gp4 unwinds dsDNA using the strand exclusion model wherein one strand of the dsDNA is threaded into the central channel of the helicase and the other strand is excluded as the helicase unwinds the dsDNA (13, 14). This strand exclusion model is accepted generally for ring-shaped helicases (15–17). The unwinding rate of T7 gp4 depends on dsDNA stability and its speed of DNA unwinding is slower than its speed of translocation along ssDNA (5). The other key component of T7 replication complex is gp5 DNA polymerase (18–20). Interestingly, the rate of DNA unwinding by T7 gp4 is accelerated by the polymerase and approaches the ssDNA translocation rate of the helicase (6). Accurate measurements of kinetic rates of translocation, unwinding, and polymerization provide both quantitative and qualitative insights about the DNA replication mechanism. 1.1. Methods for Measuring Reaction Kinetics

Unwinding and polymerization pathways comprise many interacting reaction steps such as substrate binding, catalysis, and conformational changes. The large number of these steps makes their identification and characterization increasingly difficult. Therefore, one prerequisite of a successful experimental design is its ability to decouple the pathway: to study a part of the pathway in isolation or while controlling the influence of other parts. Most clear results are produced by experiments that measure kinetics of one fully decoupled reaction step. One strategy to maximize decoupling enabled by recent technological advances involves monitoring reactions occurring with single molecules. These techniques have been successfully applied to both DNA unwinding and polymerization (21–28). Unfortunately, only parts of DNA processing pathways can be currently

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observed at single-molecule level, which brings us back to conventional measurement techniques that integrate signals originating from billions of enzyme molecules. During the course of reaction, as the system reaches a steady state, molecules of enzyme become distributed along the reaction pathway populating all of the reaction species and participating in all reactions simultaneously. Combined signal from such heterogeneous mixture is hard to interpret in terms of individual reaction rates. These problems are overcome by using pre-steady state and single round kinetic techniques. Pre-steady state kinetic measurements involve synchronization of the system – assembling the reaction mixture in a way that populates only one species of the pathway. This can be done, for example, by withholding a component required for the next reaction step. After the reaction in the synchronized mixture is started, but before it reaches a steady state, the measured signal can be attributed to a few steps that follow the synchronization point. The pre-steady state period of reaction is quite short due to a natural tendency of molecular systems to lose synchronization. To extend it and to be able to characterize more reaction steps in one experiment, single round conditions can be used. Single round conditions effectively prevent the system from reaching a steady state by allowing each molecule of enzyme transform no more than one molecule of substrate. In case of an unwinding reaction, this can be achieved by adding a helicase trap at the time of initiation. An excess of ssDNA can capture free helicase molecules from solution preventing them from re-binding to new substrate molecules. Pre-steady state single round approaches enhance our ability to decouple unwinding and polymerization pathways to measure the rates of their reaction steps. 1.1.1. Unwinding Kinetics

DNA unwinding rates have been measured using both bulk and single-molecule techniques (5, 6, 29). Single round conditions simplify interpretation of a bulk kinetic measurement result. Helicases can processively unwind stretches of dsDNA longer than their binding site. Therefore an unwinding reaction can be described as an n-step process and for experiments conducted under single round conditions, the results can be fit with a stepping equation (30, 31). Such analysis of unwinding kinetics data for dsDNA substrates of different lengths can be used for estimating helicase stepping rate and size and for assessing processivity of the helicase, i.e., how far the helicase moves along the DNA before it falls off.

1.1.1.1. Assembly of Helicase on the DNA Substrate

Ring-shaped helicases assemble around ssDNA, and assembly is usually a slow step (relative to the rate of DNA unwinding). Therefore, to measure the unwinding rate (rather than the assembly rate) and to synchronize the reactions, it is important to preassemble the

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helicase on the DNA substrate prior to reaction start. T7 gp4, like other ring-shaped helicases, binds to the DNA only in the presence of its nucleotide substrate (dTTP in the case of T7 gp4). This makes the preassembly of the helicase on the DNA substrate, without reaction occurring during the assembly period, challenging. Based on our finding that T7 gp4 forms hexamers and binds DNA in the absence of Mg(II), we arrived at the following assembly procedure (this might be applicable to other helicases that require the presence of NTP to bind DNA). We preassemble the helicase on the DNA by adding dTTP, but by leaving out Mg(II). In the absence of Mg(II) and in the presence of added EDTA (to chelate contaminating divalent metal ions), T7 gp4 does not hydrolyze dTTP or unwind DNA (32). 1.1.1.2. DNA Substrates and DNA Unwinding Kinetics

Two types of the unwinding assays are described: discontinuous gel-based radiometric assay and continuous stopped-flow fluorescence assay. They are both all-or-none unwinding assays where unwinding rates are obtained from the kinetics of end product formation, i.e., the kinetics of the appearance of the fully unwound DNA. Since the unwinding rate of T7 gp4 is fast, the kinetics are measured using a rapid quenched-flow or a stopped-flow apparatus (Fig. 5.1) that allow mixing in the millisecond time scales. A typical DNA unwinding substrate for the hexameric helicase consists of a fork DNA. Two short DNA strands (top and bottom) are annealed to generate a duplex region (40 bp, here) and ssDNA overhangs at one end. A 50 ssDNA overhang (dT35) in the top strand is needed for helicase binding (the DnaB family helicases that are 5’ to 3’ helicases), and the 30 ssDNA overhang (dT15) in the bottom strand is required for strand exclusion during unwinding (Fig. 5.2). In the gel-based assay, one of the DNA strands is radiolabeled, so the radioactive fork substrate and the ssDNA product can be quantified after they are resolved by native PAGE. The kinetics of DNA unwinding is fit to obtain the unwinding rate. In the fluorescence-based assay, a fluorescent dye (fluorescein) is incorporated at the 50 end of the bottom strand (Fig. 5.3). A run of three guanosines at the 30 end of the top strand quenches fluorescein fluorescence when the substrate is duplexed. When helicase unwinds the dsDNA and the top strand is displaced away from the dye, the fluorescence increases. The time dependent increase in fluorescence is measured continuously in a stoppedflow apparatus.

1.1.2. Polymerization Kinetics

DNA polymerization rates can be estimated from the individual nucleotide incorporation rates measured using transient state kinetic methods. DNA polymerases incorporate hundreds to thousands of nucleotides during polymerization in a templatedependent manner, adding one dNMP to the primer at a time and moving with a step-size of one nucleotide. Each nucleotide is

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Fig. 5.1. Instrumental designs for the rapid kinetic studies. (a) Chemical quenched-flow RQF-3 (www.kintek– corp.com). Sample A and sample B are loaded in sample loops from the load ports via a three-way valve. Upon firing the instrument, solution A and B are forced through the delay line by water from the drive syringes A and B, and then reactants are mixed in the valve to start the reaction. The reaction mix flows through the selected delay line and mixed with the quench solution from syringe C after predetermined time intervals. The quenched sample is collected into a tube from the exit line. Different length delay lines are selected through an eight-way valve for reaction times in the range of 2– 100 ms by selecting different reaction loops. (b) Stopped-flow instrument. The KinTek stopped flow has a stable light source and sensitive detection system that can measure absorbance and fluorescence simultaneously. There are two channels for fluorescence detection, and three drive syringes, but in normal mode of operation two syringes are connected (those shown) and used to drive mixing of two reactants A and B into the observation cell. Reaction time is under computer control allowing times from few milliseconds to several minutes. The instrument dead-time is determined as outlined in www.kintek–corp.com.

added at a different rate that depends on several factors, not all well understood, one of which is the sequence context around the base to be added. The nucleotide addition rate can be determined accurately using a combination of rapid kinetics, product analysis on a high resolution sequencing gel, and data analysis. The rapid kinetic methods providing milliseconds time resolution capture the formation and decay of intermediate products, the sequencing gels can resolve the DNA products with a single-base resolution, and data analysis extracts the single nucleotide incorporation rate and the polymerase off-rates from the observed kinetics of primer elongation. 1.1.2.1. DNA Synthesis Kinetics

DNA polymerase extends a primer annealed to a template DNA by utilizing dNTPs as substrates. When the template is single stranded, the polymerase can copy the template without the helicase, but when the template is double stranded, the polymerase

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Fig. 5.2. Gel-based radiometric assay for DNA unwinding. (a) The DNA unwinding fork substrate design with radiolabeled top strand. T7 gp4 assembles on the top strand and moves in the 5’ to 3’ direction to unwind the dsDNA substrate. (b) Representative native gel showing the ds and ss DNA resolved as a function of reaction time. Time points here are 0, 0.05, 0.08, 0.1, 0.15, 0.2, 0.3, 0.4, 0.5, 0.6, 0.7, 0.8, 1, 1.2, 1.5, 1.7, 2, 2.5, 3, 4, 6, 8, 15, 30 s (for 40 ds duplex). (c) Kinetics of ds40 unwinding at 18C in reactions containing T7 gp4, dTTP, MgCl2, and SSB as the trap. The kinetics is fit to the gfit unwinding model (unwinding.m), which provides kf (stepping rate constant) and s (kinetic step size), from which the average rate of unwinding (kf  s) was determined. A good fit for the 40 ds DNA unwinding data here gave parameters; A (maximum DNA unwound) = 0.829, kf = 9.89, minD (minimum duplex region that spontaneously separates) = 0, s = 6.34, and F0 (background) = 0.002 and hence an average rate of unwinding 62.7 bp/s (see section 3.6.2).

requires the helicase to unwind the dsDNA. T7 DNA polymerase requires T7 gp4 to catalyze strand displacement DNA synthesis. The rate of DNA unwinding by the helicase with concomitant DNA synthesis can be measured by the unwinding assays described above using a replication fork substrate. Alternatively, the kinetics of the reaction can be measured by following the primer extension reaction. We describe methods to obtain the rate of each nucleotide addition in the primer extension reaction. Replication fork substrates are made by annealing the top and bottom ssDNAs to generate a duplex region (40 bp, here) and two ssDNA overhangs.

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Fig. 5.3. Fluorescence-based stopped-flow assay for DNA unwinding. (a) DNA unwinding fork substrate design with fluorescein in the lower strand and GGG at the 3’ end in the top strand. T7 gp4 moves in the 5’ to 3’ direction to unwind the dsDNA substrate. (b) Representative kinetic trace showing the unwinding of the fork DNA by the timedependent increase in fluorescence. The kinetics is fit to the gfit unwinding model (unwinding.m), which provides kf and s, from which the average rate of unwinding (kf  s) was calculated. A good fit for the 40 ds DNA unwinding data in the presence of top ssDNA trap was obtained at N (population) = 2 and hence gave parameters for the two phases;A1= 0.862, A2 = 0.137, kf 1 = 4.488, kf 2 = 1.413, minD = 0, s = 5.395, and F 0 = 0.995 and hence an average rate of unwinding for the fast population is 24.2 bp/s and for the slow population is 7.6 bp/s (see Fig. 5.2, legend and section 3.6.2).

A third strand, a primer (24-mer, here) is annealed to the bottom strand (to 25 nt 30 ssDNA overhang, a defined sequence) to create a primer/template junction (Fig. 5.4). The primer is either radiolabeled or fluorescein-labeled (at the 5’ end) to follow the primer extension kinetics. Reaction products are separated by high percentage sequencing PAGE with single-base resolution to measure the amount of each intermediate. 1.2. Analysis of Reaction Kinetics Measurements

Even the most advanced experimental techniques cannot fully decouple all steps involved in unwinding and polymerization process. Most experimental observations arise not from one but from multiple simultaneously occurring reactions. Such results do not directly provide a value for any reaction rate or even a confirmation that the reaction step actually takes place. This information can be extracted from the results by applying model-based (regression) analysis that has the following goals: (a) to test if the proposed

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Fig. 5.4. Strand displacement DNA synthesis by helicase-polymerase replisome. (a) T7 DNA polymerase and T7 gp4 are assembled on the replication fork substrate with a radiolabeled DNA primer. T7 gp4 on top strand moves to unwind the dsDNA and DNA polymerase extends the primer in the 5’ to 3’ direction by duplicating the bottom strand. Sequencing gel shows progressive strand displacement DNA synthesis activity of the replisome. Time points here are 0, 0.004, 0.006, 0.008, 0.01, 0.015, 0.02, 0.03, 0.04, 0.05, 0.06, 0.08, 0.1, 0.12, 0.15, 0.2, 0.25, 0.3, 0.4, 0.5, 0.6, 0.75, 1, 1.5, 2, 3, 4 s (for 41 nt base extensions). The 24-mer primer is elongated to 65 nt runoff product. (b) Rate constants of individual nucleotide addition plotted against nucleotide added (rates of first three nucleotide addition are not plotted here as they are always high probably due to premelting of duplex with helicase binding). Rate constants are estimated by fitting the kinetics of individual DNA product synthesis to the polymerization model of gfit (polymerase_ni.m). Errors are calculated from the global fits to the polymerization model.

model is in agreement with the observations, (b) to estimate parameters of the model, and (c) to estimate their confidence intervals. In this chapter we describe analysis of unwinding and polymerization data using gfit, an open source program (http:// gfit.sourceforge.net) (33). Although the current version of gfit runs within MATLAB environment and uses computational models written in MATLAB language programming, it is only involved in creating new models; all analysis tasks in gfit using

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existing models are performed through graphical user interface and do not require any knowledge of programming or MATLAB environment. In the following sections we discuss details of the analysis procedure, computational problems that it presents, and the ways to address these problems. 1.2.1. Steps Involved in Model-Based Analysis

Analysis starts with creating a computational model based on existing knowledge about the system. The model should be able to compute (simulate) a predicted result of each experiment. To perform computations, the model uses two kinds of inputs: experimental conditions, known values from the experimental protocol (e.g., incubation time, nucleotide concentration), and model parameters, usually unknown, intrinsic properties of the system (e.g., translocation step size, rate of nucleotide incorporation). Although, for any given experiment, conditions and parameters are easily distinguishable, the same variable (e.g., enzyme concentration) may appear as a condition in one experiment and as a parameter in another. Correctness of a model can never be demonstrated conclusively. However, given certain parameter values, a model may produce simulations closely matching (fitting) experimental measurements. Testing if the model is consistent with all the experimental observations, a key step in data analysis, is performed by global curve fitting (optimization). Failure to find a good set of parameter values usually means that the model is not accurate and requires structural changes. Finding a good fit not only demonstrates consistency, but also provides estimates of parameters. Such result does not conclude the analysis because the estimated parameter values carry little meaning without an indication of their uniqueness and an estimate of their confidence intervals. Restarting optimization many times from randomly selected positions in the parameter space may lead to discovery of alternative parameter sets fitting the data, an indication of low confidence. Underdetermined parameters may stem from an overly complicated model or from insufficient experimental data. The problem can be solved by simplifying the model, by adding explicit constraints to parameters (e.g., equating two related rate constants, sharing a parameter between multiple experiments), or by providing more experimental results. Global analysis of experiments that highlight different aspects of system’s behavior provides most rigorous validation of the model and allows estimation of its parameters with highest confidence.

1.2.2. Software for ModelBased Analysis

Although procedures for model-based data analysis are well established, their practical applications to biological systems present several computational challenges and places rigid requirements on the analysis software. The main function of the software is to provide methods for statistical analysis that operate on

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experimental data and computational models supplied by the researcher. Biological research projects are highly dynamic. Therefore the software should make it easy to modify the model, to include new experiments into global analysis, to change statistical weights, to use same parameter value for multiple experiments, etc. The desired flexibility can be achieved by developing a projectspecific analysis program (34), although our experience shows that this approach requires frequent modifications of the program code, which slow down the analysis and lead to programming errors. Computational models are the central parts of the analysis. The models have to simulate a wide range of biological processes and experimental procedures. Ordinary differential equations (ODEs) is currently the most common method to model biology. However, defining a model strictly as a set of ODEs is unacceptable because some systems have to be simulated with a floating number of ODEs (e.g., unwinding and polymerization, as discussed below), others by solving ODEs several times while changing initial concentrations to replicate pre-incubation, dilution, and mixing performed as part of the experimental procedure, yet others may require completely different computational techniques. Since simulation is usually the slowest step in the analysis process, it should be performed as efficiently as possible. An analysis procedure may involve 103 to 108 simulations with different experimental conditions and parameters. To carry out the analysis without human intervention, each output of the model has to be directly comparable with the result of the corresponding experiment. In short, simulations require accuracy, flexibility, and efficiency. These requirements can be best met by writing models in a general purpose programming language (e.g., C++, Python, MATLAB) making it possible to accurately capture all details of the mechanism and experimental design and using most efficient algorithms. Considering the above requirements, model-based analysis of experiments in this chapter was performed using gfit software that solves the general case of this problem (33). Since gfit uses MATLAB scripts as models, it can be used for analysis of any type of experimental data – kinetic, thermodynamic, or any other. Details about writing gfit models are beyond the scope of this chapter. Instead, we described how the existing models can be used for data analysis. Models and sample datasets used inthis chapter, more detailed documentation, and more modeling examples can be found at the gfit website: http:// gfit.sourceforge.net. 1.2.3. Data Analysis Using gfit

Experimental data in gfit is represented by a collection of experiments. Each experiment may include multiple variables, which store numerical information. A variable may contain a scalar (single

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number), a vector (a column of numbers), or a matrix (arrays with more than two dimensions are also supported, but do not appear in this chapter). gfit uses variables for communicating with models. Input variables are used for sending a model the data it needs to perform a simulation; simulation results are received from the model as output variables. Since every model has its own special requirements for input data and produces different types of results, the model has to describe its input and output variables, their names, dimensional relationships, and other properties. By reading model description, gfit learns how the model should be run and what kind of simulations to expect from it. Same information is used by gfit for importing experimental data. During import, the data is checked against the requirements of the currently selected model, to determine whether it is suitable for simulation and for fitting. If a critical piece of information is missing, or if data violates some of the model’s requirements, it is rejected. For example, it is an error not to include variable time in an unwinding experiment because unwinding.m model lists this variable as a requirement. It is also an error to include a vector of ten numbers as time and a vector of 11 numbers as F, because the model stipulates that the two variables should be vectors of equal length. Experimental data is imported to gfit from a spreadsheet by copying a block of data into clipboard. Many experiments arranged side-by-side to each other can be imported in one operation. Each experiment must have a name starting with a letter and containing any characters thereafter. All names of experiments should appear in the top row of the imported block. Variables for each experiment should occupy spreadsheet cells directly below or below to the right of experiment’s name. Names and other properties of variables should be according to model’s definitions. To obtain names of variables defined by the current model, select menu Model ! Copy data headers. This command places into the clipboard a sample header for one experiment. Paste contents of clipboard into an empty spreadsheet. Note that the header contains all variables defined by the model; not all of them have to appear in actual experimental data. Variable’s data occupy rectangular blocks of spreadsheet cells. The top left corner of each block should appear immediately below the variable’s name. The number for a scalar variable may also occupy the cell immediately to the right from its name. In that case, a colon (‘:’) should be added to the name. Experimental data is imported from clipboard by selecting menu Data ! Paste-add Data. If import is successful, gfit generates parameters required for simulation and fitting of imported experiments. For each parameter, the name, optimization flag, lower bound constraint, current value, and upper bound constraint are shown. To simulate experiments, gfit combines data

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from parameters and experimental conditions and sends it to the model. The way each parameter is used for simulation of each experiment can be defined by researcher by selecting Combine Parameters and Separate Elements from Parameters menu or after right-clicking parameter name. The same parameter can be used in one or in many experiments, in one or in many input variables, and, if the input variable is an array, in multiple positions of the array. Checked optimization flag indicates that the parameter should be optimized during fitting. During fitting, parameter value may vary between lower and upper bound constraints, which can be adjusted by researcher. 1.2.4. Computational Model for Unwinding

Unwinding can be modeled as a process involving multiple steps of equal size, s, and rate, kf. The only observable product, ssDNA, is produced at the last step (Figs. 5.2, 5.3). Its appearance is simulated by the model unwinding.m as an incomplete gamma function (30, 31, 35). This method is both computationally efficient and allows using continuous (not only integers) number of steps. To account for heterogeneity in helicase population, the model calculates the sum of N unwinding processes. Other parameters of the model that can be estimated by fitting include unwinding amplitude, A, background signal, F0, and minimal stable duplex length, minD. It should be noted that the estimated step size s is strongly affected by enzyme’s heterogeneity and thus maybe inaccurate.

1.2.5. Computational Model for Polymerization

Polymerization is another example of a multi-step process with the number of steps (Fig. 5.4), N–1, equal to the length of the DNA template. At each step, the polymerase can either add a nucleotide to the growing chain with rate kf or dissociate from the substrate, with rate kd. The process is modeled as N ODEs. Since intermediate polymerization species can be resolved on a gel, a vector concentration is measured for each time point. The result of the experiment is therefore a matrix of size M by N, where M is the number of time points. To avoid writing dedicated models for each template length in gfit, it is possible to write a universal model, polymerase_ni.m, that always simulates the correct number of steps according to the experimental data. Forward and dissociation rates at each step do not have to be the same. Therefore, for each simulation the model requires N–1long vectors for kf and kd. Accordingly, for each experiment gfit generates N–1 parameters for each of the variables. By default, these parameters are linked (forced to keep the same value) into single parameters kf and kd. It is a researcher’s option to either separate the parameters completely or to group them in any desired fashion. For example, one could try fitting the data while linking the kf-s according to the incorporated base type – kfG, kfT, kfA, kfC.

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2. Materials 2.1. Proteins

1. T7 gp4 (gp4A’) was purified as described (36). The molar concentration of the protein in 8 M guanidine hydrochloride is obtained by UV absorption (extinction coefficient at 280 nm 0.0836 mM–1 cm–1) (see Note 1). 2. T7 gp5 (D5A, D7A) was purified as described (37). The molar concentration of gp5 in 8 M guanidine hydrochloride was obtained by UV absorption (extinction coefficient at 280 nm of 0.13442 mM–1 cm–1) (see Note 1). 3. E. coli thioredoxin enzyme (Sigma-Aldrich). 4. If needed purify E. coli SSB as described by Lohman et al. (38) (see Note 1).

2.2. Instruments

1. Criterion cell (BioRad) for native PAGE. 2. Preparatory vertical gel slab electrophoresis (Hoefer Scientific Instruments SE-600 series) for DNA purification. 3. Elutrap electroeluter for DNA purification (Hoefer Scientific Instruments). 4. Sequencing gel unit (Sequi Gen GT, BioRad). 5. RQF3 rapid quenched-flow instrument from KinTek corporation (Fig. 5.1a). 6. SF-2004 Stopped-flow instrument KinTek corporation (Fig. 5.1b). 7. Typhoon scanner and Phosphorimager screens from molecular dynamics.

2.3. Software Requirements for Quantification of the Data

1. ImageQuant 5.0 or later required for quantitation of scanned data. 2. MATLAB v6.5 SP 1 or later required for running simulations. 3. MATLAB Optimization Toolbox required for regression analysis.

2.4. Reagents

1. Deionized water of 18.3 Ohm conductivity (Milli-Q water) for all the solution preparation and reactions. 2. Oligonucleotides from IDT (Coralville, IA) or any other source. 3. BT1, BT2 Elutrap membranes for Elutrap electroeluter (Whatman). 4. Snake venom phoshodiesterase I, DNase I, 8 M Guanidine hydrochloride, BSA can be purchased from Sigma-Aldrich.

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5. 5’and 6’ carboxy-fluorescein succinimidyl ester (Molecular Probes). 6. Radiolabeled nucleotides (Perkin Elmer). 7. Sephadex G-25 spin columns for oligonucleotides (Roche). The gel filtration matrix can also be purchased and then packed in 1.5 ml tube spin columns available from BioRad. 8. Make Deoxy and ribo nucleotides (Sigma-Aldrich) solutions in Tris–HCl buffer and adjust pH to 7.5. Estimate concentration of the solution using respective molar extinction coefficients and store at –20C. 9. Polynucleotide kinase (New England Biolabs). 10. 0.5 mM EDTA (pH 8.0) stock solution, stored at room temperature. 11. 1 M Tris–HCl buffer, pH 7.6, stored at room temperature. 12. 1 M magnesium chloride stock solution, stored at room temperature. 13. 20% SDS stock solution, stored at room temperature. 14. 5 M sodium chloride stock solution, stored at room temperature. 15. 5 M sodium hydroxide stock solution, stored at room temperature. 16. Replication buffer: 50 mM Tris–HCl, pH 7.6, 40 mM NaCl, 10% glycerol. A 10 stock can be made and stored at room temperature in tightly capped tube. 17. Acrylamide /bis acrylamide mixture 40% (w/v) 19:1 cross linking (Amresco, Ultrapure grade), stored at 4C. 18. Loading dye 12 stock for gel-based unwinding assays: 0.25% (w/v) bromophenol blue, 40% sucrose, 12% (w/v) SDS. It is stored at 4C and brought to room temperature before use to allow SDS to dissolve back into the solution. 19. Loading dye for sequencing gels: 95% (v/v) formamide, 0.025% (w/v) bromophenol blue, 10 mM EDTA. It is stored at –20C. Avoid contact with formamide. 2.5. Sample Data and Models

A zip archive containing all models and data used in this chapter can be downloaded from http://gfit.sourceforge.net. The archive also includes readme.txt file with most current information. The data files are in tab/newline-delimited format. The files can be opened in a text editor, but it is more convenient to view them in a spreadsheet application.

3. Methods T7 gp4-catalyzed DNA unwinding activity requires a fork substrate (Figs. 5.2 and 5.3). When the activity of helicase and

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polymerase is measured, the unwinding substrate is further modified to anneal a primer to the bottom strand to create a primer/ template junction for DNA polymerase binding (Fig. 5.4). All the proteins are preassembled on the DNA substrate in the presence of dTTP, but in the absence of Mg(II). Reaction are started as ‘standing start’ by adding Mg(II) (see Note 7). The rapid quenched-flow methods for gel-based DNA unwinding assays (31) and strand displacement DNA synthesis for polymerase-helicase in replisome (6) are described here. A high-throughput stopped flow fluorescence-based DNA unwinding assay for helicase is also explained (39). 3.1. DNA Substrate Assembly for the Radiometric Assays

1. Purify the oligodeoxynucleotides by resolving them on preparatory 7 M urea-PAGE/TBE gel. 2. Identify the major DNA band by UV shadowing and excise it out with a clean blade (see Note 2). 3. Cut the gel band into 11mm pieces and elute the oligonucleotide from the gel pieces by electroelution on Elutrap electroeluter using BT1 and BT2 membranes and 1 TBE buffer (see Note 2). Carry out the electroelution at 120 V. BT2 membrane is permeable to the oligonucleotides but BT1 is not, so the eluted oligodeoxynucleotides is retained in the chamber between the two membranes at the positive electrode side. 4. Collect eluted fraction in a tube every 2–3 h, three to four times. Precipitate the DNA by ethanol/salt treatment followed by 70% ethanol wash to desalt the pellet. 5. Resuspend the air dried pellet in TE pH 8.0 buffer for storage. 6. Determine the oligodeoxynucleotide concentrations after digesting it with snake venom phosphodiesterase I using absorption at 260 nm and individual base extinction coefficient values at 260 nm (40, 41) (see Note 3). 7. Radiolabel the 5’ end of the top strand (for gel-based helicase unwinding assays) or primer (for DNA synthesis assays) with g 32 P-ATP and polynucleotide kinase at 37C for 30 min followed by incubation at 65C for 20 min to denature the PNK enzyme. 8. Clean the labeled DNA using Sephadex G-25 spin column (Roche Scientific). Check the purity of the labeled DNA by separating labeled oligonucleotide from the free label on PEI cellulose F TLC (Merck)/0.4 M potassium phosphate buffer (pH 3.4). Expose the TLC to the phosphorimager screen and scan the phosphorimage after an appropriate time. If the amount of free radiolabel is high, then do another round of clean up with Sephadex G-25 spin column (see Note 4).

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9. Assemble the DNA substrates by mixing equal molar amounts of the DNAs in replication buffer, incubate the mix at 95C for 10 min followed by slow cooling (2–3 h) to room temperature (see Note 5 and Fig. 5.2). 3.2. DNA Substrate Assembly for the Fluorescent Assays

1. Label the 5’ end of the bottom strand (for unwinding experiment) or of the primer (for DNA synthesis experiment) which has a 6-carbon linker terminated with an amino group, with 5’and 6’ carboxy-fluorescein succinimidyl ester (see Note 6). Carry out the labeling in carbonate buffer using the procedure from Molecular Probes. The fluorophore labeled oligodeoxynucleotides can also be purchased from IDT. 2. Purify the labeled oligodeoxynucleotides by resolving on a long preparatory 7 M Urea-PAGE/TBE gel followed by electroelution as described in Section 3.1 3. Anneal the DNA substrate by mixing the dye-labeled bottom strand with the complementary portion of the top strand with or with the primer as described in Section 3.1 (Fig. 5.3). 4. Purify the annealed substrate complex on long preparatory non-denaturating PAG followed by electroelution. 5. Determine the concentrations of the substrates after digestion with phosphodiesterase in combination with DNase I. Standardize the extinction coefficients of the unlabeled substrates with 0.1 M NaOH and use it to get the concentration of the labeled substrate (absorbance at 260 nm in 0.1 N NaOH) (42).

3.3. Gel-Based DNA Unwinding Assays for Hexameric Helicases

1. Set up the temperature to 18C (or desired temperature) in the circulatory cooling water bath attached to the rapid quenched-flow instrument. Consult the information brochure of the quenched-flow instrument (Fig. 5.1a.) available on Kintek corporation web site (www.kintek– corp.com) for operation details of the instrument. 2. Clean the two drive syringes and the quench syringe thoroughly with water. Fill up the two drive syringes with water avoiding any air bubble and the middle quench syringe with the 100 mM EDTA quench. Keep the top three valves in closed position for this procedure. 3. Wash the two sample loops by connecting 10 ml syringes filled with water in load ports A and B in ‘load’ mode while having the exit line connected to the vacuum. Rinse the sample loops with methanol the same way in the ‘load’ mode followed by connecting the exit line to a vacuum line, until dry.

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4. Rinse all the reaction loops (1–8) one by one with water in the ‘flush’ mode through the flush tubes, by changing the position of the reaction loop valve 1 through 8 and having the exit line connected to the vacuum. Rinse all the loops with methanol while having them connected to the vacuum until dry. 5. Turn on the motor and then the system control panel. Type ‘RUN’ after ‘–’ appears on the screen and then press enter. Type ‘0’ (no) for ‘USE CONSTANT QUENCH VOLUME’ and then again type ‘0’ (none) for ‘ENTER 2ND QUENCH DELAY (SEC)’. This mode of operation is preferred as the chances of experimental errors are less in this operation. Quench solution volumes added to the reaction vary when loops 1–6 are used and so an estimated volume of the quench solution has to be pre added in the sample collection tubes to ensure equal quench volume in all the samples. Instrument should be calibrated for the quench volumes of each loop and difference in the volumes should be known to work in this mode. 6. Select ‘ADJUST POSITION’ from the main menu followed by selecting ‘+’ for the downward movement and to bring down the ramp by pressing the start button on the motor. Lower the ramp to touch the top of the drive and quench syringe pistons. Similarly ‘–’ can be selected followed by pressing the start button to bring the ramp up if needed to readjust it or to refill the drive and quench syringes during the experiment. 7. Mark 1.5 ml reaction tubes according to the sample time points, and add required amount of 100 mM EDTA quench in the tube depending on the loop to be used for that sample. Put 23 ml of 12 loading dye for final 1% (w/v) SDS, 20% glycerol, and 0.25% (w/v) bromophenol blue in the reaction. Puncture the lids of the tubes to insert the exit line of the RQF during reaction for sample collection. 8. Select ‘QUENCH FLOW RUN’ from the main menu and type a time in seconds and press ‘enter’. Set the valves in the ‘fire’ mode; hold the 1.5 ml tube with the collection line inserted in the tube through the lid hole. Press the ‘G’ button to fire. Two to three of blank fires without reaction help force out the air bubbles from the tubings and loops. Wash and dry the loop in the ‘flush’ mode. 9. Make a solution with 5 nM radiolabeled fork DNA substrate, 200 nM hexameric amount of gp4, 2 mM dTTP, and 5 mM EDTA in replication buffer (see Note 7 and 8). Put solution in a 1-ml syringe avoiding any air bubbles and connect to sample loading port A. Total amount of solution made depends upon number of time points. Load this solution into the sample loop in the ‘load’ position.

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10. Prepare the same amount of the other solution with 2 mM dTTP, 8 mM free MgCl2 (accounting for EDTA and dNTPs), and appropriate traps if required (3 mM top strand ssDNA or 6 mM dT90 the protein traps or 3 mM SSB the DNA trap). Put solution in a 1-ml syringe and hook up to second sample loading port in the same way as above and load it into the sample loop in the ‘load’ position. 11. The two solutions are in the respective sample loops in equal volumes. Setup the knobs in the ‘fire’ mode and fire from the control panel as described above to rapidly mix 20 micro litre of the two solutions, from each syringe in the reaction loop and to initiate the reactions. The reaction is incubated according to the time set. Reactions are stopped by rapid mixing with 1.5-fold volume of a quenching solution of EDTA from the quench syringe. After sample collection, vortex and spin down the contents in the reaction tube. Wash the reaction loop with water followed by methanol to dry between each time point. Load 10–15 ml of the reaction on 10% native PAGE /1 TBE buffer, pH 8.3, with 0.2% SDS and resolve the products (see Note 9). 12. Peel off the gel from the plates and wrap in clear cling wrap. Expose the gels to the phosphorimager screen for appropriate times depending on the counts and then scan the screen in a phosphorimage scanner. 13. Quantify the bands in the scanned image using the ImageQuant software. Calculate the fraction unwound for each time point by using the equation: Fractionunwound ¼fðSS DS0 ÞðDS SS0 Þg=fDS0 ðDS þSSÞg where SS is the unwound single-stranded DNA count, DS is the double-stranded DNA substrate count, SS0 and DS0 are the respective counts for single-stranded and doublestranded DNA at 0 time. 14. Analyze the results as described in Section 3.6.2, to estimate the unwinding rate and other parameters (Fig. 5.2). 3.4. Real Time Fluorescence DNA Unwinding Assay for Hexameric Helicase

1. Set up the temperature at 18C (or desired temperature) in the circulatory cooling water bath of the SF-2004, stopped flow instrument (Fig. 5.1b). Consult the information brochure of the stopped flow instrument available on www.kintek –corp.com for details about the instrument (see Note 8). 2. Fill up the syringe chamber and the reaction cell with 2% (v/v) RBS 35 detergent (Merck) and incubate for 30 min for cleaning in the ‘load’ position of the valves.

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3. Rinse thoroughly with water to remove any traces of the detergent. Do this exercise by loading in the ‘load’ position and removing the solution out in the ‘fire’ position. Bring the valve to the ‘load’ position at the end. 4. Turn on the mercury lamp and allow it to stabilize to 75 V. 5. Set up the absorption wavelength to 480 nm. Insert a 515-nm cut-off long pass filter in the wavelength slot of the photomultiplier tube (PMT) path to be used. Adjust the slit width of light path to 2 mm (0.5 mm/turn, four turns of the slit adjustment knob). 6. Document the reactants in each syringe. Under ‘Instrument’, enter set time/channels and click OK. Set up the required parameters in the control panel; time 15 s (depending on the ds length to be unwound), PMT voltage in the range of 800– 900 V, wavelength 480 nm. 7. Use the replication buffer for a mock fire (fire as described below) to clean up the water traces from the reaction cell and the syringes. 8. Prepare a solution of 20 nM DNA substrate (you can use as low as 5 nM) with fluorescein label at the 50 end of the bottom strand, 200 nM gp4 (or desired concentration), 4 mM dTTP (or desired concentration), 6 mM EDTA, 2 mM BSA in replication buffer, and load in one of the loading syringe (see Note 7 and 8). Incubate the mix for 15 min at 18C. 9. Make a solution with free concentration of 8 mM MgCl2, 2 mM BSA, and 3 mM ssDNA or 3 mM E. coli SSB trap and load in the second loading syringe. Incubate the mix for 15 min at 18C. 10. Click on ‘Adjust syringe drive’ in the ‘Control’ menu to bring the ramp down to touch the drive syringes. 11. Ensure that the valve is in the ‘fire’ position and click on ‘Collect data’ button to fire the instrument for reaction to start. Upon firing, the instrument mixes the two solutions rapidly in the reaction chamber in equal volumes. The reaction mix is excited with 480 nm light and fluorescence is detected by a photomultiplier tube filtered through a 515 nm cut-off long pass filter. 12. Save all the traces and average the overlaying ones for further analysis. Go to ‘Data analysis’, ‘Select traces’, and delete abnormal traces. Go to File menu and save the data as ‘Stop flow data’ by clicking OK. Select ‘Data analysis’ and select ‘Average traces’ to get an average trace and save it.

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13. Save the data as an ASCII spreadsheet selecting for 1000 data points/trace. Copy the fluorescence data for each time point and paste in spreadsheet by clicking Data followed by choosing Text to columns, fixed width and save it in a spread sheet format. 14. Analyze the results as described in Section 3.6.2 to estimate unwinding rate and other parameters (Fig. 5.3). 3.5. DNA Synthesis Reactions on ds Template by the Helicase-Polymerase

1. The kinetics of strand displacement DNA synthesis by T7 helicase-polymerase replisome is measured in the rapid quenched-flow instrument at 18C (or desired temperature). Set up the instrument as described in Section 3.3. Prepare the sample collection tubes with required amounts of 300 mM EDTA quench according to the loops to be used. 2. Mix 50 mM E. coli thioredoxin with freshly made 5 mM DTT in replication buffer for 5 min at room temperature (22C). Assemble the T7 DNA polymerase by adding 10 mM of gp5 (in 1: 5 molar proportion to thioredoxin) in the mix and incubating it at room temperature for 5 min (37). Store the T7 polymerase on ice until use (2–3 h) (see Note 10). 3. Assemble 400 nM T7 gp4 hexamer on 200 nM fork DNA substrate (with a radiolabeled or fluorescein-labeled primer annealed to the bottom strand) in the presence of 2 mM dTTP, 2 mM DTT, and 2 mM EDTA in the replication buffer on ice for 30 min. (can use less polymerase and fork substrate as long as their concentrations is above the Kd value.) 4. Add 400 nM T7 DNA polymerase to the T7 gp4-DNA mix and further incubate the solution at room temperature for 30 min. All the concentrations mentioned here for replisome assembly are the concentrations in the syringe. 5. Load the replisome–DNA complex solution in one of the sample loading syringe of the quenched-flow as described in Section 3.3. 6. Make a solution with 1 mM each of dATP, dCTP, and dGTP, MgCl2 (free concentration of 8 mM) and 6 mM dT90 trap (optional) in the replication buffer. Load this solution to the second sample loading syringe. 7. Start the reaction by rapidly mixing equal volumes of the two solutions upon firing the instrument, and quench after various intervals with 300 mM EDTA from the quench syringe. 8. Dilute reactions with the sequencing dye (1:2 proportion of sample to sequencing dye for 20% GC content 40 ds duplex) boil the samples for 10 min at 95C and load

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immediately on a 24% acrylamide/7 M urea sequencing gel with 1.5  TBE buffer and run for 14 h at constant 90 W. Load just the sequencing dye with xylene cyanol in an end lane and run the gel till the xylene cyanol dye migrates to the end of plate. The 24-mer primer runs above this dye in 20–23% gel (see Note 11). 9. Let the gel cool down to room temperature for 40 min. Carefully peel the gel from the plate on a cling wrap with the help of a wet, flat spatula. 10. Cover the radioactive gel with the cling wrap on both sides and then expose it to the phosphorimager screen for required time and then scan the screen in the phosphorimager mode in the scanner. The fluorescent gel is directly scanned (best without the cling wrap) in the scanner in fluorescence mode at 526 nm Short Pass filter, Green 532 nm, Voltage 600–1000 V (depending on the intensity of the label) at normal sensitivity to get the image. 11. Quantify all visible DNA bands in the image by ImageQuant software (Fig. 5.4). 12. Paste counts data for different length bands into a spreadsheet. Add up the counts of all the bands in each lane in the gel (for each time point) to get the total intensity. Calculate the fraction of each band by dividing its count by the total count in that lane on the gel. 13. Analyze the results as described in Section 3.6.3 to estimate the nucleotide addition rates and polymerase dissociation rates for each base addition (Fig. 5.4). 3.6. Data Analysis 3.6.1. Installation of gfit

gfit software can be downloaded from http:// gfit.sourceforge.net. This website also contains most upto-date installation instructions. 1. Unzip the downloaded archive to your hard disk. For this chapter we will assume location E:/. Folder E:/Mgfit will be created. 2. Start MATLAB. 3. Change MATLAB’s current directory to E:/Mgfit. 4. To start installation, type mgfit in MATLAB’s command line after >> and press Enter. 5. Respond ‘Yes’ to the query about adding E:/Mgfit to MATLAB’s path. 6. Restart MATLAB if requested. 7. After installation, the same command, mgfit, will bring up gfit user interface window.

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3.6.2. Analysis of Unwinding Data

1. Start gfit by typing mgfit in MATLAB command line. The gfit window will appear. 2. Click on Model ! Pick Model and select unwinding.m file (included in models and data archive) and click Open. The name of the model will appear in the model field and the status line will show empty data set. 3. Arrange experimental data in a spreadsheet as described in Section 1.2.3. If using book chapter’s sample data, open file unwinding_dataset.txt (included in models and data archive) in a spreadsheet application. If using your own data, use that file as an example. 4. To transfer the data to gfit, select the data and copy it into the clipboard. In gfit window, choose menu Data ! Paste-add Data. If data import is successful, gfit shows a table of parameters that will be used for simulation or fitting of imported experiments. View data by clicking button Plot. gfit window shows following parameters: amplitude (A), stepping rate (kf), step length (s), minimum duplex length (minD), background signal (F0), and number of enzyme populations (N). 5. Adjust starting values of parameters and optimization flags. If fluorescence data are used, set the flag for F0. If analyzing data for one DNA length, clear the flag for minD. 6. Click button Fit. During optimization, the iterations are shown in the main MATLAB window. If fitting is successful, the status line shows message fitting converged followed by the sum of squared deviations. Two extra columns appear in parameter table showing the optimized parameter values and their asymptotic confidence intervals (see Note 12 and 13). Goodness of fit may be improved by increasing the number of enzyme populations. 7. Export analysis results to spreadsheet. Select menu Analysis ! Copy Report and paste table of parameters into spreadsheet. To export simulation results, select menu Data ! Export to Matlab. This command creates structure DS in MATLAB’s workspace. Substructure DS.sim contains an array of experiments from which simulated variables can be accessed one-byone and copied. 8. To improve the quality of fit and to test whether discovered optimal parameter values are unique, perform global search by selecting menu Analysis ! Random Restart. To terminate the search, click the Cancel button (see Note 14).

3.6.3. Analysis of Polymerization Data

1. Start gfit by typing mgfit in MATLAB command line. gfit window will appear.

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2. Select polymerase_ni.m model by choosing menu Model ! Pick Model. The name of the model will appear in the model field. 3. Arrange experimental data in a spreadsheet as described in Section 1.2.3. If using book chapter’s sample data, open file polymerization_dataset.txt (included in models and data archive) in a spreadsheet application. If using your own data, use that file as an example. 4. To transfer the data to gfit, select the data and copy it into the clipboard. In gfit window, choose menu Data ! Paste-add Data. If data import is successful, gfit shows a table of parameters that will be used for simulation or fitting of imported experiments. View data by clicking button Plot.gfit window shows following parameters: base addition rate (kf), dissociation rate (koff), and starting concentration (C0). 5. Click button Fit. If a good fit cannot be found, polymerization may be occurring with non-uniform kf and koff rates. Allow each step to have a unique kf rate by right-clicking the parameter and selecting Parameters ! Separate elements option. 6. Search parameter space using Random Restart and export analysis results as described in previous section.

4. Notes 1. Check all the protein preparations for any nuclease activity to avoid anomalous results due to DNA substrate degradation. 2. BT1 membranes used for Elutrap are brittle and need extra care while handling. BT2 membranes need to be stored at 4C and be moist all the time. Reverse the polarity of electrodes for about 30 s before taking an elute fraction of the oligonucleotide to dislodge any DNA that is sticking to the membranes. Use fresh gloves and blades for handling each oligonucleotide while cutting out the band form the preparatory gel and during setting up the elution to prevent any intermixing and contamination due to handling. Do not overexpose the DNA band to UV light. 3. It is very important to get an accurate oligonucleotide concentrations for preparing a correctly annealed substrate for all the assays described here. 4. Sephadex G-25 spin column purification works well for all the oligonucleotides above 8 nt; however, the recovery and purity may vary depending on the secondary structure of the

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oligonucleotide. Other matrices like P-5, P-30 from BioRad, or Sephadex G-50 could also be tried if needed. Use a medium size particle grade matrix for making the spin column. 5. It is good to have the GC proportion of the duplex region evenly distributed through the entire duplex region to avoid getting biased rates due to localized high or low GC patches. During oligonucleotide design avoid having any primase recognition site 3’CTG (on the top oligonucleotide) to prevent any primase activity when not needed. Avoid freezing and thawing the annealed substrates to retain their proper form. It is best to make them fresh or if required store at them 4C. 6. Do not have Tris or any other competing amine group in the labeling reactions with the fluorescein dye for an efficient labeling of the amine group. The fluorescein-labeled DNA substrate should always be protected from light by using dark tubes for reactions and storage. Keep the stopped flow reaction chamber covered with an aluminum foil to avoid outside light exposure of the dye. 7. In all the assays described here, it is important to keep a basal level of EDTA in the protein assembly solution to chelate any contaminating magnesium ions if present in the buffers to prevent an uncontrolled start of the reaction. 8. All the buffer solutions used should be filtered through the 0.22 mm filters whenever possible to avoid blockage or bacterial growth in the RQF or stopped flow instruments. Avoid air bubbles in tubings, loops, and syringes in both the instruments when collecting the data. A vacuum line is required for the RQF instrument for efficient flushing and drying of the loops. Do not connect the RQF instrument exit line to the vacuum line when the valves are in the ‘load’ position (reactants in the sample loading syringe) or in the ‘fire’ mode during the experiment. Clean the RQF and stopped flow instrument tubes before and after the experiment to prevent any blockage/buildup in the system. Never fire either instruments while having the valves in the ‘load’ position. 9. The quenched DNA unwinding reactions should be loaded immediately on the gel; if possible, as the experiment is being carried out, to avoid re-annealing of the complementary strands especially in the experiments without a trap. 10. Use very fresh DTT solution for T7 DNA polymerase assembly. Store the assembled T7 DNA polymerase on ice until use (2–3 h). 11. The Sequencing gel should be made with high purity acrylamide solutions and reagents. The gel should be run at high power to attain temperatures around 45–50C to get high resolution of the bands. A gel prepared with wedge spacer (0.25–0.4 mm here) gives good resolution for the

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bands of 20- to 80-mer size range. Sequencing gel loading dye to sample ratio should be increased if the length or the GC content of the duplex region of the DNA substrate is higher than mentioned here. The xylene cyanol dye interferes with the fluorescein intensity measurements and should not be used with fluorescent samples. 12. The parameters of unwinding can be estimated more reliably by globally fitting the unwinding kinetics for substrates of different length and similar GC composition. Processivity is best determined from the gel-based assay by analyzing unwinding amplitude as a function of dsDNA length (31). 13. Fitting operation can be aborted at any time by clicking Cancel. If, for any reason, fitting does not complete successfully, the latest set of parameters appears in the right column. To continue optimization, copy new parameter values to the starting ones by clicking column header <
Acknowledgments We thank the Patel lab members for proofreading the chapter and testing the models. This work was supported by National Institute of Health grant (GM55310).

References 1. Lohman T. M. (1993) Helicase-catalyzed DNA unwinding. J. Biol. Chem. 268, 2269–2272. 2. Lohman T. M., Tomko E. J., and Wu C. G. (2008) Non-hexameric DNA helicases and

translocases: mechanisms and regulation. Nat. Rev. Mol. Cell Biol. 9, 391–401. 3. Patel S. S. and Donmez I. (2006) Mechanisms of helicases. J. Biol. Chem. 281, 18265–18268.

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4. Patel S. S. and Picha K. M. (2000) Structure and function of hexameric helicases. Annu. Rev. Biochem. 69, 651–697. 5. Donmez I., Rajagopal V., Jeong Y. J., and Patel S. S. (2007) Nucleic acid unwinding by hepatitis C virus and bacteriophage t7 helicases is sensitive to base pair stability. J. Biol. Chem. 282, 21116–21123. 6. Stano N. M., Jeong Y. J., Donmez I., Tummalapalli P., Levin M. K., and Patel S. S. (2005) DNA synthesis provides the driving force to accelerate DNA unwinding by a helicase. Nature 435, 370–373. 7. Egelman E. H., Yu X., Wild R., Hingorani M. M., and Patel S. S. (1995) Bacteriophage T7 helicase/primase proteins form rings around single-stranded DNA that suggest a general structure for hexameric helicases. Proc. Natl. Acad. Sci. USA 92, 3869–3873. 8. Singleton M. R., Sawaya M. R., Ellenberger T., and Wigley D. B. (2000) Crystal structure of T7 gene 4 ring helicase indicates a mechanism for sequential hydrolysis of nucleotides. Cell 101, 589–600. 9. Toth E. A., Li Y., Sawaya M. R., Cheng Y., and Ellenberger T. (2003) The crystal structure of the bifunctional primase-helicase of bacteriophage t7. Mol. Cell 12, 1113–1123. 10. Tabor S. and Richardson C. C. (1981) Template recognition sequence for RNA primer synthesis by gene 4 protein of bacteriophage T7. Proc. Natl. Acad. Sci. U.S.A. 78, 205–209. 11. Kim D. E., Narayan M., and Patel S. S. (2002) T7 DNA helicase: a molecular motor that processively and unidirectionally translocates along single-stranded DNA. J. Mol. Biol. 321, 807–819. 12. Rasnik I., Jeong Y. J., McKinney S. A., Rajagopal V., Patel S. S., and Ha T. (2008) Branch migration enzyme as a Brownian ratchet. EMBO J. 27, 1727–1735. 13. Ahnert P. and Patel S. S. (1997) Asymmetric interactions of hexameric bacteriophage T7 DNA helicase with the 5’- and 3’-tails of the forked DNA substrate. J. Biol. Chem. 272, 32267–32273. 14. Hacker K. J. and Johnson K. A. (1997) A hexameric helicase encircles one DNA strand and excludes the other during DNA unwinding. Biochemistry 36, 14080–14087. 15. Kaplan D. L., Davey M. J., and O’Donnell M. (2003) Mcm4,6,7 uses a ‘pump in ring’ mechanism to unwind DNA by steric exclusion and actively translocate along a duplex. J. Biol. Chem. 278, 49171–49182.

16. Kaplan D. L. (2000) The 3’-tail of a forkedduplex sterically determines whether one or two DNA strands pass through the central channel of a replication-fork helicase. J. Mol. Biol. 301, 285–299. 17. Jezewska M. J., Rajendran S., Bujalowska D., and Bujalowski W. (1998) Does singlestranded DNA pass through the inner channel of the protein hexamer in the complex with the Escherichia coli DnaB Helicase? Fluorescence energy transfer studies. J. Biol. Chem. 273, 10515–10529. 18. Doublie S., Tabor S., Long A. M., Richardson C. C., and Ellenberger T. (1998) Crystal structure of a bacteriophage T7 DNA replication complex at 2.2 A resolution. Nature 391, 251–258. 19. Modrich P. and Richardson C. C. (1975) Bacteriophage T7 Deoxyribonucleic acid replication in vitro. A protein of Escherichia coli required for bacteriophage T7 DNA polymerase activity. J. Biol. Chem. 250, 5508–5514. 20. Tabor S., Huber H. E., and Richardson C. C. (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. 21. Ha T., Rasnik I., Cheng W., Babcock H. P., Gauss G. H., Lohman T. M., and Chu S. (2002) Initiation and re-initiation of DNA unwinding by the Escherichia coli Rep helicase. Nature 419, 638–641. 22. Dessinges M. N., Lionnet T., Xi X. G., Bensimon D., and Croquette V. (2004) Singlemolecule assay reveals strand switching and enhanced processivity of UvrD. Proc. Natl. Acad. Sci. U.S.A. 101, 6439–6444. 23. Dumont S., Cheng W., Serebrov V., Beran R. K., Tinoco I., Jr., Pyle A. M., and Bustamante C. (2006) RNA translocation and unwinding mechanism of HCV NS3 helicase and its coordination by ATP. Nature 439, 105–108. 24. Lee J. B., Hite R. K., Hamdan S. M., Xie X. S., Richardson C. C., and van Oijen A. M. (2006) DNA primase acts as a molecular brake in DNA replication. Nature 439, 621–624. 25. van Oijen A. M. (2007) Single-molecule studies of complex systems: the replisome. Mol. Biosyst. 3, 117–125. 26. Johnson D. S., Bai L., Smith B. Y., Patel S. S., and Wang M. D. (2007) Single-molecule studies reveal dynamics of DNA unwinding by the ring-shaped T7 helicase. Cell 129, 1299–1309.

Experimental and Computational Analysis 27. van Oijen A. M. (2008) Cutting the forest to see a single tree? Nat. Chem. Biol. 4, 440–443. 28. Tanner N. A., Hamdan S. M., Jergic S., Schaeffer P. M., Dixon N. E., and van Oijen A. M. (2008) Single-molecule studies of fork dynamics in Escherichia coli DNA replication. Nat. Struct. Mol. Biol. 15, 170–176. 29. Lionnet T., Spiering M. M., Benkovic S. J., Bensimon D., and Croquette V. (2007) Real-time observation of bacteriophage T4 gp41 helicase reveals an unwinding mechanism. Proc. Natl. Acad. Sci. U.S.A. 104, 19790–19795. 30. Ali J. A. and Lohman T. M. (1997) Kinetic measurement of the step size of DNA unwinding by Escherichia coli UvrD helicase. Science 275, 377–380. 31. Jeong Y. J., Levin M. K., and Patel S. S. (2004) The DNA-unwinding mechanism of the ring helicase of bacteriophage T7. Proc. Natl. Acad. Sci. U.S.A. 101, 7264–7269. 32. Picha K. M. and Patel S. S. (1998) Bacteriophage T7 DNA helicase binds dTTP, forms hexamers, and binds DNA in the absence of Mg2+. The presence of dTTP is sufficient for hexamer formation and DNA binding. J. Biol. Chem. 273, 27315–27319. 33. Levin M. K., Hingorani M. H., Holmes R. M., Patel S. S. and Carson J. H. (2009) Model-based global analysis of heterogeneous experimental data using gfit. Methods Mol. Biol. 500, 335–359, Humana Press Inc. 34. Patel S. S., Bandwar R. P., and Levin M. K. (2002) Transient-state kinetics and computational analysis of transcription initiation. The practical approach series/Kinetic analysis of macromolecules (Johnson K. A., Ed.), Oxford University Press, Oxford.

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35. Lucius A. L., Maluf N. K., Fischer C. J., and Lohman T. M. (2003) General methods for analysis of sequential ‘n-step’ kinetic mechanisms: application to single turnover kinetics of helicase-catalyzed DNA unwinding. Biophys. J. 85, 2224–2239. 36. Patel S. S., Rosenberg A. H., Studier F. W., and Johnson K. A. (1992) Large scale purification and biochemical characterization of T7 primase/helicase proteins. Evidence for homodimer and heterodimer formation. J. Biol. Chem. 267, 15013–15021. 37. Patel S. S., Wong I., and Johnson K. A. (1991) Pre-steady-state kinetic analysis of processive DNA replication including complete characterization of an exonucleasedeficient mutant. Biochemistry 30, 511–525. 38. Lohman T. M., Green J. M., and Beyer R. S. (1986) Large-scale overproduction and rapid purification of the Escherichia coli ssb gene product. Expression of the ssb gene under lambda PL control. Biochemistry 25, 21–25. 39. Donmez I. and Patel S. S. (2008) Coupling of DNA unwinding to nucleotide hydrolysis in a ring-shaped helicase. EMBO J. 27, 1718–1726. 40. Cavaluzzi M. J. and Borer P. N. (2004) Revised UV extinction coefficients for nucleoside-5’-monophosphates and unpaired DNA and RNA. Nucleic Acids Res. 32, e13. 41. Kallansrud G. and Ward B. (1996) A comparison of measured and calculated singleand double-stranded oligodeoxynucleotide extinction coefficients. Anal. Biochem. 236, 134–138. 42. Sjoback R., Nygren J., and Kubista M. (1995) Absorption and fluorescence properties of fluorescein. Spectrochim. Acta. [A] 51, 7–21.

Chapter 6 Protein Displacement by Helicases Laxmi Yeruva and Kevin D. Raney Abstract Helicases are ubiquitous enzymes that are vital to all living organisms. They are motor proteins that move in a specific direction along the nucleic acid and unwind the nucleic acid (DNA and RNA). ATP hydrolysis provides energy for helicase translocation and unwinding. The unwinding process provides ssDNA intermediates necessary for replication, recombination, and repair. Mutations in specific DNA helicases can lead to disruption in DNA metabolism. For example, mutations in helicases genes resulted in diseases such as xeroderma pigmentosum, cockayne’s syndrome, Bloom’s syndrome, and Werner’s syndrome. During unwinding, helicases are most likely to encounter proteins while moving along the nucleic acid. Several different research groups have demonstrated that helicases shift or displace proteins from one nucleic acid-bound location to another. These protein–protein collisions could result in displacement of proteins from nucleic acid or dissociation of helicase from nucleic acid. This report describes several different methods developed to study protein displacement by DNA and RNA helicases. Key words: DNA helicase, RNA helicase, FRET.

1. Introduction Helicases are motor proteins that are involved in DNA and RNA metabolism, replication, recombination, transcription, and repair (1– 4). Helicases convert chemical energy of nucleoside triphosphate (NTP) hydrolysis to mechanical energy to separate/unwind the strands of duplex DNA (2, 5, 6). In addition to unwinding, helicases also play a major role in disruption of protein–nucleic acid interactions (7–13). These protein–protein interactions can lead to displacement of nucleic acid binding protein, or helicase dissociation from the DNA, or temporary halt of the motor. The biological relevance of removing proteins from DNA by helicases or other enzymes is gaining researchers’ attention. For example, UvrD and Srs2 helicases have M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_6, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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been shown to remove proteins from DNA while DExH/D helicases have been demonstrated to disrupt RNA–protein interactions (13–15). NPH-II (DExH protein) is an RNA helicase from vaccinia virus that unwinds DNA in 30 to 50 direction. Studies demonstrated that NPH-II displaces protein U1A from an RNA substrate (16). Displacement of U1A is important because it is involved in spliceosomal machinery and it controls its own gene expression by feedback regulation. Chromatin remodelers contain regions of homology to helicases and disrupt DNA histone interactions (17, 18). For example, Mot1, an essential transcriptional regulator in yeast appears to displace TATA-binding protein (TBP) from DNA (19). Chromatin remodelers such as Snf2-like ATPases also have non-transcriptional functions such as homologous recombination (RAD54), transcription-coupled DNA repair (ERCC6.CSB), and histone deacetylation (Mi-2/ CHD3/CHD4). DNA repair protein (Mfd) reportedly pushes RNA polymerase from a DNA damage site or it completely displaces RNA polymerase allowing DNA repair to occur (20). From all these observations it is clear that protein displacement studies are gaining interest in the field. Four different methods that have been developed to study protein displacement by helicases are described below. 1.1. Helicases Displace Protein (Streptavidin) from Biotinylated Oligonucleotides

Hepatitis C virus (HCV) non-structural protein 3 (NS3) helicase is a SF2 helicase. The N-terminal 180 amino acids contribute to protease and C-terminal 450 amino acids possess helicase activities. NS3 facilitates DNA unwinding in a 30 to 50 direction by a 30 single-stranded DNA (ssDNA) attached to the duplex. Protein displacement studies indicated ATP-dependent unidirectional translocation of NS3 along the DNA (9). Method developed to study streptavidin displacement by NS3 is described in Section 3.1.

1.2. Displacement and Unwinding of Trp Repressor by Dda Helicase

Dda (DNA dependent ATPase) is a T4 bacteriophage helicase and has been shown to be active as a monomer for unwinding DNA and displacing streptavidin from 30 -biotinylated ss oligonucleotides. Method developed by Byrd et al. allows studying the DNA unwinding with protein displacement and is described in Section 3.2 (11).

1.3. PcrA Displaces RecA from ssDNA

RecA plays major role in genetic recombination. RecA polymerizes on the DNA in a 50 to 30 direction and is active as nucleoprotein filament with ATP and DNA. PcrA can affect RecA function by displacing RecA from DNA, by binding directly to RecA, or by competing for DNA binding. PcrA is a conserved DNA helicase and its absence shows high levels of recombination. Anand et al. have demonstrated that PcrA displaces RecA from ssDNA and dsDNA (12). Mutation of PcrA showed that helicase activity of PcrA is not essential for RecA displacement. Two methods were developed to study RecA displacement from ssDNA and dsDNA, which are described in methods Section 3.3 (12).

Protein Displacement by Helicases

1.4. Protein Displacement by DExH/ D RNA Helicases Without Duplex Unwinding

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DExH/D proteins are involved in all phases of RNA metabolism and these are often referred as RNA helicases. However, DExH/D proteins are likely to encounter RNA–protein complexes rather than pure RNA duplexes. Margaret et al. demonstrated that two RNA helicases readily remodel ribonucleo-protein complexes (13). Two methods were developed to study protein displacement by DExH/D RNA helicases and are detailed in Section 3.4.

2. Materials 2.1. Streptavidin Displacement Assay

1. Poly (dT) (Amersham Pharmacia Biotech). 2. Streptavidin, phosphoenol pyruvate (tricycle hexylammonium salt), ATP (disodium salt), pyruvate kinase/lactate dehydrogenase (in glycerol), bovine serum albumin, and sephadex G-25 (Sigma). 3. [g32-P]ATP (PerkinElmer Life Sciences) (see Note 7). 4. T4 polynucleotide kinase (New England Biolabs). 5. 30 or 50 biotinylated oligonucleotides (Integrated DNA technologies, IDT). Different lengths and sequence of substrates labeled with biotin either 30 or 50 were used for streptavidin displacement assay. Example sequences: 30 -bio-62mer (61 nucleotides plus biotin) 50 -TAACGTATTCAAGATACCTCGTACTCTGTAC TGACTGCGATCCGACGTCCTGCATGATGXT-30 or 50 bio-30mer (29 nucleotides plus biotin) 5-GXACGTA TTCAAGATACCTCGTACTCTGTA-30 or 50 bio-30mer (29 thymidines (T) plus biotin) or 50 -bio-60mer (59 T’s plus biotin). 6. Prepare Streptavidin stock (50 mM) in buffer containing 25 mM HEPES, pH 7.4, 10% glycerol, and 10 mM NaCl. Freeze down the protein in liquid nitrogen at 4C and store at –80C (see Notes 4 and 5). Dilute streptavidin to 5 mM prior to the experiment in buffer consisting of 25 mM HEPES, pH 7.5, 0.1 mg/ml BSA, 0.1 mM Disodium ethylenediamine tetraacetate (EDTA), 1 mM b-mercaptoethanol (BME). 7. Reaction buffer: 25 mM HEPES, pH 7.5, or 25 mM MOPS, pH 7 (see Note 8), 12.5 mM Mg(OAc)2, 150 mM KOAc, 4 mM phosphoenol pyruvate (PEP), 1 mM b-mercaptoethanol (BME), and 0.1 mg/ml bovine serum albumin (BSA). Make the reaction buffer fresh just prior to use. 8. 100 mM biotin stock, store at –20C. 9. Radiolabeled DNA at 50 end with [r–32P] ATP (see Note 7). 10. Quenching solution: 0.6% sodium dodecyl sulfate (SDS) (see Note 6), 200 mM EDTA, 10 mM poly (dT), 0.08% xylene cyanol, 0.08% bromophenol blue, and 10% glycerol.

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11. 10  TBE: 890 mM Tris, 890 mM Boric acid, and 20 mM EDTA. Store at room temperature. Dilute 10  TBE to 1  prior to use. 12. Acrylamide/bis acrylamide solution: 30% of acrylamide and 8% of N0 N0 methylene bisacrylamide in 1  TBE and store at 4C in a brown bottle (see Note 1). 13. 10% ammonium per sulfate (APS) prepared fresh prior to use. 14. NS3 was recombinant expressed and purified as described before (21, 22). 2.2. DNA Unwinding and Protein Displacement Assay

1. Poly(dT) (Amersham Pharmacia Biotech), BSA, L-Trp, and Sephadex G-25 from Sigma (St. Louis, MO). 2. ATP, HEPES, KOAc, BME, and EDTA from Fisher (Houston, TX). 3. [g32-P]ATP and T4 polynucleotide kinase. 4. 30 bp duplex substrate. The first 18 nt are palindromic sequence of the Escherichia coli trpEDCBA operator and the remaining 12 nt are a random, non-palindromic sequence. The 50 -single-stranded overhang consisted of varying number of thymidines (8, 12, and 24). (T)n GTACTAGTTAACTAGTACCGCTGATGTCGC – Loading strand CATGATCAATTGATCATGGCGACTACAGCG – Displaced strand 5. Quench solution: 200 mM EDTA. 6. Loading solution: 0.1% xylene cyanol, 0.1% bromophenol blue, and 10% glycerol.

2.3. Protein Displacement by PcrA Helicase

1. Streptavidin magnetic particles (Stratagene, Cedar Creek, TX). 2. The sequences of oligonucleotides are 50 C/BiodT/T GGC GAC CGC AGC GAG GC (dT)20 30 and 50 (dT)20 GGC GAC GGC AGC GAG GC/BiodT/C 30 . BiodT is biotinylated dT. 3. Sodium dodecyl sulfate, phosphoenol pyruvate (tricycle hexylammonium salt), ATP (disodium salt), pyruvate kinase/lactate dehydrogenase, TEMED, Streptavidin, Glucose, b-mercaptoethanol, Glucose oxidase, and catalase (Sigma). A higher stock of ATP can be prepared and stored at –20C. 4. RecA buffer: 20 mM Tris acetate, pH 7.5, 1 mM DTT, 10 mM magnesium acetate. 5. Separating buffer (4  ): 1.5 M Tris-HCl, pH 8.7, 0.4% SDS (see Note 6). Store at room temperature.

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6. Stacking buffer (4  ): 0.5 M Tris-HCL, pH 6.8, 0.4% SDS. Store at room temperature. 7. 30% acrylamide/bis solution (see Note 1). 8. 10% ammonium persulfate (APS). 9. Water-saturated isobutanol: mix equal volumes of water and isobutanol in a glass bottle, allow to separate, and use the top layer. Store at room temperature. 10. Running buffer (5  ): 125 mM Tris, 960 mM glycine, 0.5% (w/v) SDS. Store at room temperature. 11. Prestained molecular weight markers (Bio-Rad, Hercules, CA). 12. Commassie stain: 0.24% comassie blue, 60% methanol, 12% of acetic acid. 13. Destaining solution: 10% methanol and 10% of acetic acid. 14. Sample loading buffer: 62.5 mM Tris, 10% Glycerol, 5% bME, 1.05% SDS (see Note 6), and 0.004% bromophenol blue. 2.4. Protein Displacement by NPHII or DEDI

1. Oligonucleotides. 2. Tris, Nonidet P40, sodium chloride, magnesium chloride, and tryptophan from Sigma (St. Louis, MO). 3. 8% non-denaturing gel: 8 M urea and 8% polyacrylamide. 4. T4 DNA ligase (New England Biolabs).

3. Methods 3.1. Streptavidin Displacement by NS3

1. Purify the oligos by denaturing 20% polyacrylamide gel electrophoresis (PAGE). 2. Electroelute the DNA from the gel with an Elutrap apparatus. 3. Desalt the DNA with Waters Sep-Pak column and dry via Speed-Vac (Savant). 4. Resuspend oligos in 10 mM HEPES, pH 7.5, and 1 mM EDTA. 5. Determine the concentration by using the absorbance at 260 nm in 0.2 M KOH and calculated extinction coefficients. 6. Radiolabel the DNA at 50 end with T4 polynucleotide kinase at 37C for 1 h (see Note 7). 7. Inactivate (Denature) the kinase by incubating at 72C in a water bath for 10 min. 8. Remove the unincorporated [r-32p] ATP by passing the labeled oligos twice through a Sephadex G-25 spin column.

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9. Store labeled oligos at –20C and these can be used for 1 week. 10. Incubate 50 -radiolabeled oligos in reaction buffer (25 mM HEPES, pH 7.5, or 25 mM MOPS, pH 7, 12.5 mM Mg(OAc)2, 150 mM KOAc, 4 mM PEP, 1 mM BME, and 0.1 mg/ml BSA) with 300 nM streptavidin, 5 mM ATP, and PK/LDH (10.8 and 16.6 units/ml) at 37C for 3 min. 11. Following incubation add 6 mM biotin and initiate streptavidin displacement assay by the addition of helicase (see Notes 4 and 5). 12. Reaction can also be carried out by incubating helicase with oligonucleotide and reaction will be initiated by the addition of ATP. 13. At various time intervals collect 10 ml aliquots and mix with 10 ml of quench solution. 14. Analyze samples by electrophoresis on a 15% native polyacrylamide gel (PAGE) in 1  TBE. 15. Determine the quantity of radioactivity in streptavidin bound oligonucleotide and free oligonucleotide in each sample by using Molecular Dynamics 445-SI Phorphorimager with Image Quant software. 16. The fraction of free oligonucleotide, with a correction for the free oligonucleotide in the blank sample, can be determined by using the following formula. 17. FDc,t is the fraction of free oligonucleotide corrected for the amount of free oligonucleotide in the blank sample.FDt is the radioactivity of free oligonucleotide DNA for each sample at time t. SD t is the radioactivity of streptavidin-bound oligonucleotide DNA at time t. FDb and SDb are the radioactivity of free oligonucleotide DNA and streptavidin bound oligonucleotide DNA, respectively, in the blank (b) at time zero. Schematic of method is shown in Fig. 6.1. An example of results produced is shown in Fig. 6.2.  FDc;t ¼

3.2. Streptavidin Displacement and DNA Unwinding by Dda

FDt FDt þSDt

1

 





FDb FDb þSDb

FDb FDb þSDb





1. DNA duplexes and oligonucleotides were gel purified by PAGE (described in Section 3.1) and concentrations were determined by using the absorbance at 260 nm in 0.2 M KOH and calculated extinction coefficients. 2. The loading strand was radiolabeled at the 50 end (described in Section 3.1). The re-annealing trap was 12 nt complimentary to the non-palindromic portion of the displaced strand.

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Fig. 6.1. Illustration of experimental protocol of helicase-mediated streptavidin displacement from biotin oligonucleotide. Biotin oligonucleotide was preincubated with ATP in HEPES (pH 7.5) or MOPS (7.0), helicase was added with free biotin. At various time, aliquots were collected and quenched by the addition of SDS and EDTA. Free biotin prevents any displaced streptavidin from rebinding to the DNA. Samples were electrophoresed on 15% polyacrylamide gel. A representative gel picture is shown in Fig. 6.2.

Fig. 6.2. Results represent NS3-catalyzed displacement of streptavidin from 50 -bio60mer oligonucleotides. NS3 (500 nM) was incubated with 10 nM of substrate in a reaction buffer. Samples were collected at various time points and quenched. Streptavidin-bound oligonucleotide was separated from free oligonucleotide on 15% polyacrylamide gel. (Results are reproduced from ref. 9 with permission from ACS.)

3. Displacement of DNA binding protein trp repressor by Dda was carried out at 25C using a Kintek rapid chemical quenchflow instrument. 4. Incubate Dda (750 nM or 1mM, see Notes 4 and 5) with 32plabeled DNA substrate (100 nM) in reaction buffer (25 mM HEPES, pH 7.5, 10 mM KOAc, 0.1 mM EDTA, 2 mM BME, 0.1 mg/ml BSA, and 0.5 mM L-trp). 5. Initiate the reaction by adding ATP (5 mM), Mg(OAc)2 (10 mM), 75 uM (in nucleotides) poly (dT), and 25-fold excess of re-annealing trap. 6. Quench the reaction at various times with 200 mM EDTA (conc. before mixing). 7. Heat control sample to 95C for 10 min to test the efficiency of annealing trap. 8. Load samples on a 20% native polyacrylamide gel containing 10 mM TrisCl, pH 7.5, and 0.1 mM L-Trp and run at 250 V for 2 h in a buffer containing 10 mM TrisCl, pH 7.5, and 0.1 mM L-Trp with a circulating water bath maintaining the temperature at 4C.

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9. Determine the fraction of protein-bound dsDNA, free dsDNA, and ssDNA (unwound DNA after protein displacement) in each sample by using Molecular Dynamics 445-SI PhosphorImager and Image Quant software. Schematic of the method is shown in Fig. 6.3. An example result is shown in Fig. 6.4.

Fig. 6.3. Illustration of Dda catalyzed protein-displacement reaction. Dda in the presence of ATP and Mg2+displaces E. coli trp repressor from trpEDCBA operator and unwinds the duplex. Annealing trap prevents rebinding of the displaced strand to the loading strand. Helicase displaced Trp repressor protein only binds to the dsDNA. This assay allows one to measure unwinding and protein displacement simultaneously.

Fig. 6.4. Results represent Dda displacement of trp repressor and unwinding of DNA. One hundred nanomoles of substrate (24 nt overhang) was incubated with 750 nM of Dda and samples were collected at various time points. Samples were quenched immediately as described in methods section followed by 20% polyacrylamide gel electrophoresis. (Reproduced from ref. 11 with permission from Oxford Journals).

. 3.3. RecA Displacement by PcrA

1. Equilibrate streptavidin magnetic particles (SM) in Rec A buffer and use 20 ml SM slurry per reaction.

3.3.1. Method 1

2. Add 1 ml of 20 pM of biotinylated oligonucleotides to SM in RecA buffer and incubate for 20 min at 37C. 3. Remove the unbound oligonucleotides by magnetic separation and wash the beads in RecA buffer supplemented with ATP regenerating system (ATP (5 mM), pyruvate kinase, lactate dehydrogenase, and phosphoenol pyruvate).

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4. Add RecA protein (see Notes 4 and 5) to a final concentration of 6 mM to the SM slurry to a total volume of 20 ml and incubate the mixture at 37C for 5 min. 5. Separate the RecA bound to the oligonucleotides by magnetic capture of SM and elute the protein by 1  SDS sample buffer. 6. Analyze the eluted fractions and RecA released in the supernatant during the incubation by 10% SDS-PAGE and Coomassie blue staining. 7. To study RecA displacement by PcrA (or mutants) add 460 nM PcrA (helicase mutants PcrA3H– or PcrA3sau), to the RecA-bio-oligonucleotide complexes and incubate at 37C for various time periods. 8. Remove the RecA-bio-oligonucleotides by magnetic separation and collect the displaced RecA from the supernatant. 9. Analyze displaced and RecA bound to the oligonucleotides by SDS-PAGE and Coomassie blue staining. Schematic of the method is shown in Fig. 6.5.

Fig. 6.5. Schematic representation of RecA displacement from ssDNA by PcrA. Biotin oligonucleotides were coated with streptavidin magnetic beads. RecA was added to allow RecA-DNA bead complex formation. To the complex, PcrA was added and displacement of RecA was observed from supernatant fractions as explained in methods section.

3.3.2. Method 2

1. Design substrate (134 bp duplex with dT20 ss tail and 74 bp duplex with dT20 tail) containing four pairs of Cy3/Cy5 with a Forster distance [R0] of 11 nm to prevent FRET between the pairs.

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2. Coat slide surface with streptavidin and add biotinylated DNA substrates to 2 fmol/ml. 3. Wash unbound DNA, add RecA buffer containing oxygen scavenger (3% [wt/wt] glucose, 1% BME, 0.1 mg/ml glucose oxidase, and 0.02 mg/ml catalase), and measure fluorescence intensities for Cy3 and Cy5. 4. Add RecA at a concentration of 6 mM in a buffer containing ATP regenerating system (described previously). 5. Incubate the slide for 5 min in a water bath chamber to allow RecA binding and measure the fluorescence intensities of Cy3 and Cy5. 6. To study RecA displacement, add 500 nM of PcrA, and measure fluorescence intensities immediately. 7. The apparent efficiency of FRET (Eapp) is calculated by using IPLab software and the results are plotted using Origin and Excel. 8. Calculate mean values by Gaussian curve fitting of the Eapp values. 9. Separate the ssDNA containing dye pairs from dsDNA to analyze the Eapp value coming from the same molecule of DNA when it is in single-stranded form. Schematic of the method is shown in Fig. 6.6.

Fig. 6.6. Schematic representation of RecA displacement from dsDNA by PcrA in FRET based assay. Distance between FRET pair is 1  ¼ 4.4 nm. RecA binding lengthens dsDNA to 1.5-fold and decreases the FRET signal (Eapp). Addition of PcrA increases FRET signal (Eapp), indicating RecA displacement from dsDNA.

3.4. Protein Displacement by RNA Helicases 3.4.1. Method 1

1. RNA oligonucleotides are purchased from DHARMACON or transcribed from linearized DNA plasmids using T7 polymerase. 2. For TRAP (tryptophan RNA binding attenuation protein)RNP remodeling, 78 nucleotide RNA containing 30 24 nt single-strand overhang and 53 nt TRAP binding site (underlined) (50 GAGAUGAGAUGAGAUGAGAUGAGAUGAGAUGAGAUGAGAUGAGAUGAGAUGAGGUACCCACACUACACAUAGCCACC) is prepared by template directed ligation of two RNA pieces using T4 DNA ligase. 3. Bipartite control RNA duplexes were also prepared. Prepare TRAP-RNA complex 10 min prior to the reaction.

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4. TRAP-RNA remodeling reactions are performed in a buffer containing 40 mM Tris-HCl, pH 8.0, 0.01% (v/v) Nonidet P40, 0.5 nM radio labeled RNA, and 30 mM NaCl. 5. Add 20 nM TRAP and then add 20 nM DExH/D (NPH-II or DED1, see Notes 4 and 5), 4 mM MgCl2, 10 mM tryptophan. 6. Initiate the reaction by adding a mixture of 3.5 mM ATP and 600 nM of TRAP RNA scavenger DNA (DNA oligonucleotide that hybridize to the TRAP binding site and thus prevent re-binding of TRAP to its cognate RNA once it has been displaced). 7. Collect the aliquots at various time periods and stop the reaction by 5 mM EDTA and NPH-II (DED1) scavenger RNA (prevents helicase binding to the RNA from which TRAP has been displaced). 8. Electrophoreses the samples on 8% non-denaturing PAGE (1.5 mm thick, run at 10 V/cm at 4C). 9. Dry the gels and visualize the bands corresponding to TRAP bound RNA and free RNA by using a Phosphor Imager (GE Healthcare). 10. Quantify the radioactivity corresponding to the respective bands using the Image Quant software (GE Healthcare). 11. Calculate the amount of TRAP displaced by helicases from the amount of radioactivity in TRAP complexes and the amount of radioactivity in free RNA. Fraction [P] ¼ [RNA]/([RNA]+[TRAP]). Schematic of the method is shown in Fig. 6.7.

Fig. 6.7. TRAP displacement from RNA by DExH/D proteins. Prepare TRAP-RNA complex 10 min prior to the reaction. Add DExH/D protein to the TRAP-RNA complex and initiate the reaction by the addition of ATP. Quench the reaction with EDTA and scavenger RNA, which prevents helicase rebinding to the RNA after TRAP displacement. Collect the aliquots at various time points and separate by electrophoresis on non-denaturing gel as explained in methods. 3.4.2. Method 2

1. Prepare EJC complex by splicing radio labeled RNA in vitro. 2. Radiolabel mRNA at position –21 relative to the 50 splice site by template-directed ligation using T4 DNA ligase.

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3. RNA is spliced in vitro at 30C for 60 min using Hela Cell extract (source of exon junction proteins), followed by glycerol gradient sedimentation. 4. All the fractions were collected from gradient sedimentation. RNA is digested with micrococcal nuclease to identify the fractions containing the radiolabeled RNA with bound EJC followed by denaturing PAGE. 5. Pool the fractions containing EJC-RNA complex and desalt by size exclusion chromatography (Biorad P6 spin columns). 6. Tailless EJC-RNA complex is prepared by splicing mRNA with micrococcal nuclease prior to desalting. 7. EJC-RNP remodeling reactions are performed at room temperature in a buffer containing 40 mM Tris-HCl, pH 8.0, 0.01% (v/v) Nonidet P40, 0.1 nM EJC bound to radio labeled RNA, and 40 mM NaCl. Add 2 mM MgCl2, 30 nM NPH-II (600 nM DED1), and 0.5 nM RNA control duplex (wherever is applicable) and incubated for 10 min. 8. Remodeling reaction is initiated by the addition of ATP (10 mM). Reactions are stopped at appropriate time periods with EDTA solution (2 mM final) and immediately placed on ice.

Fig. 6.8. DExH/D helicases displace exon junction complex from RNA. Prepare EJC-RNA complex as explained above. DExH/D helicase (NPH-II/DED1) was added to the EJC-RNA complex and reaction was initiated by the addition of ATP. Collect the samples at various time points and quench with EDTA. Perform gel electrophoresis and quantitate the radioactivity in the respective bands by Image Quant software.

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9. Subsequently, ATP is removed using P6 spin columns by size exclusion chromatography (Biorad, Hercules, CA). 10. Samples are digested with micrococcal nuclease for 10 min, followed by phenol extraction and ethanol precipitation. 11. Add a 22-nucleotide DNA standard to normalize for loading and other deviations from denaturing PAGE (10%). 12. Gels are dried and bands corresponding to EJC-RNA complex and free RNA are visualized by using a Phosphor Imager (GE Healthcare, Piscataway, NJ). 13. Radioactivity in the respective bands is quantified by Image Quant software (GE Healthcare). Schematic of the method is shown in Fig. 6.8. .

4. Notes 1. Acrylamide is a neurotoxin when it is not polymerized; hence, care should be taken not to receive exposure. 2. Unless stated otherwise, use water for reagents preparation. 3. TEMED is best stored at room temperature in desiccators. Purchase small bottles as quality may decline (gel may take longer time to polymerize) after opening. 4. Avoid air bubbles with protein solutions, as it may denature the protein. 5. Avoid repeated freeze thaw cycles with protein solutions, it may reduce the activity of the protein. 6. Prevent SDS powder inhalation/exposure as it may cause irritation to eyes and nose. 7. Avoid exposure to radioactive substances by proper shielding and wearing eye protection. 8. MOPS buffer should be stored in brown bottles to avoid exposure to light. References 1. Delagoutte E. and von Hippel P. H. (2002) Helicase mechanisms and the coupling of helicases within macromolecular machines. Part I: Structures and properties of isolated helicases. Q. Rev. Biophys. 35, 431–478. 2. Lohman T. M. and Bjornson K. P. (1996) Mechanisms of helicase-catalyzed DNA

unwinding. Annu. Rev. Biochem. 65, 169–214. 3. Soultanas P. and Wigley D. B. (2001) Unwinding the ‘Gordian knot’ of helicase action. Trends Biochem. Sci. 26, 47–54. 4. Matson S. W., Bean D. W., and George J. W. (1994) DNA helicases: enzymes with

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5.

6.

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8.

9.

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11.

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13.

essential roles in all aspects of DNA metabolism. BioEssays 16, 13–22. Lahue E. E. and Matson S. W. (1988). Escherichia coli DNA helicase I catalyzes a uniderctional and highly processive unwinding reaction. J. Biol. Chem. 263, 3208–3215. Delagoutte E. and von Hippel P. H. (2002) Helicase mechanisms and the coupling of helicases within macromolecular machines. Part I:structures and properties of isolated helicases. Q. Rev. Biophys. 35, 431–478. Eggleston A. K., O’Neill T. E., Bradbury E. M., and Kowalczykowski S. C. (1995) Unwinding of nucleosomal DNA by a DNA helicase. J. Biol. Chem. 270, 2024–2031. Morris P. D. and Raney K. D. (1999) DNA helicases displace streptavidin from biotinlabeled oligonucleotides. Biochemistry 38, 5164–5171. Morris P. D., Byrd A. K., Tackett A. J., Cameron C. E., Tanega P., Ott R., et al. (2002) Hepatitis C virus NS3 and simian virus 40 T antigen helicases displace streptavidin from 50 -biotinylated oligonucleotides but not from 30 -biotinylated oligonucleotides: evidence for directional bias in translocation on single-stranded DNA. Biochemistry 41, 2372–2378. Byrd A. K. and Raney K. D. (2004) Protein displacement by an assembly of helicase molecules aligned along single-stranded DNA. Nat. Struct. Mol. Biol. 11, 531–538. Byrd A. K. and Raney K. D. (2006) Displacement and unwinding of trp repressor by Dda helicase. Nucleic Acids Res. 34, 3020–3029. Anand S. P., Zheng H., Bianco P. R., Leuba S. H., and Khan S. A. (2007) DNA helicase activity of PcrA is not required for the displacement of RecA protein from DNA or inhibition of RecA-mediated strand exchange. J. Bact. 189, 4502–4509. Fairman M. E., Maroney P. A., Wang W., Bowers H. A., Gollnick P., Nilsen T. W.,

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et al. (2004) Protein displacement by DExH/D ‘‘RNA helicases’’ with out duplex unwinding. Science 304, 730–734. Flores M. J., Sanchez N., and Michel B. (2005) A fork-clearing role for UvrD. Mol. Microbiol. 57, 1664–1675. Macris M. A. and Sung P. (2005) Multifaceted role of the Saccharomyces cerevisiae Srs2 helicase in homologous recombination regulation. Biochem. Soc. Trans. 33, 1447–1450. Jankowsky E., Gross C. H., Shuman S., and Pyle A. M. (2001) Active disruption of an RNA-protein interaction by a DExH/D RNA helicase. Science 291, 121–125. Fyodorov D. V. and Kadonaga J. T. (2001) The many faces of chromatin remodeling: switching beyond transcription. Cell 106, 523–525. Lusser A. and Kadonaga J. T. (2003) Chromatin remodeling by ATP-dependent molecular machines. Bioessays 25, 1192–1200. Sprouse O. R., Brenowitz M., and Auble D. T. (2006) Snf2/Swi2-related ATPase Mot1 drives displacement of TATA-binding protein by gripping DNA. EMBO J. 25, 1492–1504. Park J., Marr M. T., and Roberts J. W. (2002) Escherichia coli transcription repair coupling factor (Mfd protein) rescues arrested complexes by promoting forward translocation. Cell 109, 757–767. Gohara D. W., Ha C. S., Kumar S., Gosh B., Arnold J. J., Wisniewki T. J., et al. (1999) Production of ‘‘authentic’’ poliovirus RNAdependent RNA polymerase (3D(pol)) by ubiquitin-protease-mediated cleavage in Escherichia coli. Protein. Expr. Purif. 17,128–138. Tackett A. J., Wei L., Cameron C. E., and Raney K. D. (2001) Unwinding of nucleic acids by HCV NS3 helicase is sensitive to the structure of the duplex. Nucleic Acids Res. 29, 565–572.

Chapter 7 In Vivo Investigation of Protein–Protein Interactions for Helicases Using Tandem Affinity Purification Matthew Jessulat, Terry Buist, Md Alamgir, Mohsen Hooshyar, Jianhua Xu, Hiroyuki Aoki, M. Clelia Ganoza, Gareth Butland, and Ashkan Golshani Abstract A key component in determining the functional role of any protein is the elucidation of its binding partners using protein–protein interaction (PPI) data. Here we examine the use of tandem affinity purification (TAP) tagging to study RNA/DNA helicase PPIs in Escherichia coli. The tag, which consists of a calmodulin-binding region, a TEV protease recognition sequence, and an IgG-binding domain, is introduced into E. coli using a lred recombination system. This method prevents the overproduction of the target protein, which could generate false interactions. The interacting proteins are then affinity purified using double affinity purification steps and are seperated by SDS-PAGE followed by mass spectrometry identification. Each protein identified would represent a physical interaction in the cell. These interactions may potentially be mediated by an RNA/DNA template, for which the helicase would likely be needed to disrupt the secondary structures. Key words: Helicase, protein–protein interaction, tandem affinity purification, homologous recombination, RNA/DNA secondary structures.

1. Introduction Within the cell, individual proteins must frequently interact with one another to fulfill their function. As a result, protein– protein interactions (PPIs) can frequently yield clues as to what an uncharacterized protein’s function might be, or may reveal novel functions for partially characterized genes (1). In other cases, proteins form complexes, which have a combined activity that the individual parts may not accomplish in isolation. In addition, recruiting factors, chaperones, and a variety of other additional proteins along with non-protein elements such M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_7, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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cofactors must often be available for an individual protein to function. For example, various nucleic acid modifying proteins may require a helicase such as the DeaD protein in E. coli to unwind structured RNA templates so that the target region becomes accessible (2). The analysis and characterization of unknown proteins by observing PPIs has been widely used in functional genomics. Techniques used to observe these interactions include yeast two-hybrid analysis, microarray-based binding strategies, and purifying complexes from tagged proteins (1). Isolating complexes by using a ‘bait’ protein fused with an affinity tag has the advantage of detecting interactions that exist in vivo (3). The tandem affinity purification (TAP) tagging method is based on the principle of double affinity tagging a target protein and isolating the complex it makes with other proteins, by highly enriching the tagged protein and its binding partners using two rounds of affinity purification. This procedure was first applied for use in the yeast, Saccharomyces cerevisiae, but has also been shown effective in E. coli (4), and similar procedures have been successfully applied to plants (5), insects (6), and mammalian cells (4, 7, 8). By combining TAP tagging with mass spectrometry, reliable identification of interacting proteins can be achieved (1). High-throughput studies based on the TAP methodology have also been undertaken (9–11). The TAP method of isolating protein complexes requires a protein of interest (the bait) to be fused with a calmodulinbinding domain, a TEV protease cleavage site, and an IgGbinding domain (derived from Protein A). The complex is enriched through two affinity purification steps: initially, the bait and bound proteins are bound to beads coated in IgG, and in the second affinity purification step, they bind to calmodulin beads. The complex is released first by TEV cleavage of the tag (separating the protein of interest from the IgG-binding domain), then by washing with EGTA to release the proteins from the calmodulin beads (3). An important consideration in the TAP procedure is that the protein in question must remain at its native expression level and localization in order to provide reliable results. Alteration of protein levels through overproduction of the protein of interest can create false interaction results (4). To minimize these potential errors, the bait protein should ideally be kept under its native promoter and at the same copy number, rather than introducing a plasmid expression vector. Using a lred recombination system (12) for transformation of E. coli cells with a tagging-PCR product followed by standard TAP protocols allows for rapid isolation of proteins that physically interact with the bait protein, under natural or near-natural conditions in E. coli.

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Here we describe the methods used to fuse the TAP tag to the C-terminal end of a generic target helicase, isolate the tagged helicase along with any bound proteins using two rounds of affinity purification steps, and finally separate and examine the proteins found to interact with the tagged protein using polyacrylamide gel electrophoresis. The overview of this approach is summarized in Fig. 7.1. Please note that due to space limitation, we refer to mass spectrometry as a tool for protein identification only, technical details for which could be found elsewhere (13).

Fig. 7.1. A schematic overview of the TAP procedure. (a) The linear PCR product used to transform DY330 cells. It contains both the TAP tag sequence and a selectable kanamycin marker. Plasmids are not used to avoid overexpression of the protein of interest. (b) Incorporation of the TAP tag into the chromosomal DNA is achieved by inducing l recombination. Homologous regions on the PCR product direct the tag to the desired region. Cells are then lysed with sonication. (c) Cell lysate contains both complexes containing the bait protein and contaminants, under natural expression levels. (d) Two rounds of affinity purification, first with IgG beads, and second with calmodulin beads. Contaminants are washed out, the protein complex of interest is released by TEV or EGTA, respectively. (e) Identification of bound proteins is conducted by first separating proteins on an SDS-PAGE gel, followed by MALDI-TOF mass spectrometry.

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2. Materials 2.1. Tag Incorporation

1. Plasmid pJL72 (4), suspended in sterile water. 2. LongAmp Taq PCR kit (New England Biolabs). Contains enzyme (store at –20C), buffer, and dNTP mix. 3. Primers (order from Sigma Genosys). The PCR primers must be designed with a 50 region containing a 40- to 50-bp region homologous to the region being replaced by the tag and an inner 30 conserved region that will allow amplification of the tag, which is attached to the kanamycin (depending on the cassette used) resistance marker. In case of pJL72, the conserved regions used were TCCATGGAAAAGAGAAG (forward primer) and CATATGAATATCCTCCTTAG (reverse primer). The forward primer must be immediately adjacent and in-frame with the stop codon of the gene being tagged. 4. QIAgen PCR purification kit (QIAgen). Contains spin columns, collection tubes, buffer PB, and buffer PE. Store at room temperature. 5. LB medium: dissolve 10 g tryptone, 5 g yeast extract, and 10 g NaCl in 900 ml of distilled water. Adjust pH to 7.5 and bring to final volume of 1 1. Sterilize by autoclave and store at 4C. 6. Falcon tubes (14 ml) (Ultident scientific). 7. Centrifuge tubes (1.5 ml) (Diamed). 8. Electroporation cuvettes (Bio-Rad). 9. LB plates with kanamycin: Dissolve 10 g tryptone, 5 g yeast extract, 10 g NaCl and 20 g agar in 900 ml of distilled water. Adjust pH to 7.5 and bring to final volume of 1 1. Sterilize by autoclave and allow to cool to 65C. Add 1 ml of kanamycin stock solution (50 mg/ml) and pour into petri plates (Ultident). Allow to solidify at room temperature, then store at 4C. 10. Kanamycin stock solution: Dissolve 500 mg of kanamycin (Fisher) in 10 ml distilled water. Filter-sterilize and store at –20C in 1-ml aliquots until ready to use. Use 1 ml per ml of final solution volume.

2.2. Affinity Purification

1. Terrific Broth: combine 12 g tryptone, 24 g yeast extract, and 5 ml glycerol in 900 ml final volume distilled water and autoclave. Add 100 ml of 0.17 M KH2PO4, 0.73 M K2HPO4, autoclaved separately. Store at 4C. 2. Sonication buffer: 10 mM Tris–HCl, pH 7.9, 100 mM NaCl, 0.2 mM EDTA, 10% glycerol, 0.5 mM DTT. Add DTT immediately before use. Store at room temperature. 3. Bio-spin columns (Bio-Rad).

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4. IgG Sepharose beads (Amersham Biosciences) washed in M2 buffer prior to use. 5. M2 buffer: 10 mM Tris–HCl, pH 8, 100 mM NaCl, 0.1% Triton X-100, 10% glycerol, 1 mM DTT. Add DTT immediately before use. Store at room temperature. 6. TEV buffer: 50 mM Tris–HCl, pH 7.9, 100 mM NaCl, 0.2 mM EDTA, 0.1% Triton X-100, 1 mM DTT. Add DTT immediately before use. Store at room temperature. 7. TEV protease (Invitrogen): store at –20C. 8. 1 M CaCl2: store at room temperature. 9. Calmodulin-sepharose beads (Amersham Biosciences). Wash with binding buffer prior to use. 10. Binding buffer: 10 mM Tris–HCl, pH 7.9, 100 mM NaCl, 2 mM CaCl2, 0.1% Triton X-100, 10 mM 2-mercaptoethanol. Store at room temperature. 11. Wash buffer: 10 mM Tris–HCl, pH 7.9, 100 mM NaCl, 0.1 mM CaCl2, 0.1% Triton X-100, 10 mM 2-mercaptoethanol. Store at room temperature. 12. Elution buffer: 10 mM Tris–HCl, pH 7.9, 100 mM ammonium bicarbonate, 3 mM EGTA, 10 mM 2-mercaptoethanol. Store at room temperature. 2.3. Separation and Characterization of Interacting Proteins

1. Electrophoresis equipment (Bio-Rad or any other brand): requires two glass plates (one smaller front plate and one larger back plate), two spacers, comb, clamp assembly, casting stand, cooling core, electrophoretic cell, and power supply. 2. 50 ml centrifuge tubes (Ultident). 3. Separating gel: combine in a 50-ml centrifuge tube, in order, 2.14 ml of 30% acrylamide/bis acrylamide (97% acrylamide, 3% bis acrylamide, in sterile water), 1.53 ml of distilled water, 1.25 ml of 1 M Tris–HCl, pH 8.8, 50 ml 10% SDS, and 5 ml TEMED. Add 22 ml of 30% ammonium persulfate. Must be used immediately. 4. Stacking gel: in a 50-ml centrifuge tube, in order, mix 0.233 ml 30% acrylamide/bis acrylamide, 1.23 ml sterile water, 0.5 ml 1 M Tris–HCl, pH 6.8, 20 ml 10% SDS, 1.3 ml TEMED. Add 13 ml ammonium persulfate. Must be used immediately. 5. Electrode buffer: dissolve 3 g Tris base, 14.4 g glycine, and 1 g SDS in 1 l water. Can be prepared as a 5  concentrate solution in advance. Store at 4C. 6. Reducing buffer: 1 ml 0.5 M Tris–HCl, pH 6.8, 0.8 ml glycerol, 1.6 ml 10% SDS, 0.4 ml mercaptoethanol, 4.2 ml distilled water, 10 ml bromophenol blue.

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7. Protein molecular weight marker (Bio-Rad or any other brand). 8. Fixing solution: 120 ml glacial acetic acid, 500 ml 96% ethanol, and 500 ml formalin. Bring to a final volume of 1 l with distilled water. Store at room temperature. Note that glacial acetic acid should not be allowed to come into contact with skin. 9. 20% ethanol: bring 208 ml of 96% ethanol to a final volume of 1 l with distilled sterile water. Store at room temperature. 10. Sensitizing solution: add 200 mg sodium thiosulfate anhydrate to a small amount of distilled sterile water. Mix and bring to a final volume of 1 l. Store at room temperature. 11. Silver staining solution: combine 2 g silver nitrate, 50 ml water, and 760 ml formalin. Bring to 1 l final volume with water. Cool to 4C and use immediately. 12. Developing solution: 60 g sodium carbonate, 4 mg sodium thiosulfate anhydrate, 0.5 ml of formalin. Dissolve in a small volume of water, then bring to final volume of 1 l. Store at room temperature. 13. Terminating solution: mix 120 ml of glacial acetic acid with water, to a final volume of 1 l. Store at room temperature. 14. Acetonitrile (Sigma). 15. Trypsin in gel digestion buffer: 12.5 mg/ml trypsin in 50 mM ammonium bicarbonate. Prepare only what is needed and use immediately. 16. 25 mM ammonium bicarbonate: dissolve 0.198 g of ammonium bicarbonate in 100 ml of distilled water. Store at room temperature. 17. Extraction solution: 50% acetonitrile, 5% formic acid.

3. Methods To properly examine protein-binding partners, it is important not to alter the expression level or localization of the protein of interest. Use of a gene expression vector such as a plasmid would result in increased protein production, and it is possible the overproduction of the bait protein may lead to interactions that do not reflect the protein’s natural behavior. As a result, lred recombination is used to incorporate a DNA cassette encoding an affinity tag to the 30 end of a gene of interest, at its chromosomal locus, taking care to remove the native stop codon, to allow translational fusion of the gene of interest and the affinity tag and maintain natural gene

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expression levels. This allows the gene of interest to maintain its native promoter region, while a selective marker allows for the detection of tag incorporation. When affixing a tag to the bait protein, there is the possibility that this tag will block a binding site or other region of physiological importance on the C-terminal region of the protein of interest, thus preventing the detection of some protein interactions or creating a mutant phenotype. To examine this possibility, the protein tag which is affixed here to the C-terminal end of the protein can instead be linked to the N-terminal end (14). However, this requires the use of a separate tag template and may also result in the protein of interest no longer being expressed under its natural promoter. As an alternative, reciprocal tagging can be used to examine a target interaction. Regardless of the circumstances, reciprocal tagging is often used to minimize false positives and confirm the observed interactions (14). It is also possible to use different tags, such as the sequential peptide affinity (SPA) tag (4), which may provide better yields with some proteins. The SPA tag consists of a calmodulin-binding sequence, a TEV cleavage site, and a 3  FLAG sequence. Using this tag requires small changes to the procedure, specifically the use of anti-FLAG M2 agarose beads (from Sigma) in place of IgG beads in step 3, and the removal of DTT from the M2 buffer recipe. It should be re-emphasized that in most cases, PPIs can indicate the function of a poorly characterized target protein, or new functions to previously characterized proteins. However, a helicase, such as the DeaD protein (2), may generally interact with proteins of many functions and in many pathways. Because helicases are needed to unwind structured nucleic acids, it is reasonable for a single helicase to show physical interactions across many pathways. The common feature would not indicate a functional relationship between interacting proteins, but rather a reliance on nucleic acid unwinding for the various proteins or complexes identified (2). 3.1. Tag Incorporation

1. Amplify the TAP-tag and kanamycin marker section of plasmid pJL72 using standard PCR protocols (see Notes 1 and 2) and primers listed in Section 2. 2. Purify the linear DNA using a QIAgen PCR purification kit. Add 5 volumes of buffer PB to the PCR reaction tube. Mix by inversion, and transfer to a spin column placed inside a collection tube. Centrifuge for 1 min at maximum speed on a tabletop microcentrifuge. Discard flowthrough. Add 0.75 ml of PE buffer and spin at same speed. Discard flowthrough, spin again, and discard any additional flowthrough.

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Place spin column in a clean 1.5-ml centrifuge tube. Add 30 ml of sterile distilled water to the column, let stand for 1 min, and centrifuge to elute DNA. 3. Inoculate a 5-ml culture of E. coli strain DY330, containing the lred prophage sequence under temperature-sensitive regulation, in LB media. Incubate at 32C, with shaking, overnight. 4. Transfer 1 ml of culture to a flask containing 50 ml of fresh LB media and continue to incubate at 32C with shaking until the culture’s OD600 reaches between 0.4 and 0.6 (approximately 2–4 h). 5. Toinducecellsforrecombinationactivity,transfer10mlofculture to a 50-ml sterile flask and transfer to a water bath at 42C for 15 min. Immediately transfer to an ice-water bath for 10 min. 6. Transfer cells to a sterile 14-ml falcon tube and centrifuge at 7000 rpm for 2 min. Discard supernatant and resuspend pellet in 1 ml ice-cold sterile water. Transfer to a 1.5-ml centrifuge tube. Centrifuge at maximum speed at 4C on a tabletop microcentrifuge to wash cells and discard supernatant. Repeat wash a second time with a second 1-ml volume of ice-cold sterile water. 7. Resuspend cells in 300 ml sterile ice-cold water (see Note 3). 8. Transfer 100 ml of competent cells to an electroporation cuvette. Add 1 ml of linear PCR product. 9. Electroporate cells at settings of 1.8 KV, 200 , and 25 mFD in a Bio-Rad Gene Pulser or equivalent electroporation machine. Transfer cells to 1 ml of fresh LB media in a 1.5ml centrifuge tube and incubate at 32C for 1 h. 10. Centrifuge cells on a tabletop microcentrifuge for 1 min at maximum speed. Discard supernatant and resuspend pellet in 100 ml of fresh LB. 11. Spread over an LB plate containing 50 ml/ml kanamycin. Grow at 32C for 1–2 days. Choose individual transformant colonies for further study. 3.2. Affinity Purification

1. Inoculate a selected transformant in 4 l of Terrific Broth (TB) and allow to grow at 32C with shaking to late log phase. Harvest cells by centrifugation (7000 rpm, 6 min) and keep pellet at –80C until ready to use. 2. Prepare cell extract by sonication (see Note 4) of the pellet in 35 ml of sonication buffer (see Notes 5 and 6). All following steps should be performed at 4C. 3. To remove cellular debris, centrifuge at 20,000  g for 30 min. 4. Transfer 10 ml of sample to a Bio-Spin column containing 100 ml IgG sepharose beads. Incubate for 3 h with rotation.

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5. Drain the Bio-Spin column by opening the plug at the bottom of the tube and the cap at the top. Allow eluate to filter out by gravity. Wash the beads by adding 5 ml M2 buffer and draining the column to remove unbound proteins and contaminants (see Note 7). 6. Add 200 ml TEV buffer to the column with 50 units TEV protease. Incubate for 12 h (see Note 8). 7. Drain eluate directly into a new Bio-spin column. Close the plug and add 400 ml TEV buffer, 1.2 ml 1 M CaCl2, and 50 ml calmodulin-sepharose beads, washed with binding buffer. Incubate with rotation for 3 h. Drain eluate. 8. Wash the column with 400 ml of the binding buffer and drain. Wash the column with 100 ml wash buffer and drain. 9. Add 300 ml of elution buffer. Incubate for 1–2 min. Drain eluate, containing protein of interest and interacting proteins, into a microtube. 3.3. Separation and Characterization of Interacting Proteins

1. Separate two glass plates (one small front plate, one larger back plate) using a pair of spacers set to the outer left and right edges of the plates. Ensure the bottom and sides of the two glass plates are perfectly aligned. Screw the separated plates into the clamp assembly to secure in place. Fit the clamp assembly into a casting stand, making sure the bottoms of the plates are pressed firmly against the bottom of the casting stand and ensure all sides are tightly sealed to prevent leaking when the gel is cast. 2. In a 50-ml centrifuge tube, mix fresh separating gel. Swirl gently to mix, avoiding air bubbles. Note that unpolymerized acrylamide is toxic and avoid direct contact. 3. Use a blue-tip micropipette to transfer 1 ml at a time of the acrylamide solution between the glass plates. Fill the space between the glass plates to approximately 2 cm from the top. 4. Fill the rest of the space with sterile isobutanol and allow gel to set for 30 min. 5. Rinse off isobutanol with distilled water. Remove water by blotting the top of the gel with a kimwipe. 6. Mix stacking gel. Swirl immediately to mix, careful not to create bubbles. 7. Insert a comb. Use a blue-tip micropipette to transfer enough stacking gel to almost fill the space between the glass plates. Allow to set for 30 min. 8. Remove the comb and transfer the gel assembly from the casting stand to the cooling core in the electrophoretic cell. Fill the reservoir at the top and bottom of the apparatus with

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electrode buffer. Add 1 volume of protein sample to 4 volumes of reducing buffer in a 1.5-ml centrifuge tube and load wells with 10 ml of this sample. One lane should also contain a protein molecular weight marker for size comparisons. Attach to a power supply and run the gel at 180 V for 45 min (see Note 9). 9. Separate the gel from the electrophoresis apparatus and transfer toacontainer filledwith asmallvolume of fixing solution(enough to immerse the gel in). Allow the gel to sit for 2 h or overnight. 10. Pour off the fixing solution and wash the gel three times with 20% ethanol. 11. Discard the last ethanol wash and add a volume of sensitizing solution sufficient to immerse the gel. Incubate with gentle rocking for 2 min. 12. Pour off the sensitizing solution and rinse the gel twice with water. 13. Add silver staining solution to the container and incubate for 20 min. Do not pour the solution directly on the gel, as this may cause uneven staining. 14. Discard the staining solution and wash the gel for 30 s to 1 min with an excess volume of water. 15. Wash the gel briefly with developing solution. Discard the solution and replace it with 300 ml of fresh developing solution. Incubate for 2–5 min or until bands reach a desired intensity. 16. Add 50 ml of terminating solution to the developing solution. Incubate with gentle rocking for 10 min. The gel can now be photographed and the size of protein bands can be compared to the protein molecular weight marker. 17. Transfer the gel to a clean glass plate. Excise visible bands with a clean razor blade, cutting the smallest piece possible while taking the entire band. Transfer the band to a 1.5-ml centrifuge tube. Cover with acetonitrile for 15 min at room temperature. 18. Centrifuge in a microcentrifuge at maximum speed and remove acetonitrile with a micropipette. If gel is not an opaque white color, repeat addition and removal of acetonitrile. 19. Add enough trypsin in gel digestion buffer to cover the gel. Incubate at 4C for 45 min to cause the gel to swell. Transfer to 37C for trypsin digestion and incubate overnight. 20. Centrifuge at max speed in a microcentrifuge for 2 min and collect the supernatant. Add a volume of 25 mM ammonium bicarbonate sufficient to cover the gel and incubate for 25 min. Remove ammonium bicarbonate with a micropipette and keep it in case subsequent protein extraction fails.

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21. Extract peptides by addition of 30 ml of extraction solution, incubate for 10 min, and centrifuge as before. Transfer to a clean centrifuge tube. Repeat this step three times, combining all extracted solution. 22. Using a speed-vac, evaporate extracted sample until the desired volume is reached. Sample is now ready for Matrix Assisted Laser Desorption /Ionization-Time Of Flight (MALDI-TOF) mass spectrometry and subsequent protein identification.

4. Notes 1. We recommend an annealing temperature of 50C for 30 s, an extension temperature of 68–72C for 2 min, and a melting temperature of 92C for 30 s, repeated for 30–35 cycles. 2. Due to the size of the PCR product and importance of sequence fidelity, we recommend using a high-fidelity polymerase for PCR, or one specifically suited to longer PCR products. Errors or frame-shifts in the extension phase could disrupt the tag or kanamycin-resistance marker, making the PCR product useless in further experiments. 3. It is possible to store the prepared cells at –80C for up to 10 days. However, efficiency of cells stored in this way will be reduced. Unless a large number of transformations requires the preparation of large batches of cells in advance, it is better to use only freshly prepared cultures. 4. Length and intensity of sonication treatment must be initially optimized. A longer, lower intensity treatment would yield results similar to a more intense but shorter treatment. However, the sonication treatment should not result in a significant increase in temperature. Sonication should lead to the disruption of the majority of cells (a sample can be visualized under a microscope, with 90–95% cell disruption being desired), but should fall short of disrupting protein complexes or denaturing proteins through temperature changes. 5. Sonication is one of many methods used to prepare a cell extract. Different methods may be appropriate for different cell types. For example, a yeast cell extract can be prepared by passing harvested cells through a french press. 6. To eliminate binding effects mediated by DNA or RNA bound to the helicase, benzonase or RNase A can be added to the sonication buffer. However, high concentrations of nuclease may interfere with subsequent affinity capture of the protein of interest.

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7. Initially, a sample of each elution should be saved. If the procedure does not yield the ‘bait’ protein, each elution step can then be tested for the presence of the tagged protein, in order to determine where binding may have failed. 8. Incubation at warmer temperatures (for example, 16C) will allow for shorter incubation times (2 h). 9. Voltage and time can be varied considerably to give the best results in any given experiment. Lower voltage would require more time for the same amount of migration, but may result in cleaner banding patterns. Increasing the gel concentration may also improve the appearance of protein bands, but again a longer time frame would be required for the same migration as a less concentrated band.

Acknowledgments This work was supported by a grant from the Natural Sciences and Engineering Research Council of Canada (NSERC) and is dedicated to the loving memory of Minoo Rajabian. References 1. Chepelev N., Chepelev L., Alamgir M., and Golshani A. (2008). Large-scale proteinprotein interaction detection approaches: past, present and future. Biotechnol. Biotechnol. Eq. 22, 513–529. 2. Butland G., Krogan N., Xu J., Yang X., Yang W., Aoki H., et al. (2007) Investigating the in vivo activity of the DeaD protein using preotien-protein interactions and the translational activity of structured chloraphenicol acetyltransferase mRNAs. J. Cell Biochem. 100, 642–652. 3. Rigaut G., Shevchenko A., Rutz B., Wilm M., and Seraphin B. (1999). A generic protein purification method for protein complex characterization and proteome exploration. Nat. Biotechnol. 17, 1030–1032. 4. Zeghouf M., Li J., Butland G., Borkowsa A., Canadien V., Richards D., et al. (2004) Sequential peptide affinity (SPA) system for the identification of mammalian and bacterial protein complexes. J. Proteome Res. 3, 463–468. 5. Rubio V., Shen Y., Saijo Y., Liu Y., Gusmaroli G., Dinish-Kumar S., et al. (2005) An

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