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

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Author: Mohamed M. Abdelhaleem

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METHODS

IN

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

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

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

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

1

2.

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

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3.

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

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

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

9.

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.

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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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alternative tandem affinity purification strategy applied to Arabidopsis protein complex isolation. Plant J. 41, 767–778. Forler D., K¨ocher T., Rode M., Gentzel M., Izaurralde E., and Wilm M. (2003) An efficient protein complex purification method for functional proteomics in higher eukaryotes. Nat. Biotechnol. 21, 89–92. Zhou D., ren J. X., Ryan T. M., Higgins N. P., and Townes T. M. (2004) Rapid tagging of endogenous mouse genes by recombineering and ES cell complementation of tetraploid blastocysts. Nucleic Acids Res. 32, e128. Gregan J., Riedel C. G., Petronczki M., Cipak L., Rumpf C., Poser I., et al. (2007) Tandem affinity purification of functional TAP-tagged proteins from human cells. Nature Protoc. 2, 1145–1151. Gavin A. C., B¨osche M., Krause R., Grandi P., Marzioch M., Bauer A., et al. (2002) Functional organization of the yeast proteome by systematic analysis of protein complexes. Nature 415, 141–147. Gavin A. C., Aloy P., Grandi P., Krause R., Boesche M., Marzioch M., et al. (2006)

Detecting Protein–Protein Interactions Proteome survey reveals modularity of the yeast cell machinery. Nature 440, 631–636. 11. Krogan N. J., Cagney G., Yu H., Zhong G., Guo X., Ignatchenko A., et al. (2006) Global landscape of protein complexes in the yeast Saccharomyces cerevisiae. Nature 440, 637–643. 12. Yu D., Ellis H. H., Lee E.-C., Jenkins N. A., Copeland N. G., and Court D. L. (2000) An efficient recombination system for chromosome engineering in Escherichia coli. Proc. Natl. Acad. Sci. U.S.A. 97, 5978–5983.

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13. Mann M., Højrup P., and Roepstorff P. (1993) Use of mass spectrometric molecular weight information to identify proteins in sequence databases. Biol. Mass Spectrom. 22, 338–345. 14. Puig O., Caspary F., Rigaut G., Rutz B., Bouveret E., Bragado-Nilsson E., Wilm M., and Seraphin B. (2001). The tandem affinity purification (TAP) method: a general procedure of protein complex purification. Methods 24, 218–229.

Chapter 8 Mapping Genomic Targets of DNA Helicases by Chromatin Immunoprecipitation in Saccharomyces cerevisiae Jennifer Cobb and Haico van Attikum Abstract DNA helicases utilize the energy of nucleotide hydrolysis to unwind the two annealed strands of the DNA helix and are involved in many aspects of DNA metabolism such as replication, recombination, and repair. Chromatin immunoprecipitation (ChIP) has been instrumental in determining the genomic targets of many DNA helicases and DNA helicase-containing complexes including the minichromosome maintenance (Mcm) proteins 2–7, the RecQ helicase Sgs1 as well as the Rvb1 and Rvb2 helicase-containing INO80 and SWR1 chromatin remodeling complexes. Here we describe a ChIP method that has been successfully used to map these proteins at chromosomal double-strand breaks and replication forks in the model organism Saccharomyces cerevisiae. Key words: DNA helicases, chromatin immunoprecipitation (ChIP), Saccharomyces cerevisiae, DNA repair, DNA replication.

1. Introduction DNA helicases are involved in many aspects of DNA metabolism, including replication, recombination, and repair. ChIP has been used to show the temporal recruitment of helicases and helicase-containing complexes to sites of replication and DNA double-strand breaks (DSBs). One group is formed by the Rvb1 and Rvb2 helicases, which are members of the large AAAþ class of ATP-dependent ATPases. Rvb1 and Rvb2 form a heterohexameric complex that unwinds DNA (1). Moreover, they are integral subunits of multi-protein complexes, including the INO80 and SWR1 chromatin remodeling complexes, which have been implicated in transcription regulation, replication, and repair (2–5). ChIP has demonstrated the recruitment of INO80 M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_8, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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and SWR1 complexes to chromosomal DSBs (6–8). The same method has also demonstrated the association of INO80 with replication forks that encounter replicative stress and INO80 is thought to facilitate both fork progression and the recovery of stalled forks (9, 10). DNA unwinding driven by the Rvb1 and Rvb2 helicases is presumed to facilitate INO80 and SWR1 chromatin remodeling, which may explain why these helicases are associated with diverse cellular functions (2, 11). ChIP has also been performed on components of the Mcm2–7 helicase complex. Each Mcm protein is a member of the AAAþ class of ATPases and has an ATP binding site. However, no individual Mcm protein alone has significant ATPase activity (12). ATPase activity is generated by the interaction of at least two MCM proteins and requires an ATP binding site from one subunit and a catalytic residue from another subunit (13). The Mcm2–7 six component complex is presumed to function as the replicative helicase (14), loading onto well-characterized origins of replication at the end of mitosis. Lastly, the RecQ helicase Sgs1 is recruited to sites of replication only after initiation. By ChIP both Sgs1 and Mcm2–7 helicases have been shown to move with the fork during replication elongation (15, 16). The Mcm2–7 helicase is presumed to move ahead of the fork unwinding the double-stranded helix so that DNA polymerases can replicate the DNA, whereas Sgs1 is thought to have a role behind replication forks preventing the formation of aberrant recombination intermediates during replication. Determining the genomic loci of DNA helicases by ChIP in combination with in vitro assays showing their enzymatic activity has been extremely powerful for studying the function of these helicases in living cells. We will outline the method of ChIP that has been successfully used to study these proteins at replication forks and chromosomal DNA double-strand breaks (DSBs) in budding yeast (Saccharomyces cerevisiae) (6, 8, 15–17). Figure 8.1 schematically shows the different steps of the ChIP procedure. The ChIP starts with the formaldehyde treatment of cells to crosslink protein–protein and protein–DNA complexes. After crosslinking, the cells are lysed and crude extracts are sonicated to shear the DNA. Proteins together with crosslinked DNA are immunoprecipitated. Because S. cerevisiae is amenable to genetic manipulations the challenge of antibody selection for immunoprecipitation in many cases can be overcome by the addition of an epitope tag (e.g., Myc or HA) to the endogenous protein. Protein–DNA crosslinks in the immunoprecipitate and input (non-immunoprecipitated whole cell extract) are then reversed and the DNA fragments are purified. Quantitative real-time PCR is then performed to amplify the region where either a protein or protein modification is present. DNA fragments of this genomic locus should be enriched in the immunoprecipitate compared to that in the input (which represents all portions of the genome). In principle, the method of ChIP is

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Fig. 8.1. Schematic showing the strategy of ChIP for the mapping of genomic targets of chromatin associated proteins. The method starts with the in vivo formaldehyde treatment of cells to crosslink protein–protein and protein–DNA complexes (a). After crosslinking, the cells are lysed and crude extracts are sonicated to shear the DNA (b). Proteins together with crosslinked DNA are immunoprecipitated (c). Protein–DNA crosslinks in the input (nonimmunoprecipitated whole cell extract) and immunoprecipitate are then reversed and the DNA fragments are purified (d). Quantitative real-time PCR is then performed to amplify the region where either a protein or protein modification is present. DNA fragments of this genomic locus should be enriched in the immunoprecipitate compared to that in the input and a control immunoprecipitate (from no antibody or non-tagged strain) (e).

straightforward; the process itself, however, can be extremely challenging and success with ChIP depends a great deal on the protein of interest and appropriate antibodies.

2. Materials 2.1. Growth, In Vivo Crosslinking, and Harvest of Yeast Cells

1. YPLGg medium: 1% (w/v) yeast extract (Difco Laboratories), 2% (w/v) bactopeptone (Difco Laboratories), 2% (v/v) lactic acid (Fluka), 3% (v/v) glycerol, and 0.05% (w/ v) glucose (Fluka). Dissolve components in double distilled H2O, adjust pH to 6.6, and autoclave for 15 min at 115C.

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2. YPAD medium: 1% (w/v) yeast extract, 2% (w/v) bactopeptone, 0.0025% (w/v) adenine (Sigma), 2% (w/v) glucose. Dissolve components in double distilled H2O, adjust pH to 5.8, and autoclave. 3. YPAD, pH 5. 4. 20% (w/v) glucose and 20% (w/v) galactose stock solutions: dissolve components in double distilled H2O and sterilize by filtration. 5. Alpha-factor. 6. 37% formaldehyde. 7. 2.5 M Glycine: dissolve glycine in double distilled H2O and sterilize by filtration. 8. Phosphate buffered saline (PBS): 140 mM NaCl, 2.5 mM KCl, 8.1 mM Na2HPO4, 1.5 mM KH2PO4. Dissolve components in double distilled H2O, adjust pH to 7.5, and autoclave for 15 min at 115C. 9. 2 ml tubes with screw cap. 10. Hydroxyurea. 2.2. Preparation of Whole Cell Extracts

1. Lysis buffer: 50 mM HEPES/KOH, pH 7.5, 140 mM NaCl, 1 mM EDTA, 1% (v/v) Triton X-100, 0.1% (w/v) Na-deoxycholate, and protease inhibitors. 2. Protease inhibitors: 100 stocks of aprotinin (100%, store at 4C), 100 mM PMSF in isopropanol (freshly prepared), 1000 stocks of 2 mg/ml Antipain in H2O, 300 mg/ml benzamidin in H2O, 1 mg/ml PepstatinA in ethanol, 20 mg/ ml TPCK in ethanol, 10 mg/ml TLCK in H2O, 2000 stock of 1 mg/ml Leupeptin in H2O (all stored at –20C). 3. Zirconia/Silica beads (0.5 mm diameter). 4. 25G Needles (0.5 mm diameter). 5. 3 ml polypropylene tubes (12 55 mm). 6. 12 ml Falcon tubes.

2.3. Immunoprecipitation and DNA Isolation

1. Sheep anti-mouse IgG or sheep anti-rabbit IgG Dynabeads. 2. Monoclonal or polyclonal antibodies: monoclonal anti-Myc (9E10) and anti-HA (F-7, Santa-Cruz) work well for tagged proteins. 3. PBS-BSA: dissolve 5 mg/ml BSA (Sigma) in 1 PBS and sterilize by filtration. 4. Wash buffer: 100 mM Tris–HCl, pH 8, 250 mM LiCl, 0.5% (v/v) NP-40, 0.5% (w/v) Na-deoxycholate, 1 mM EDTA, and protease inhibitors. 5. TE: 10 mM Tris–HCl, pH 8, 1 mM EDTA.

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6. TE/1% SDS. 7. 5 M LiCl: dissolve LiCl in 50 mM Tris–HCl, pH 8, and autoclave for 15 min at 115C. 8. Phenol:chloroform:isoamylalcohol (25:24:1) 9. 20 mg/ml glycogen. 10. 20 mg/ml Proteinase K. 11. Isopropanol. 2.4. Quantitative PCR Analysis

1. TaqManTM probe: double-dye oligonucleotide that has a fluorescent reporter dye (FAMTM) and a quencher (TAMRATM) at its 50 and 30 end, respectively (see Note 1). 2. Oligonucleotides that anneal upstream and downstream of TaqManTM probe (see Note 1). 3. 2 TaqManTM Universal PCR master mix (Applied Biosystems). 4. MicroAmpTM optical 96-well reaction plate (Applied Biosystems). 5. MicroAmpTM optical adhesive film (Applied Biosystems). 6. Optical cover compression pad (Applied Biosystems). 7. Quantitative real-time PCR machine (Applied Biosystems).

3. Methods 3.1. Growth, In Vivo Crosslinking and Harvest of Yeast Cells 3.1.1. Growth of Yeast Cells Prior to ChIP of Proteins at DSBs

We have used strains with JKM179 background to study binding of proteins at DSBs (19). This strain expresses the HO endonuclease from a galactose-inducible promoter (HO is repressed when glucose is present in the medium). HO induces a unique DSB at the MAT locus on chromosome III. This break is usually repaired by recombination. However, in this strain the donor loci for repair were deleted in order to prevent repair by recombination. This allows us to monitor protein binding at the HO-induced DSB. 1. Inoculate YPAD medium with strain JKM179 or one of its derivatives and incubate and rotate at 180–210 rpm overnight at 30C. 2. Dilute the overnight culture in YPLGg medium such that the volume is consistent with the number of time points (100 ml/ time point). Incubate and rotate at 180–210 rpm overnight at 30C until OD600 ¼ 0.3–0.4 (see Note 2). 3. Pour 100 ml culture into a new flask and add glucose to a final concentration of 2%. 4. Add galactose to a final concentration of 2% to the rest of the overnight culture.

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5. Incubate and rotate the cultures at 180–210 rpm at 30C. 6. At each desired time point take 100 ml culture and pour it into a new flask for formaldehyde crosslinking as described in Section 3.1.3 (see Note 3); galactose-containing samples are taken at 0.5, 1, 2, and 4 h, whereas the glucose-containing control sample is taken at 2 h after sugar addition. 3.1.2. Growth of Yeast Cells Prior to ChIP of Proteins at Replication Forks

1. Inoculate YPAD medium with the strain(s) of interest and incubate and rotate at 180–210 rpm overnight at 30C. 2. Dilute the overnight culture to 1 106 cells/ml in YPAD medium such that the volume is consistent with the number of time points (50 ml/time point). Incubate and rotate at 180–210 rpm at 30C until the cell density of the culture is 5 106 cells/ml. 3. Collect cells by centrifugation at 1620 g at room temperature for 3 min. Remove supernatant and resuspend cells in YPAD, pH 5. 4. Collect cells by centrifugation at 1620 g at room temperature for 3 min. Remove supernatant and resuspend cells in YPAD, pH 5. 5. Add alpha-factor and incubate and rotate at 180–210 rpm at 30C for 1.5 h to arrest cells in G1 (see Note 4). 6. Collect cells by centrifugation at 1620 g at room temperature for 3 min. Remove supernatant and resuspend cells in YPAD to wash. 7. Collect cells by centrifugation at 1620 g at room temperature for 3 min. Remove supernatant and resuspend cells in YPAD without or with 200 mM hydroxyurea. 8. Incubate and rotate at 180–210 rpm at 16C when cultures do not contain hydroxyurea, and at 30C when cultures contain hydroxyurea. 9. At each desired time point take 50 ml culture and pour it into a new flask for formaldehyde crosslinking as described in Section 3.1.3 (samples are taken at 0 (cells in G1), 10, 20, 40, and 60 min after release into S phase).

3.1.3. In Vivo Crosslinking and Harvest of Yeast Cells

1. Add 1.5 ml (3 ml) 37% formaldehyde to 50 ml (100 ml) of yeast cell culture (1% final) and shake slowly at 100 rpm at 30C for 15 min (see Note 5). 2. Add 2.5 ml (5 ml) 2.5 M glycine to 50 ml (100 ml) of yeast culture and shake slowly at 100 rpm at 30C for 5 min. 3. Collect cells by centrifugation at 1620 g at room temperature for 3 min. Remove supernatant and resuspend cells in 25 ml 1 PBS to wash.

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4. Collect cells by centrifugation at 1620 g at room temperature for 3 min. Remove supernatant, resuspend cells in 1 ml 1 PBS and transfer them to a 2-ml tube with screw cap. 5. Collect cells by centrifugation at 9500 g at room temperature for 2 min, remove supernatant and freeze pellet at –80C. 3.2. Preparation of Beads Covered with Antibodies

1. Pellet 2 40 ml (80 ml) Dynabeads per sample by centrifugation at 10,000 g for 5 s (see Note 6). Place tube in magnetic stand and remove supernatant. 2. Resuspend Dynabeads in 0.5 ml ice-cold 1 PBS-BSA (5 mg/ml) and wash by constant rotation for 30 min at 4C. 3. Pellet Dynabeads by centrifugation at 10,000 g for 5 s. Place tube in magnetic stand and remove supernatant. 4. Resuspend Dynabeads in 0.5 ml ice-cold 1 PBS-BSA and add 5 mg antibody to 40 ml Dynabeads (see Note 6). Add a (non-related) control antibody or do not add antibody to the other 40 ml Dynabeads. Rotate for 2 h at 4C. 5. Pellet Dynabeads by centrifugation at 10,000 g for 5 s. Place tube in magnetic stand and remove supernatant. 6. Resuspend Dynabeads in 0.5 ml ice-cold wash buffer and wash by rotation for 5 min at 4C. 7. Repeat the last two steps. Pellet Dynabeads by centrifugation at 10,000 g for 5 s. Remove supernatant and resuspend each portion of Dynabeads in 40 ml ice-cold 1 PBS-BSA (5 mg/ml).

3.3. Preparation of Whole Cell Extracts (WCE)

1. Add 400 ml Zirconia/Silica (Biospec Products, Inc.) beads and 600 ml ice-cold lysis buffer to the cell pellet from Section 3.1.3 step 5. 2. Beadbeat 3 1 min with 1 min rest intervals at maximum at 4C to lyse the cells (see Note 7). 3. Make a hole in the bottom of the tube using a 25-G needle (0.5 mm diameter). 4. Place the tube in a 3-ml polypropylene tube and put both tubes in a 12-ml falcon tube. 5. Centrifuge at 380 g at 4C for 5 min to recover the extract. Remove and discard the 2-ml tubes with screw cap. Add 600 ml ice-cold lysis buffer to the WCE and resuspend the pellet. 6. Shear chromatin by sonication with four pulses of 20 s each with rest intervals of 1 min at 4C at power setting 2.5, 3.5, 3.5, and micro, respectively (see Note 8). 7. Transfer WCE to a pre-chilled eppendorf tube and centrifuge at 4650 g at 4C for 2 min to pellet insoluble debris.

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8. Transfer supernatant, which contains the soluble chromatin, to new pre-chilled eppendorf tubes. Save 25 ml aliquots (input samples) in 2 ml safe capslock eppendorf tubes and freeze at –20C until crosslinking reversal in Section 3.4 step 9 (see Note 9). 3.4. Immunoprecipitation and DNA Isolation

1. Divide WCE from Section 3.3 step 8 over two eppendorf tubes. 2. Add 40 ml Dynabeads with antibody from Section 3.2 step 7 to one half of the extracts and 40 ml Dynabeads without antibody or a control antibody, also from Section 3.2 step 7, to the other half of the extracts. Rotate for 2 h at 4C. 3. Pellet Dynabeads by centrifugation at 10,000 g for 5 s, place tubes in a magnetic stand, remove supernatant, and resuspend the Dynabeads in 600 ml ice-cold lysis buffer (see Note 10). Mix the Dynabeads for 5 min at 4C in a mixer. 4. Repeat step 3. 5. Pellet Dynabeads by centrifugation for 5 s at 10,000 g, place tubes in a magnetic stand, remove supernatant, and resuspend the Dynabeads in 600 ml ice-cold wash buffer (see Note 10). Mix Dynabeads for 5 min at 4C. 6. Pellet Dynabeads by centrifugation for 5 s at 10,000 g, place tubes in a magnetic stand, remove supernatant, and resuspend the Dynabeads in 600 ml ice-cold TE buffer. Mix Dynabeads for 1 min at 4C. 7. Pellet Dynabeads by centrifugation for 5 s at 10,000 g, place tubes in a magnetic stand, remove supernatant, and resuspend Dynabeads in 120 ml TE/1% SDS. Mix Dynabeads for 10 min at 65C in a thermomixer. 8. Pellet Dynabeads by centrifugation for 5 s at 10,000 g and place tubes in a magnetic stand. Transfer supernatant to 2 ml safe capslock eppendorf tubes (see Note 9). 9. Add 130 ml and 100 ml TE/1% SDS to the immunoprecipitate and input (from Section 3.3 step 8) samples, respectively, submerge the tubes in H2O, and incubate overnight at 65C to reverse crosslinks. 10. Add 240 ml and 370 ml TE buffer to the immunoprecipitate and input samples, respectively. 11. Add 20 ml proteinase K (20 mg/ml) and incubate for 2 h at 37C. 12. Add 50 ml 5 M LiCl and 400 ml phenol:chloroform:isoamylalcohol, vortex, and centrifuge for 5 min at 10,000 g.

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13. Transfer supernatant to new eppendorf tubes. Add 2 ml glycogen and 0.7 volume of isopropanol. Incubate for 30 min at –80C and pellet DNA by centrifugation for 30 min at 10,000 g at 4C. 14. Remove supernatant and add 0.5 ml 70% ethanol. Pellet DNA by centrifugation for 30 min at 10,000 g at 4C. 15. Remove supernatant and dry DNA under vacuum. Resuspend DNA in 30 ml H2O (see Note 11). 3.5. Quantitative PCR Analysis

Input and immunoprecipitated DNA are analyzed by quantitative real-time PCR. Regions from an HO-induced DSB or a control region (SMC2 gene) are amplified using TaqManTM PCR. For replication forks, origin regions ARS607 (early origin), ARS501 (late origin), and a non-origin region are amplified using TaqManTM PCR. This involves a double-dye oligonucleotide (TaqManTM probe) that has a fluorescent reporter dye (FAMTM) and a quencher (TAMRATM) at its 50 and 30 end, respectively, and primers that anneal upstream and downstream of the probe (see Note 1). During the amplification process the probe is cleaved by the 50 ! 30 exonuclease activity of the Taq DNA polymerase. This removes the fluorophore from the probe and separates it from the quencher. As a consequence the fluorophore starts to fluoresce. This fluorescence can be measured and the level is directly proportional to the amount of DNA amplified during the PCR reaction. Primers and probes are tested for comparable and linear amplification of specific and control fragments from a genomic DNA template (see Note 1). 1. Setup PCR reactions in 20 ml with 1, 6, or 18 pMol of each primer, 4 pMol of TaqManTM probe, 10 ml 2 TaqManTM Universal PCR master mix, H2O, and either 1 ml of input or immunoprecipitated DNA from Section 3.4 step 15 (see Note 1). Prepare a premix containing primers, TaqManTM probe, and H2O and a premix containing DNA, 2 TaqManTM Universal PCR master mix, and H2O. 2. Mix aliquots of the appropriate premixes in a 96-well reaction plate. Seal the plate with an optical adhesive film and cover the plate with a compression pad. 3. Place the plate in the real-time PCR machine and perform PCR using the following program: 2 min at 50C and 10 min at 95C followed by 40 cycles with 15 s at 95C and 1 min at 60C (see Note 1). 4. Retrieve the ct values, export them into an Excel spreadsheet and calculate the fold enrichment for your locus of interest (see Note 1). Examples for how the enrichment for INO80 at a DSB and Mcm7 at a replication fork can be presented are shown in Fig. 8.2

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Fig. 8.2. Examples of experiments that used the ChIP method described to examine the binding of INO80 near a DSB and the association of Mcm7 with an origin of replication. (a) A genomic region flanking the DSB (+1.6 kb) on Chr III, which was induced by a galactose-inducible HO endonuclease in strain JKM179, was amplified by PCR after ChIP. ChIP was performed using an antibody against the Myc epitope (9E10) in the tagged INO80 protein and a control antibody against the HA epitope (12CA5) on cells that were grown in the presence of glucose (for 2 h) to repress HO or galactose (1, 2, and 4 h) to induce HO. The enrichment for INO80 was calculated as described in the text and presented as the ratio of the signal in the Myc immunoprecipitate over that in the HA immunoprecipiate. Error bars represent the standard deviation in fold enrichment of multiple runs of at least two different ChIP experiments (b) Genomic regions amplified for ChIP analysis of Chr VI correspond to early-firing origin ARS607 (filled symbol) and a non-origin site, +14 Kb (open symbols). ChIP was performed using an antibody against the Myc epitope (9E10) in the tagged Mcm7 protein on cells released from -factor into YPAD + 0.2 M HU. The enrichment for Mcm7 at replication forks was calculated as described in the text and is presented as the increase in signal in the Myc immunoprecipitate over that in the immunoprecipitate with beads alone for both loci. Error bars represent the standard deviation in fold enrichment of multiple runs of at least three different ChIP experiments.

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4. Notes 1. Primers and probes are designed using the Primer Express software (Applied Biosystems). Primers are 20–30 bp in length with a melting temperature of 59C, whereas the Taqman probeTM has a melting point of at least 69C and does not start with a guanine (G) base. Primers and probes are tested for comparable and linear amplification of specific and control fragments from input samples; 1, 6 and 18 pMol of forward primer is combined with 1, 6, and 18 pMol of reverse primer, probe, 2 Taqman probeTM Universal PCR master mix and undiluted to 4096-fold diluted input DNA on, e.g., a 7000 Sequence Detector System (Applied Biosystems). The quantitative real-time PCR monitors a ‘threshold cycle (ct)’ at which the exponential curve of accumulated product passes a certain threshold. A standard curve is made by plotting the ct values against the log of DNA quantity. The slope (S) of the curve can be calculated and used to determine the amplification efficiency (AE) by using the formula: AE = 10(–1/S)–1. When the AE for a target and control sequence are comparable and near 2, then these can be used. The enrichment for a given target locus after ChIP can be represented by the ratio of the difference of signal accumulation rates for the immunoprecipitate versus input, divided by the difference in rates for a control immunoprecipitate using beads alone, beads covered with a (non-related) control antibody or a non-tagged strain (see Note 9). It can be calculated by using, e.g., the formula 2(ct_Input – ctI_IP)/2(ct_Input – ct_Control). Alternatively, a SYBR Green-based real-time PCR method, which does not require TaqmanTM probes, or classical PCR combined with agarose gel electrophoresis may be used to determine the enrichment for a given target locus after ChIP (20). 2. Wild-type strains and their isogenic mutant derivatives may differ in growth properties. Moreover, more complex media such as YPLGg or synthetic selective media may influence the doubling time of your yeast strains. As it is desirable to have synchronously growing strains and to harvest cells in the same growth phase, preferably in log-phase, we suggest that the doubling time of your strains is determined in a pilot experiment. This allows you to calculate the number of cells from a preculture required to reach a certain cell density after a specific period of growth. 3. The HO endonuclease can also be induced by adding galactose directly to cell cultures that have been pre-grown in YPA raffinose-containing media. However, the galactose-inducible

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promoter, which drives HO expression, may be leaky on raffinose-containing media and therefore HO-induced DSBs may be induced and cells may die. 4. The optimal concentration of alpha-factor depends on the strain background and should be determined empirically. FACS analysis is the best method to measure good synchronization in G1 and release into S phase after removal of alphafactor. 5. The optimal crosslinking time varies from few minutes to few hours and should be determined empirically. Addition of protein–protein crosslinkers may improve signals for some targets. 6. We have successfully used sheep anti-mouse IgG and sheep anti-rabbit IgG Dynabeads (Dynal Biotech ASA/Invitrogen) for ChIP. We usually use up to 5 mg of antibody to completely saturate the Dynabeads. Saturation of your beads may be tested by incubating beads with increasing amounts of affinity purified antibody as described in Section 3.2 steps 1–7. Save the recovered supernatant. Resuspend the beads in sample buffer, incubate for 5 min at 95C and recover the supernatant as well. Perform Western blot analysis on aliquots of these supernatants. The concentration at which the beads cannot completely remove all antibodies from the solution should be used in experiments. Depending on the affinity of your antibody for proteinA or proteinG, proteinA or proteinG sepharose beads can also be used. Alternatively, biotinylated proteins may be immunoprecipitated with streptavidincoupled beads (Dynal Biotech ASA/Invitrogen) (9, 21). The advantage of this method may be the use of high salt and SDS concentrations in the lysis and wash buffers, which may efficiently reduce non-specific precipitation. 7. The conditions mentioned in Section 3.3 step 2 resulted in more than 80% cell lysis with a Mini Beadbeater-8 (Biospec Products, Inc.). Alternatively, cell breakage may also be achieved using a vortex shaker, such as Eppendorf model 5432 at 4C for 40 min. 8. The extent of sonication will determine the average size of your chromatin fragments and thus the resolution of mapping in your ChIP experiment. The smaller the fragments the higher the resolution of mapping. The conditions for sonication have to be established for each sonicator. To establish these conditions some WCE can be prepared and subjected to an increasing number of sonication cycles with different time and power settings. Small aliquots of WCE should be removed after each cycle. The DNA should be cleaned up as described in Section 3.4 steps 9–15 and analyzed by agarose gel electrophoresis in order to determine the amount of

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cycles, time, and power settings that shear your chromatin to a certain size range. The conditions mentioned in Section 3.3 step 6 yield DNA fragments of 0.5–1 kb in size with a Sonic Dismembrator 550 (Fischer Scientific). As Quantitative PCR usually amplifies DNA fragments of 100–150 bp in size, the average fragment size after sonication should be > 150 bp in order to prevent a decrease in the amplification efficiency. 9. 10 and 25 ml aliquots of the supernatants recovered in Section 3.3 step 8 and Section 3.4 step 8, respectively, may be used for Western blot analysis to examine the presence of your protein of interest in WCE and immunoprecipitate, respectively. 10. The stringency of the ChIP procedure is determined by the composition of the lysis and wash buffers. Increasing salt and detergent concentrations will increase the stringency and reduce non-specific precipitation, yet may also be harmful for your antibody and reduce the immunoprecipitation efficiency. Varying the nature and concentration of salt and detergent may sometimes be necessary to determine the optimal conditions for the procedure for your protein of interest and application (depending on the protein up to 500 mM LiCl or NaCl can be used in place of 250 mM LiCl to reduce background). To test whether your protein of interest specifically precipitates a given DNA sequence, control experiments in which the antibody or crosslinking step is omitted can be performed. Alternatively, the immunoprecipitation from a tagged strain could be compared to that from a non-tagged strain. 11. After reversal of crosslinks in Section 3.4 step 9, the subsequent steps 10–15 for DNA recovery can be omitted and DNA can also be recovered without Protease K treatment using commercial DNA recovery kits, such as the Qiagen PCR cleanup kit.

Acknowledgments The authors would like to thank Professor Susan Gasser and members of the Gasser Laboratory for support during the development of these protocols. JC is an Alberta Heritage Foundation for Medical Research Scholar and work in the JC laboratory is funded by grants from the Canadian Institutes for Health Research # MOP-82736 and the Alberta Cancer Board # 23575. HvA is a Human Frontiers Science Program long-term fellow and work in HvA’s group is funded by a VIDI grant from the Netherlands Organisation for Scientific Research (NWO).

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References 1. Gribun A., Cheung K. L., Huen J., Ortega J., and Houry W. A. (2008) Yeast Rvb1 and Rvb2 are ATP-dependent DNA helicases that form a heterohexameric complex. J. Mol. Biol. 376, 1320–1333. 2. Shen X., Mizuguchi G., Hamiche A., and Wu C. (2000) A chromatin remodelling complex involved in transcription and DNA processing. Nature 406, 541–544. 3. Mizuguchi G., Shen X., Landry J., Wu W. H., Sen S., and Wu C. (2004) ATPdriven exchange of histone H2AZ variant catalyzed by SWR1 chromatin remodeling complex. Science 303, 343–348. 4. Krogan N. J., Keogh M. C., Datta N., et al. (2003) A Snf2 family ATPase complex required for recruitment of the histone H2A variant Htz1. Mol. Cell. 12, 1565–1576. 5. Kobor M. S., Venkatasubrahmanyam S., Meneghini M. D., et al. (2004) A protein complex containing the conserved Swi2/ Snf2-related ATPase Swr1p deposits histone variant H2A.Z into euchromatin. PLoS Biol. 2, E131. 6. van Attikum H., Fritsch O., Hohn B., and Gasser S. M. (2004) Recruitment of the INO80 complex by H2A phosphorylation links ATP-dependent chromatin remodeling with DNA double-strand break repair. Cell 119, 777–788. 7. Morrison A. J., Highland J., Krogan N. J., et al. (2004) INO80 and gamma-H2AX interaction links ATP-dependent chromatin remodeling to DNA damage repair. Cell 119, 767–775. 8. van Attikum H., Fritsch O., and Gasser S. M. (2007) Distinct roles for SWR1 and INO80 chromatin remodeling complexes at chromosomal double-strand breaks. EMBO J. 26, 4113–4125. 9. Shimada K., Oma Y., Schleker T., et al. (2008) Ino80 chromatin remodeling complex promotes recovery of stalled replication forks. Curr. Biol. 18, 566–575. 10. Papamichos-Chronakis M., and Peterson C. L. (2008) The Ino80 chromatin-remodeling enzyme regulates replisome function and stability. Nat. Struct. Mol. Biol. 15, 338–345. 11. Jonsson Z. O., Jha S., Wohlschlegel J. A., and Dutta A. (2004) Rvb1p/Rvb2p recruit Arp5p and assemble a functional Ino80 chromatin remodeling complex. Mol. Cell. 16, 465–477.

12. Biswas-Fiss E. E., Khopde S. M., and Biswas S. B. (2005) The Mcm467 complex of Saccharomyces cerevisiae is preferentially activated by autonomously replicating DNA sequences. Biochemistry 44, 2916–2925. 13. Davey M. J., Indiani C., and O’Donnell M. (2003) Reconstitution of the Mcm2-7p heterohexamer, subunit arrangement, and ATP site architecture. J. Biol. Chem. 278, 4491–4499. 14. Maiorano D., Lutzmann M., and Mechali M. (2006) MCM proteins and DNA replication. Curr. Opin. Cell. Biol. 18, 130–136. 15. Aparicio O. M., Weinstein D. M., and Bell S. P. (1997) Components and dynamics of DNA replication complexes in Saccharomyces cerevisiae: redistribution of MCM proteins and Cdc45p during S phase. Cell 91, 59–69. 16. Cobb J. A., Bjergbaek L., Shimada K., Frei C., and Gasser S. M. (2003) DNA polymerase stabilization at stalled replication forks requires Mec1 and the RecQ helicase Sgs1. EMBO J. 22, 4325–4336. 17. Bjergbaek L., Cobb J. A., Tsai-Pflugfelder M., and Gasser S. M. (2005) Mechanistically distinct roles for Sgs1p in checkpoint activation and replication fork maintenance. EMBO J. 24, 405–417. 18. Cobb J. A., Schleker T., Rojas V., Bjergbaek L., Tercero J. A., and Gasser S. M. (2005) Replisome instability, fork collapse, and gross chromosomal rearrangements arise synergistically from Mec1 kinase and RecQ helicase mutations. Genes Dev. 19, 3055–3069. 19. Lee S. E., Moore J. K., Holmes A., Umezu K., Kolodner R. D., and Haber J. E. (1998) Saccharomyces Ku70, mre11/rad50 and RPA proteins regulate adaptation to G2/ M arrest after DNA damage. Cell 94, 399–409. 20. Dubrana K., van Attikum H., Hediger F., and Gasser S. M. (2007) The processing of double-strand breaks and binding of single-strand-binding proteins RPA and Rad51 modulate the formation of ATRkinase foci in yeast. J. Cell. Sci. 120, 4209–4220. 21. van Werven F. J., and Timmers H. T. (2006) The use of biotin tagging in Saccharomyces cerevisiae improves the sensitivity of chromatin immunoprecipitation. Nucleic Acids Res. 34, e33.

Chapter 9 Methods to Study How Replication Fork Helicases Unwind DNA Daniel L. Kaplan and Irina Bruck Abstract Replication fork helicases unwind DNA at a replication fork, providing polymerases with single-stranded DNA templates for replication. In bacteria, DnaB unwinds DNA at a replication fork, while in archaeal and eukaryotic organisms the Mcm proteins catalyze replication fork unwinding. Unwinding in archaea is catalyzed by a single Mcm protein that forms multimeric rings, whereas eukaryotic helicase activity is catalyzed by the heterohexameric Mcm2–7 complex acting in concert with Cdc45 and the GINS complex. A subcomplex of eukaryotic Mcm proteins, the Mcm4,6,7 complex, unwinds DNA in vitro, and studies of this assembly reveal insight into the mechanism of the eukaryotic Mcm helicase. Detailed methods for the investigation of replication fork helicase mechanism are described in this chapter. Described herein are methods for the design of DNA substrates for unwinding and branch migration studies, annealing DNA, purifying replication fork helicase proteins, and analyzing DNA unwinding activity. Key words: Helicase, unwinding, DNA replication, mechanism, Mcm, DnaB, method, minichromosome maintenance, replication fork, annealing.

1. Introduction 1.1. Replication Fork Helicase Function

DnaB catalyzes DNA unwinding at a replication fork in bacteria, whereas the minichromosome maintenance proteins (Mcms) catalyze DNA unwinding during chromosomal replication in eukaryotes (1–5) and in archaea (6–8). In archaea, there is a single Mcm protein that forms ring-shaped structures that are generally hexamers or double-hexamers (8–11). Since the discovery of archaeal Mcm helicase activity by three different groups (6–8), there have been detailed mechanistic studies of the archaeal Mcms. Since

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many of the sequence motifs of the archaeal Mcms are conserved in eukaryotic Mcms, these studies are important for understanding archaeal and eukaryotic Mcm mechanisms. In eukaryotes, a heterohexameric ring composed of Mcm2, Mcm3, Mcm4, Mcm5, Mcm6, and Mcm7 (Mcm2–7) acting in concert with Cdc45 and the GINS complex unwinds DNA at a replication fork (1–5). Biochemical studies of Mcm2–7 have thus far been limited. Recently, an assembly composed of Mcm2–7, Cdc45, and GINS purified from Drosophila embryo extracts exhibited helicase activity in vitro (4), and Mcm2–7 alone from budding yeast has helicase activity in vitro under certain conditions (5). In contrast, a more extensive literature exists on a subcomplex of eukaryotic Mcm proteins composed of Mcm4, Mcm6, and Mcm7 (Mcm4,6,7) (12–16). The Mcm4,6,7 complex has helicase activity in vitro in many different species, including Xenopus, mouse, budding yeast, and fission yeast (12–16). Insight into Mcm2–7 mechanism can likely be learned from studies of the Mcm4,6,7 complex, and therefore these Mcm4,6,7 mechanistic studies are very important in the understanding of eukaryotic replication fork helicase mechanism. 1.2. Design of DNA Substrates for Unwinding and Branch Migration Studies

There are many important considerations for designing a DNA substrate to monitor helicase unwinding activity. One consideration in the choice of DNA substrate is that of DNA length. Helicases that are highly processive will unwind DNA that is long, whereas helicases that are not very processive require short DNA stretches to observe unwinding. A second consideration for the construction of a DNA substrate is that of melting temperature. It is important that the melting temperature of the DNA substrate is higher than the incubation temperature of the unwinding assay. For example, in studies of Thermus aquaticus DnaB, the enzyme is most active at 55C, and the melting temperature of the DNA substrate used was 15C higher, at 70C (17). The DNA substrate used in these studies is shown in Fig. 9.1. Notice that although this DNA substrate is only 22 bp long, it has a high melting temperature because the GC content is high. The DNA reannealing time is a third consideration for the design of a DNA substrate for unwinding studies. If the DNA reanneals faster than the helicase unwinds, no unwinding will be observed. Therefore, if a helicase is very slow, the DNA reannealing time should be long. In studies with the replication fork helicase DnaB from T. aquaticus, the half-time for reannealing of the DNA substrate was approximately 30 min (17, 18). For Saccharomyces cerevisiae Mcm4,6,7 studies, the DNA substrate has a half-time for reannealing of 75 min (Fig. 9.1), which is substantially longer than the 16 min incubation time used in the unwinding assays (15). It is also important to study the reannealing time in the presence of the helicase, since the enzyme may

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Fig. 9.1. DNA substrates used in replication fork helicase unwinding studies. (a) This substrate has a high GC content and was used to study T. aquaticus DnaB activity at 55C. (b) This substrate was used to study S. cerevisiae Mcm4,6,7 activity at 37C.

affect the reannealing rate. To accomplish this, one must start with a DNA substrate that the helicase cannot unwind, and then directly measure reannealing time in the presence of protein and a nucleoside triphosphate. For T. aquaticus DnaB, the presence of helicase increased the half-time for reannealing of the DNA substrate from 30 to 120 min (17). A related consideration is that of local reannealing. In this process, the helicase unwinds the DNA strands, but as the helicase continues to progress, the DNA strands reanneal behind the DNA helicase because of the high local concentration of the DNA strands. A final consideration for the design of a DNA substrate for a helicase assay is that of GC content. A substrate with a high GC content was used to study T. aquaticus DnaB (17, 18), while a substrate with a lower GC content was used to study Escherichia DnaB and S. cerevisiae Mcm4,6,7 (Fig. 9.1) (15, 19, 20). E. coli DnaB and S. cerevisiae Mcm4,6,7 can also drive branch migration of four-way DNA Holiday junctions (15, 19, 20). When studying branch migration, it is important to distinguish bona fide branch migration activity from unwinding activity. For example, in Fig. 9.2, the Holiday junction used to study E. coli DnaB activity has four DNA strands, labeled *1, 2, 3, and 4. Strand *1 is radiolabeled (*). If bona fide branch migration occurs, the *1,4 product will accumulate. In contrast, if unwinding occurs, the *1 product will accumulate. However, if strand 4 is also unwound by

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Fig. 9.2. E. coli DnaB drives branch migration of this 4-way DNA junction, producing the *1,4 duplex. Strand 1 is radiolabeled in this assay (*).

the enzyme, and if the *1 and 4 strands reanneal with one another, then the *1,4 product will accumulate. Thus, the accumulation of *1,4 product may be the result of DNA unwinding followed by reannealing, and not bona fide branch migration, if the reannealing rate of *1 and 4 is rapid relative to the time course of the assay. In the Holiday junctions used to study DnaB and Mcm4,6,7, the half-time for reannealing strand *1 and strand 4 is 75 min, far longer than the incubation time of the assay (Fig. 9.2) (15, 19– 21). DNA strands have widely variable annealing times; thus, the empirical study of DNA reannealing time is critical for the proper investigation of DNA branch migration activity. In this chapter we describe methods for annealing DNA, purifying replication fork helicase proteins, and analyzing DNA helicase activity.

2. Materials 2.1. Conditions for Annealing DNA

1. T4 polynucleotide kinase (New England Biolabs). 2.

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P-g–ATP (NEN-Perkin Elmer).

3. TE: 10 mM Tris–HCl, pH 8.0, 0.1 mM EDTA. 4. G-25 sephadex column (Roche). 5. 6 loading dye: 15% Ficoll and 0.1% Xylene Cyanol.

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6. 5 EDBG þ Mg: 100 mM Tris–HCl, pH 7.5, 20% glycerol, 0.5 mM EDTA, 200 mg/ml BSA, 25 mM DTT, and 50 mM magnesium acetate. All reagents listed above are stored at –20C. 2.2. Replication Fork Helicase Protein Preparation

1. Buffer A: 10% sucrose, 50 mM Tris–HCl, pH 7.5, 50 mM NaCl, 5 mM DTT, and two tablets of complete, EDTA-free, protease inhibitor cocktail (Boehringer Mannheim). 2. Buffer B: 10% glycerol, 50 mM Tris–HCl, pH 7.5, 50 mM NaCl, 5 mM DTT, and 5 mM MgCl2. 3. Buffer C: 10% glycerol, 20 mM Tris–HCl, pH 7.5, 50 mM NaCl, 5 mM DTT, 5 mM MgCl2, and 0.02% Na Azide. 4. Buffer D: 10% glycerol, 10 mM Tris–HCl, pH 7.5, 100 mM NaCl, 5 mM DTT, 5 mM MgCl2, and 0.02% Na Azide.

2.3. Unwinding Assay

1. MMM: mix3.2 ml of 5 EDBG+Mg, 3.2 ml of 80% glycerol, 0.8 ml of 100 mM ATP (make sure the ATP is freshly made), 160 ml of 0.5 M creatine phosphate (stored at –80C), and 320 ml of 1 mg/ml creatine kinase (stored at –80C); store in aliquots at –80C. 2. 4 Stop buffer: 2% SDS and 80 mM EDTA; store at room temperature. 3. Proteinase K (Sigma-Aldrich).

3. Methods 3.1. Conditions for Annealing DNA

1. T4 polynucleotide kinase is used to radiolabel 50 pmoles of the DNA strand of interest (S460T, Fig. 9.1). 2. The kinase reaction contains 2 ml of 10 kinase buffer, 5 ml of 10 mM S460T (Fig. 9.1), 7 ml of H2O, 5 ml 32P-g–ATP, and 1 ml of T4 polynucleotide kinase. 3. The kinase reaction is allowed to proceed for 30 min at 37C. 4. The kinase is then denatured by heat inactivation to 65C for 20 min. 5. The unincorporated 32P-g–ATP is then removed with a G-25 sephadex spin column. 6. TE is then added to the *S460T to achieve a final volume of 100 ml, for a final concentration of 500 nM (see Note1). 7. This sample is then serially diluted to run as a marker in the gel. Four microliters of 500 nM *S460T are mixed with 96 ml TE to yield 100 ml of 20 nM *S460T.

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8. 51 ml of TE are then mixed with 4 ml 20 nM *S460T and 25 ml 6 loading dye to yield 80 ml 1 nM *S460T in dye (run in gel as marker). 9. The radiolabeled DNA stand is then mixed in a 1:2 ratio with the unlabeled, complementary strand. 10. 4 ml of 5 EDBG þ Mg are mixed with 4 ml of 500 nM *S460T and 4 ml of 1 mM 60T1S (complementary DNA strand, Fig. 9.1). 11. The sample is then annealed at the assay temperature for 4 h (see Notes 2 and 3). 12. 38 ml TE is then added to achieve a final concentration of 40 nM (working stock of *S460T/60T1S). This 40 nM stock is used for the helicase assay. This sample is also serially diluted to use as marker in gel. 13. 53 ml TE are mixed with 2 ml 40 nM *S460T/60T1S and 25 ml 6 loading dye to yield 80 ml of 1 nM *S460T/60T1S in dye (run in gel as marker). 3.2. Replication Fork Helicase Protein Preparation 3.2.1. Preparation of DnaB from Thermus aquaticus

1. The E. coli BL21 cells are transformed with the pET22b vector, which contains the T. aquaticus dnaB helicase gene. 2. The transformed cells are grown in 10 l of LB media containing 50 mg/ml ampicillin in a stirred, aerated fermentor at 37C. When the cells reach as A600 of 0.6, the temperature is decreased to 30C and isopropyl-b-D-thiogalactopyranoside is added to a final concentration of 100 mM to induce gene expression. 3. Three hours later, cells are harvested by centrifugation (typically yield is 34 g). From this point, all manipulations are carried out at 4C unless otherwise stated. 4. The cells are resuspended in 100 ml of buffer A. The cells are then lysed with a Microfluidizer (Microfluidics Corp.). MgCl2 is then added to a final concentration of 5 mM. 5. The lysate is then heated at 65C for 20 min followed by chilling on ice for 20 min. The cells are then spun at 38,000 RPM in a Ti45 rotor (Beckman) for 60 min. 6. The supernatant is filtered with a 0.22-mm low protein-binding filter (Fraction I). A Q source column (50 ml, Pharmacia) is pre-equilibrated with buffer B. 7. Fraction I (130 ml) is loaded onto the column and the column is then washed with 5 column volumes of buffer B. 8. The protein is eluted with a linear gradient (1 l) of buffer B (containing 0.1 mM PMSF) to buffer B containing an additional 450 mM NaCl. DnaB helicase elutes at 300–345 mM NaCl (fraction II).

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9. Fraction II (250 ml) is dialyzed against buffer C. Following dialysis the sample is loaded onto a series of five 5 ml HiTrap heparin sepharose columns (Pharmacia) that are preequilibrated with buffer C. 10. The columns are then washed with 5 column volumes of buffer C followed by elution with a linear gradient (500 ml) of buffer C to buffer C containing an additional 450 mM NaCl. DnaB helicase elutes between 200 and 250 mM NaCl (fraction III). 11. Fraction III (125 ml) is dialyzed against buffer D. After dialysis the sample is concentrated with an Amicon ultrafiltration device to a final volume of 13 ml and then loaded onto a Superdex 200PG column (320 ml, Pharmacia) that is pre-equilibrated with buffer D. The sample elutes as a single peak at a ratio of elution volume to void volume of 1.16 (fraction IV). 12. Fraction IV (60 ml) is then concentrated with an Amicon ultrafiltration device followed by an Amicon Centriplus device until the concentration of protein is 11 mg/ml. The final yield is 112 mg of protein from a 10-l cell paste. Aliquots are snap-frozen in liquid nitrogen and stored at –80C. When ready for use, an aliquot is rapidly thawed. If the protein is going to be used in a biochemical assay, 80% glycerol is added to the sample to achieve a final concentration of 50% glycerol. Samples containing 50% glycerol are stored at –20C for up to 1 week. 3.2.2. Purification of E. coli DnaB and S. cerevisiae Mcms

3.3. Unwinding Assay

Purification of E. coli DnaB and S. cerevisiae Mcms have been described previously in detail (15, 22, 23).

1. Fresh master mix (MM) is made by mixing 145 ml MMM with 5 ml 40 nM *S460T/60T1S DNA substrate (1 nM final concentration). Eight microliters MM is added to each reaction tube on ice. Three microliters of helicase protein is then added. 2. The sample is mixed and then incubated at the reaction temperature for the desired time. When the reaction is complete, 1 ml of 10 mg/ml proteinase K is added and the sample is then incubated an additional 1 min at 37C. 3. The sample is then mixed with 5 ml of 4 stop buffer, and then 5 ml of 6 loading dye is added and mixed. The sample is snap frozen on dry ice. 4. Samples are thawed in a room temperature water bath for 4 min (see Note 4). 5. The samples are then analyzed with an 8% Polyacrylamide gel containing 1 TBE.

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6. The samples are electrophoresed at 175 V until the dye front approaches the bottom of the gel. 7. The gel is then dried with a gel dryer for 1 h at 80C. The gel is then exposed to a phosphorimaging screen.

4. Notes 1. 65 ml TE is typically added to 35 ml of DNA. 2. Pre-equilibrating the annealed DNA at the assay temperature ensures that the fraction of radiolabeled single-stranded DNA will not change during the unwinding assay in the absence of protein. 3. If a water bath is used, the sample should be completely submerged to prevent accumulation of water at the top of the tube. 4. This incubation step reduces intra-strand base pairing.

Acknowledgments The authors thank Dr. Thomas A. Steitz and Dr. Mike O’Donnell for their support and encouragement. References 1. Labib K., Tercero J. A., and Diffley J. F. X. (2000) Uninterrupted MCM2–7 function required for DNA replication fork progression. Science 288, 1643–1647. 2. Pacek M., Tutter A., Kubota Y., Takisawa H., and Walter J. (2006) Localization of MCM2–7, Cdc45, and GINS to the site of DNA unwinding during eukaryotic DNA replication. Mol. Cell 21, 581–587. 3. Calzada A., Hodgson B., Kanemaki M., Bueno A., and Labib K. (2005) Molecular anatomy and regulation of a stable replisome at a paused eukaryotic DNA replication fork. Genes Dev. 19, 1905–1919. 4. Moyer S., Lewis P., and Botchan M. (2006) Isolation of the Cdc45/ Mcm2–7/GINS (CMG) complex, a candidate for the eukaryotic DNA replication fork helicase. Proc. Natl. Acad. Sci. USA 103, 10236–10241.

5. Bochman M. and Schwacha A. (2008) The Mcm2-7 complex has in vitro helicase activity. Mol. Cell 31, 287–293. 6. Shechter D. F., Ying C. Y., and Gautier J. (2000) The intrinsic DNA helicase activity of Methanobacterium thermoautotrohicum delta H minichromosome maintenance protein. J. Biol. Chem. 275, 15049–15059. 7. Kelman Z., Lee J.-K., and Hurwitz J. (1999) The single minichromosome maintenance protein of Methanobacterium thermoautotrophicum H contains DNA helicase activity. Proc. Natl. Acad. Sci. USA 96, 14783–14788. 8. Chong J. P. J., Hayashi M. K., Simon M. N., Xu R.-M., and Stillman B. (2000) A doublehexamer archaeal minichromosome maintenance protein is an ATP-dependent DNA helicase. Proc. Natl. Acad. Sci. U.S.A. 97, 1530–1535.

Study How Replication Fork Helicases Unwind DNA 9. Costa A., Pape T., van Heel M., Brick P., Patwardhan A., and Onesti S. (2006) Structural basis of the Methanothermobacter thermautotrophicus MCM helicase activity. Nucleic Acids Res. 34, 5829–5838. 10. Gomez-Llorente Y., Fletcher R., Chex X., Carazo X., and San Martin C. (2005) Polymorphism and double hexamer structure in the archaeal minichromosome maintenance (MCM) helicase from Methanobacterium thermoautrophicum. J. Biol. Chem. 280, 40909–40915. 11. Fletcher R., Shen J., Gomez-Llorente Y., Martin C. S., Carazo J. M., and Chen X. (2005) Double hexamer disruption and biochemical activities of Methanobacterium thermoautotrophicum MCM. J. Biol. Chem. 280, 42405–42410. 12. Ishimi Y. (1997) A DNA helicase activity is associated with an MCM4, -6, and -7 protein complex. J. Biol. Chem. 272, 24508–24513. 13. Lee J.-K., and Hurwitz J. (2000) Isolation and characterization of various complexes of the minichromosome maintenance proteins of Schizosaccharomyces pombe. J. Biol. Chem. 275, 18871–18878. 14. You Z., Komamura Y., and Ishimi Y. (1999) Biochemical analysis of the intrinsic Mcm4Mcm6-Mcm7 DNA helicase activity. Mol. Cell. Biol. 19, 8003–8015. 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.

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16. Lee J.-K., and Hurwitz J. (2001) Processive DNA helicase activity of the minichromosome maintenance proteins 4,6, and 7 complex requires forked DNA structures. Proc. Natl. Acad. Sci. USA 98, 54–59. 17. Kaplan D. L., and Steitz T. A. (1999) DnaB from Thermus aquaticus unwinds forked duplex DNA with an asymmetric tail length dependence. J. Biol. Chem. 274, 6889–6897. 18. 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. 19. Kaplan D. L., and O’Donnell M. (2002) DnaB drives DNA branch migration and dislodges proteins while encircling two DNA strands. Mol. Cell 10, 647–657. 20. Kaplan D. L., and O’Donnell M. (2004) Twin DNA pumps of a hexameric helicase provide power to simultaneously melt two duplexes. Mol. Cell 15, 453–465. 21. Kaplan D. and O’Donnell M. (2006) RuvA is a sliding collar that protects holliday junctions from unwinding while promoting branch migration. J. Mol. Biol. 355, 473–490. 22. Yuzhakov A., Turner J., and O’Donnell M. (1996) Replisome assembly reveals the basis for asymmetric function in leading and lagging strand replication. Cell 86, 877–886. 23. Davey M. J., Indiani C., and O’Donnell M. (2003) Reconstitution of the Mcm2–7p heterohexamer, subunit arrangement, and ATP site architecture. J. Biol. Chem. 278, 4491–4499.

Chapter 10 Simple Enzymatic Assays for the In Vitro Motor Activity of Transcription Termination Factor Rho from Escherichia coli Marc Boudvillain, Ce´line Walmacq, Annie Schwartz, and Fre´de´rique Jacquinot Abstract The transcription termination factor Rho from Escherichia coli is a ring-shaped homo-hexameric protein that preferentially interacts with naked cytosine-rich Rut (Rho utilization) regions of nascent RNA transcripts. Once bound to the RNA chain, Rho uses ATP as an energy source to produce mechanical work and disruptive forces that ultimately lead to the dissociation of the ternary transcription complex. Although transcription termination assays have been useful to study Rho activity in various experimental contexts, they do not report directly on Rho mechanisms and kinetics. Here, we describe complementary ATP-dependent RNA–DNA helicase and streptavidin displacement assays that can be used to monitor in vitro Rho’s motor activity in a more direct and quantitative manner. Key words: Rho, transcription, termination, ring shaped, helicase, hexamer, molecular motor.

1. Introduction In Escherichia coli, about half of the transcription termination events require the participation of the endogenous Rho protein. Since its discovery in 1969 (1), a considerable amount of work has been devoted to the function, structure, and mechanisms of Rho (2, 3). The active form of the Rho factor is a complex of six protomers arranged in a ring-shaped configuration (4, 5). In vitro, Rho hexamers exhibit enzymatic activities that are characteristic of NTP-driven nucleic acid motors such as RNA-dependent ATPase (6) and ATP-dependent helicase activities (7). Rho preferentially binds to cytosine-rich, scarcely structured, nucleic acids M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_10, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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(NAs) including poly(C) oligomers and Rut (Rho utilization) segments of nascent transcripts (8–11). This is due to the presence of a binding pocket on each Rho monomer that can specifically accommodate a 50 -YC dimer (Y and C being a pyrimidine and a cytosine, respectively) (12). Altogether the six binding pockets form a crownlike primary interaction site (4, 5) that anchors, somewhat specifically, the NA chain to the Rho factor. Once bound, C-rich RNA strands, but not DNA strands, can trigger the ATPase activity of Rho (9). It is believed that this differential activation of Rho’s ATPase is due to specific binding of RNA to a secondary interaction site within the hexamer central channel (5, 13). A productive Rhotranscript complex should therefore adopt a topology such as the one depicted in Fig. 10.1a. This configuration has important mechanistic implications that cannot be tested easily with classical transcription termination assays. This is because, in such assays, transcription requirements complicate the manipulation of experimental parameters for structure–function and mechanistic studies and because individual thermo-kinetic parameters cannot be measured precisely and attributed unambiguously to specific steps of the Rho-dependent termination process. To get around these problems, we have developed complementary ATP-dependent RNA– DNA helicase and streptavidin displacement assays that can be used to analyze and quantitate Rho’s motor activity in isolation (14–17). The assays rely on the use of composite NA substrates that contain a well-defined synthetic Rut sequence (aRut; Fig. 10.1b) (8) and that can be readily prepared and manipulated by classical molecular biology techniques in order to explore the mechanistic framework of the Rho enzyme.

2. Materials 2.1. Expression and Purification of the Rho Enzyme

1. LB broth: dissolve LB powder (Sigma-Aldrich) at 20 mg/mL in water. Prepare a 500-mL aliquot in a 2-L baffled Erlenmeyer culture flask and a 250-mL aliquot in an autoclavable bottle. Sterilize by autoclaving and store at room temperature. 2. Kanamycin (Sigma-Aldrich) stock solution ( 1000) at 50 mg/mL in water. Sterilize with a 0.22 mm filter and store at –20C. 3. Chloramphenicol (Sigma-Aldrich) stock solution ( 1000) at 50 mg/mL in ethanol. 4. 20% glycerol stock of E. coli BL21(DE3)pLysS cells (Stratagene) transformed with the pEt28b-Rho plasmid (kindly provided by Pr. J.M. Berger, University of California at Berkeley). Store at –80C.

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Fig. 10.1. Rho-dependent termination of transcription. (a) Schematic of the interaction between a Rho hexamer and the nascent RNA transcript. Each Rho monomer bears a primary interaction sub-site (white oval ) that can bind specifically a 50 -YC dimer of the Rut region (in grey ) in order to anchor the transcript to the hexamer. The RNA chain subsequently interacts with Rho’s secondary binding site, which is located within the hexamer central channel in the vicinity of the ATPase pocket (5). (b) Design of composite substrates for testing the in vitro RNA–DNA helicase and streptavidin displacement activities of Rho. The DNA template contains the aRut sequence (in grey ) inserted between a promoter for T7 RNA polymerase and a reporter cassette. The reporter cassette should encode for a single-stranded RNA segment of > 15 nts (14) followed by another RNA segment which is complementary to the DNA oligonucleotide (helicase assay) or to which is linked the biotin moiety (streptavidin displacement assay).

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5. IPTG solution: 1 M isopropyl b-d-thiogalactopyranoside (Fluka) in ethanol. Store at –20C. 6. Lysis buffer: 0.23 M NaCl, 50 mM Tris–HCl, pH 7.5, 5 mM EDTA, 0.1 mM DTT, 1 mM b-mercaptoethanol, 5% (v/v) glycerol. Sterilize with a 0.22-mm filter and store at 4C. 7. PMSF stock solution: 0.1 M phenylmethanesulfonyl fluoride (Sigma-Aldrich) in ethanol. Store at –20C. 8. Lysozyme from chicken egg white (Sigma) is dissolved at 10 mg/mL in water. Store at 4C. 9. Sodium deoxycholate (Sigma-Aldrich) at 5% (w/v) in water. Store at room temperature. 10. DnaseI from bovine pancreas (Roche). 11. 1 M MgCl2 solution (see Note 1). 12. Polymin-P stock solution: 0.2 M NaCl, 0.1 mM EDTA, 10% (v/v) poly[ethyleneImine] solution (Fluka), 5% (v/v) glycerol. The solution is adjusted to pH 8.0 with HCl, filtered on Whatman paper, and dialyzed twice against 1 L of TGEN buffer (5% [v/v] glycerol, 10 mM Tris–HCl, pH 8.0, 0.1 mM EDTA, 0.2 M NaCl) for 10 h at 4C. Store in the dark and at room temperature. 13. Ammonium sulfate (Sigma-Aldrich) (see Note 1). 14. 25-mL HiLoad SP Sepharose FF and 5-mL HiTrap heparin HP columns (GE Healthcare). Store the columns filled with 20% ethanol at 4C. 15. Buffer A (low salt): 10 mM Tris–HCl, pH 7.6, 0.1 mM EDTA, 0.1 mM DTT, 5% (v/v) glycerol, 0.1 M NaCl (see Note 2). 16. Buffer B (high salt): 10 mM Tris–HCl, pH 7.6, 0.1 mM EDTA, 0.1 mM DTT, 5% (v/v) glycerol, 1 M NaCl (see Note 2). 17. Storage buffer: 0.2 M KCl, 20 mM Tris–HCl, pH 7.9, 5% (v/v) glycerol, 0.2 mM EDTA, 0.2 mM DTT. 18. 99% glycerol (Sigma-Aldrich). 2.2. Preparation of Nucleic Acid Substrates

1. RNA-grade water (see Note 3). M10E1 buffer: 10 mM MOPS, pH 6.0, 1 mM EDTA (see Note 4). 2. Linearized plasmid, PCR-amplified DNA template, or chemically synthesized double-stranded oligodesoxyribonucleotides containing the aRut sequence downstream from a T7 promoter (Fig. 10.1b) at a concentration of 0.5 mM in M10E1 buffer (see Note 4). 3. 100 mM rNTP set (GE Healtcare). 4. T7 RNA polymerase at 200 U/mL (Epicentre), Superasin at 20 U/mL (Ambion), and RQ1 DNase at 1 U/mL (Promega).

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5. Transcription buffer ( 5): 0.12 M MgCl2, 0.4 M HEPES, pH 7.5, 0.1 M DTT, 0.05% Triton X-100 (Sigma), 5 mM Spermidine (see Note 4). 6. 3 M sodium acetate, pH 6. 7. 0.5 M EDTA, pH 7.5. 8. TBE buffer ( 10): 0.89 M Tris base, 0.89 M boric acid, 0.02 M EDTA. Filter on Whatman paper and store at room temperature. 9. Denaturing loading buffer: 95% formamide, 5 mM EDTA, 0.01% (w/v) xylene cyanol, 0.01% (w/v) bromophenol blue. 10. Denaturing acrylamide solution: 6% Acrylamide:bis-acrylamide (19:1 ratio; Interchim) and 7 M urea in 1 TBE buffer. Heat the solution to dissolve urea completely and cool down to room temperature. Prepare 30 mL of fresh solution right before use. 11. N,N,N,N0 -tetramethyl-ethylenediamine (TEMED, SigmaAldrich) and 10% (w/v) ammonium persulfate (APS) in water. 12. Elution buffer ( 1): a 1:9 (v/v) mixture of 3 M NaAc:M10E1 buffer (see Note 4). 13. Alkaline phosphatase (AP) from calf intestine at 1 U/mL (Roche diagnostics). 14. Extraction solutions: phenol:chloroform:isoamyl alcohol (25:24:1) mix saturated with 10 mM Tris, pH 8.0, 1 mM EDTA (Sigma-Aldrich) and chloroform (Sigma-Aldrich) saturated with 0.1 volume of M10E1 buffer. Store the solutions at 4C. 15.

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P-gATP (3000 Ci/mmol [10 mCi/mL]; see Note 5).

16. T4 polynucleotide kinase at 10 U/mL with 10 kinase reaction buffer (New England Biolabs). 17. G-50 Quick Spin columns (Roche). 18. Annealing buffer ( 10): 1.5 M KAc, 0.2 M HEPES, pH 7.5, 0.01 M EDTA (see Note 4). 19. Complementary oligodesoxyribonucleotide for pairing to the RNA transcript downstream from the aRut region (helicase substrates; see Fig. 10.1b). 20. Native loading buffer ( 5): 25% [w/v] Ficoll-400 (SigmaAldrich), 25 mM EDTA, 0.05% (w/v) xylene cyanol, 0.05% (w/v) bromophenol blue. 21. Native acrylamide solution: 9% acrylamide:bis-acrylamide (19:1 ratio; Interchim) in 1 TBE buffer. Prepare 30 mL of solution right before use.

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22. Autoradiography BioMax XAR films (Kodak) with exposure cassette and developer and fixer baths (Sigma-Aldrich). Store and manipulate in a dark room equipped with a dim red light. The following materials are only for the preparation of biotinylated RNA substrates: 23. Oxidation buffer: 0.3 M sodium acetate, pH 5.0. 24. Sodium periodate (Sigma-Aldrich) at 0.2 M in water. Prepare a fresh solution right before use. 25. 3 M KCl solution. 26. Dissolve biotin hydrazide (Pierce) at 50 mM in anhydrous DMSO (Sigma-Aldrich). Aliquots (20 mL) are stored at –20C. 27. Core streptavidin (Promega) at 4 mM in water (see Note 4). 28. Bovine serum albumin (BSA) at 10 mg/mL (New England Biolabs). 29. TBES buffer ( 10): 0.89 M Tris base, 0.89 M boric acid, 0.02 M EDTA, 3% (w/v) SDS. 30. Helicase gel solution: 7.5% Acrylamide:bis-acrylamide (19:1 ratio; Interchim) in 1 TBES buffer. Prepare 30 mL of fresh solution right before use. 2.3. Helicase and Streptavidin Displacement Assays

1. Rho stock solution (see Note 6). 2. ‘Standard’ helicase buffer ( 10): 1.5 M KAc, 0.2 M HEPES, pH 7.5, 1 mM EDTA, 5 mM DTT (see Note 4). 3. Poly[rC] (GE Healtcare) is dissolved at 10 mg/mL in water (see Note 3) and loaded on a 100-kDa Centricon column (Millipore). The column is repeatedly (at least 5 times) filled with 1 mL of water and centrifuged at 1000 g until the column volume is reduced to 100 mL (but avoid drying of the membrane). The solution in the bottom reservoir, which contains the small poly[rC] fragments that have flown through the membrane, is removed after each centrifugation cycle. The top reservoir is then equipped with its recovery cap and centrifugated at 1000 g in an inverted position to collect the solution containing long (>300 nucleotides [nt]) poly[rC] fragments. The concentration of the poly[rC] solution is determined by UV absorbance (em,260[rC] 25 (g/l)–1 cm–1) and adjusted to 10 mg/mL with water. 4. Initiation mix ( 10): 10 mM ATP, 10 mM MgCl2 in 1 standard helicase buffer. The solution is supplemented with 5 mM Trap oligodesoxyribonucleotide (whose sequence is complementary to the one of the DNA strand of the RNA–DNA hybrid) or 100 mM biotin for helicase or streptavidin displacement reactions, respectively. For single-run helicase reactions, 3 mg/mL poly[rC] is also added to the initiation mix.

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5. Quench buffer: 0.1 M EDTA, 0.5% (w/v) sodium dodecyl sulphate (SDS), 7% Ficoll-400 (Sigma-Aldrich). 6. TBES buffer ( 10) and Helicase gel solution as described in the preceding section.

3. Methods 3.1. Preparation of Active Rho Factor

The pEt28b-Rho plasmid was initially designed by the Berger lab to prepare large amounts of purified Rho protein for crystallographic studies (4). To this end, a Met-Gly-His triplet has been added at the beginning of Rho’s open reading frame in order to maximize over-expression of the protein in BL21(DE3)pLysS/ pEt28b-Rho cells (Fig. 10.2a). However, MALDI-TOF mass spectrometry indicates that the first methionine is absent from the purified Rho variant while the remaining amino acids (Gly-His) have no detectable effect on RNA binding or ATPase, helicase, and streptavidin displacement activities (F.J., A.R.R., and M.B., unpublished data). We thus now routinely

Fig. 10.2. Preparation of the Rho enzyme. (a) IPTG-induced overexpression of Rho variants in BL21(DE3)pLysS cells containing the pEt28b-Rho, pEt28b-RhoMGH, or pCB111 (kindly provided by Pr. J. Richardson, Indiana University) plasmid. The pEt28b-RhoMGH and pCB111 plasmids encode the sequence of wild-type Rho while pEt28b-Rho encodes for a protein variant with an N-terminal Gly-His addendum (see text). (b) Isolation of the Rho protein by cation-exchange chromatography on a SP sepharose column (first purification step). A representative chromatogram is shown together with SDS-PAGE analysis of a selection of collected fractions. (c) Rho purification (second step) by affinity chromatography on a heparin column.

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use the pEt28b-Rho plasmid to prepare the purified Rho enzyme in high yields (50 mg/L of bacterial culture) for structure-function and mechanistic studies. The protocols for Rho preparation were adapted from Nowatzke et al. (18). Following cell lysis, the Rho protein is readily purified to >95% homogeneity by a two-step (ion exchange/affinity) chromatography procedure that assumes the use of a Pharmacia FPLC system equipped with motor valves for sample injection and column selection, with a 150-mL sample superloop, with conductimetry and UV (280 nm filter) monitors and with a FRAC-920 fraction collector. The FPLC apparatus (or any equivalent low pressure chromatographic system) and accessories should be kept at 4C in a cold room or refrigerated cabinet. 3.1.1. Production of Cell Paste and Isolation of Cellular Proteins

1. The glycerol stock of BL21(DE3)pLysS/pEt28b-Rho cells is used to inoculate 5 mL of LB containing 50 mg/mL kanamycin and 50 mg/mL chloramphenicol in a sterile 14-mL culture tube. The culture is grown overnight at 37C before being transferred to a 2-L Erlenmeyer flask containing 500 mL of the same sterile medium. The flask is incubated at 37C in a rotary shaker (300 rpm in most incubator shakers is sufficient for good aeration) until A600 reaches 0.6 (exponential phase; this usually takes 3 h). Then, 0.4 mM IPTG is added and the growth is continued for 3 h at 37C. Equal volumes of the culture are transferred into two 500-mL centrifuge bottles and centrifuged for 15 min at 8,000 g and 4C in a Sorvall SLA3000 (superlite) rotor. The supernatants are discarded and the pellets (3 grams total) stored at –20C. 2. Cell pellets are gently thawed in ice-cold lysis buffer (10 mL per gram of pellet) supplemented with 1 mM PMSF. Then, lysozyme is added (5 mg per gram of pellet) and the mixture is left 20 min at room temperature (it should become viscous during that time) before addition of 0.01 volume of 5% sodium deoxycholate. The mixture is gently mixed before further incubation for 7 min at room temperature and 15 min on ice. Then, MgCl2 (24 mM, final concentration) and DNaseI (4 mg/mL) are added and the mixture is incubated for 20 min on ice. One volume of lysis buffer is added before centrifugation for 15 min at 30,000 g and 4C in a Sorvall SS-34 rotor. The supernatant is recovered and 0.5 volume of Polymin-P solution is added to precipitate the bulk of nucleic acids. The mixture is gently stirred for 5 min on ice before further centrifugation for 15 min at 30,000 g. The supernatant is carefully recovered and ammonium sulfate (0.5 grams per mL of supernatant; see Note 1) is slowly added upon gentle stirring on ice. After 1 h of incubation on ice, the mixture is centrifuged for 20 min at 30,000 g and 4C. The pellet is dissolved in 40 mL of buffer A and dialyzed twice against 1 L of the same buffer at 4C.

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1. To eliminate any insoluble material, the dialysed solution is centrifuged for 15 min at 30,000 g and 4C. The supernatant is recovered, filtered through a 0.22-mm filter and kept on ice while connecting the HiLoad SP Sepharose FF column to the FPLC system. The flow rate is set at 3 mL/min and the volume of collected fractions at 7 mL. The column is rinsed with two volumes of water and equilibrated with two volumes of buffer A, two volumes of buffer B, and three volumes of buffer A (see Note 2). The sample is loaded onto the column using the 150-mL superloop and proteins are eluted with a linear gradient of 0–60% of buffer B over a 70-min period. Analysis of the fraction contents by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) is performed by loading 8 mL of each fraction on a 10% gel (one can use a precast gel system from Biorad or any other supplier). The fractions that are richest in Rho protein (Fig. 10.2b) are pooled and stored at 4C until the next day during which the second purification step should be performed. 2. The HiTrap Heparin HP column is connected to the FPLC (flow rate: 2 mL/min; fraction volume: 4 mL), rinsed and equilibrated as described above. The sample is diluted with three volumes of buffer A, filtered through a 0.22 mm filter, and loaded onto the column. The proteins are eluted with a linear gradient of 0–100% of buffer B over a 40-min period. The fractions are analyzed by SDS-PAGE (see above). The fractions richest in Rho and poorest in contaminants (Fig. 10.2c) are pooled, dialyzed twice against 1 L of storage buffer for 4 h at 4C, and filtered through a 0.22-mm filter. The absorbance at 280 nm is measured to determine Rho concentration using an extinction coefficient e280 = 91,600 (mol/L)–1 cm–1 for the hexamer. One volume of 99% glycerol is added to the Rho solution which is stored in 2 mL aliquots at –20C (see Note 7). We routinely verify the concentration of Rho hexamers (MM: 283,200 g/mol) in the glycerol stocks using a colorimetric assay (Quick Start Bradford protein assay kit; Biorad) and bovine serum albumin as a standard. Deviation from the concentration deduced from the A280 measurements never exceeds 15%.

3.2. Preparation of Nucleic Acid Substrates for Helicase and Streptavidin Displacement Assays

The minimal substrates for testing Rho’s motor activity contain long RNA transcripts (> 100 nts) bearing an upstream 47-nt-long aRut motif (Fig. 10.1b). The substrates also contain a DNA oligonucleotide annealed to the downstream portion of the RNA (helicase assay) or a streptavidin tetramer bound to a biotin that has been linked chemically to the 30 end of the RNA chain (streptavidin displacement assay). To investigate specific aspects of the Rho mechanism, the NA strands can sometimes be split into several smaller components (15, 16), provided they all assemble

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(through base pairings) into a single specific NA complex sufficiently stable and homogeneous for purification by native PAGE. Yet, in most cases, the RNA components of the substrates exceed the size limit of commercial oligoribonucleotides (80 nts) and need to be prepared by splint-directed ligation (17) or, as described below, by in vitro transcription. 3.2.1. RNA Production and Radio-Labeling with 32P

1. Transcription components are gently thawed on ice, homogenized with a vortex, and briefly centrifuged in a microcentrifuge. A mixture of 134 mL of water, 50 mL of transcription buffer, 12.5 mL of each 100 mM rNTP stock, 1 mL of Superasin, and 5 mL of DNA template (see Note 8) solution is assembled on ice. Then, 10 mL of T7 RNA polymerase are added and the mixture is incubated for 2 h at 37C. The DNA template is then digested through the addition of 5 mL of RQ1 DNase to the mixture followed by incubation for 20 min at 37C. The mixture is mixed with 12 mL of EDTA (0.5 M), 28 mL of NaAc (3 M) and 900 mL of ethanol and incubated overnight at –20C. The sample is centrifuged for 30 min at 10,000 g in a refrigerated benchtop centrifuge. The pellet is separated from the supernatant, dried in a Speedvac apparatus, and dissolved in a mix of 10 mL of M10E1 buffer and 20 mL of denaturing loading buffer. The sample is left at room temperature while preparing the denaturing polyacrylamide gel for purification. We use custom-made 20 20 cm gel plates equipped with 0.8 mm spacers, a 15-teeth comb, and a bottom tape seal (similar gel sets are available from Fisher scientific or VWR). The denaturing gel solution is mixed with 90 mL of APS and 45 mL of TEMED, quickly poured between the gel plates, and the comb is inserted. Once the gel has polymerized (it takes 15 min), the comb is removed and the wells washed with 1 TBE using a 5-mL syringe. The gel is then installed into an electrophoresis unit and the top and bottom tanks are filled with 1 TBE. After a preelectrophoresis of 20 min at 20 W, the power is turned off and the RNA sample, which has been heat-denatured for 2 min at 95C, is distributed into two wells using a flat gel loading tip (right before loading, flush diffusing urea from the wells using a syringe containing 1 TBE). Typically (i.e., for 100–130 nt transcripts), the gel is run at 20 W until the band corresponding to xylene cyanol is 5 cm from the bottom of the gel. Then, the gel is carefully separated from the plates, wrapped in saran sheets, and placed on an X-ray intensifying screen (or a fluor-coated TLC plate). The band corresponding to the transcript is visualized by UV shadowing in a dark room with a hand-held 254 nm lamp (see Note 9). The band is cut with a scalpel, crushed by passage through a 1-mL syringe, and soaked with 3 mL of elution buffer in a sterile 14-mL culture tube. The tube is shacked overnight at 4C. Then, the gel slurry is passed through

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a 5-mL syringe equipped with a glass wool or cotton plug (to retain most gel particles) and a 0.45-mm filter (e.g., Luerlocked Millex filter from Millipore). The volume of the resulting solution is measured before being mixed with three volumes of ethanol and incubated overnight at –20C. After centrifugation for 30 min at 8000 g in a refrigerated benchtop centrifuge, the supernatant is discarded and the RNA pellet is washed carefully with 300 mL of 70% ethanol, dried in a Speedvac apparatus, and dissolved in 50–100 mL of M10E1 buffer before being stored at –20C. The RNA concentration is determined from the absorbance of the solution at 260 nm, assuming e260 [104 number of nucleotides] L/mol/cm (see Note 10). Typical yields range between 1 and 4 nmoles of purified transcript for a 250 mL transcription. 2. To remove the 50 triphosphate groups, transcripts (10 pmoles) are mixed in an eppendorf tube with M10E1 buffer (50 mL, final volume) and three units of alkaline phosphatase. The mixture is incubated for 30 min at 52C before addition of 5 mL of NaAc (3 M) and 50 mL of phenol extraction solution. The mixture is vortexed vigorously and then centrifuged for 1 min at 10,000 g. The top (aqueous) phase is transferred to another eppendorf tube and, following the same protocol, is extracted once more with one volume of phenol solution and thrice with one volume of saturated chloroform. The aqueous phase is then mixed with three volumes of ethanol and incubated for 2 h at –20C to precipitate the RNA transcripts. After centrifugation for 20 min at 10,000 g in a refrigerated benchtop centrifuge, the supernatant is discarded and the RNA pellet is dried in a Speedvac apparatus before being dissolved in 4 mL of M10E1 buffer. Then, 4 mL of 32 P-gATP (see Note 5), 1 mL of kinase reaction buffer, and 1 mL of polynucleotide kinase are added to the mixture which is incubated for 30 min at 37C. The mixture is then loaded onto a G-50 quick spin column to remove free 32P-gATP. After centrifugation of the column for 4 min at 1100 g in a swinging-bucket rotor, the eluate is mixed with 0.1 volume of NaAc (3 M) and is extracted with phenol as described above. The aqueous phase is mixed with 3 volumes of ethanol and incubated for 2 h at –20C. After centrifugation for 20 min at 10,000 g and removal of the supernatant, the pellet contains the 32P-labeled transcripts (see Note 11) that will be used either for the preparation of RNA–DNA hybrids or in biotinylation reactions. 3. To prepare RNA–DNA hybrids for helicase reactions, the pellet of 32P-labeled transcripts is dissolved in 20 mL of annealing buffer containing 20 pmoles of complementary DNA oligonucleotide. The mixture is heated for 2 min at

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95C and then cooled to 20C over a 30-min period (this is best done in a thermocycler) before being mixed with 5 mL of native loading buffer, and loaded in a single well of a 9% native polyacrylamide gel. (The gel is prepared and run as described in Section 3.2.1.1 with the following exceptions: (1) the native acrylamide solution is used; (2) the power supply is controlled by voltage set at 150 V; and (3) the electrophoresis is stopped after 4 h of migration.) After electrophoresis, the gel is separated from the plates and wrapped in saran sheets. Phosphorescent markers (e.g., Identi-kit from Sigma-Aldrich) are taped to the gel and briefly illuminated with a neon lamp. In a dark room and under dim red light, an autoradiography film is placed into an exposure cassette, exposed to the gel for 10 min, and processed into the developer and fixer bathes. The autoradiographic prints of the phosphorescent markers are then used to position the gel on top of the film and to locate the position of the band corresponding to the 32P-labeled RNA–DNA hybrid (Fig. 10.3a). The band is

Fig. 10.3. Composite substrates for the in vitro assays of Rho’s motor activity. (a) A representative autoradiography of a preparative gel for the purification of 32P-labeled RNA–DNA substrates (helicase assay). (b) The nature of the biotinylated RNA-streptavidin complexes (i.e., number of biotinylated transcripts bound per streptavidin tetramer) depends on the concentration of streptavidin in the initial reaction mixture and elution solution. The gel shows the distribution of complexes as a function of the amount of streptavidin present in the reaction mixture. (c) Representative gels and graphs showing the ATP-dependent RNA–DNA helicase and streptavidin displacement activities of Rho (multi-run conditions). Note that the RNA components of the substrates are different in the two reactions and that all helicase aliquots have been loaded at once on the gel (see Note 14).

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excised from the gel, crushed by passage through a 1-mL syringe, and soaked with 600 mL of elution buffer into an eppendorf tube. The gel slurry is shacked overnight at 4C and then passed through a 5-mL syringe equipped with a small wad of glass wool and a 0.22-mm filter. The filtered solution is mixed with three volumes of ethanol and incubated for 2 h at –20C. After centrifugation for 30 min at 8000 g, the supernatant is discarded and the RNA pellet is washed carefully with 300 mL of 70% ethanol, dried in a Speedvac apparatus, dissolved in 50 mL of 1 annealing buffer (see Note 11) and stored at –20C. 4. To introduce biotin moieties at the 30 ends of transcripts, the pellet of 32P-labeled transcripts is dissolved in 22 mL of water and 33 mL of oxidation buffer before addition of 50 mL of 0.2 M sodium periodate. The mixture is then incubated in the dark for 90 min at 25C before the addition of 10 mL of 3 M KCl and further incubation in the dark for 60 min on ice (excess periodate will precipitate as KIO4). The sample is then centrifuged for 20 min at 10,000 g in a refrigerated benchtop centrifuge, the supernatant is collected, mixed with 0.1 volume of NaAc (3 M) and three volumes of ethanol, and incubated for 2 h at –20C. After centrifugation for 20 min at 10,000 g, the supernatant is discarded and the RNA pellet is washed carefully with 300 mL of 70% ethanol, dried in a Speedvac apparatus, and dissolved in 10 mL of water. Then, 33 mL of oxidation buffer and 20 mL of biotin hydrazide solution are added before incubation for 4 h at 25C. Unreacted hydrazide is eliminated by filtering the solution through a G-50 quick spin column. The eluate is mixed with 0.1 volume of NaAc (3 M) and three volumes of ethanol, and incubated for 2 h at –20C. After centrifugation for 20 min at 10,000 g, the RNA pellet is separated from the supernatant, dried in a Speedvac apparatus, and dissolved in 12.5 mL of water. Then, 0.5 mL of Superasin, 6 mL of native loading buffer, 1.5 mL of KCl (3 M), 2 mL of BSA, and 7.5 mL of streptavidin are added to the mixture before incubation for 20 min at 30C. The sample is distributed into two wells of a 7.5% polyacrylamide gel containing 0.3% SDS (prepared with the TBES buffer and helicase gel solution as described in Section 3.2.1.1), which is run for 3 h at 130 V. The band corresponding to the biotinylated RNA-Streptavidin complex is located by autoradiography as explained above (see also Fig. 10.3a), excised from the gel, crushed by passage through a 1-mL syringe, and soaked with 600 mL of elution buffer supplemented with 200 nM streptavidin (see Note 12). The gel slurry is shacked for 1 h at 20C and then passed through a 5-mL syringe equipped with a glass wool plug and a 5-mm filter. The filtered solution is mixed with three volumes of

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ethanol and incubated for 2 h at –20C. After centrifugation for 30 min at 8000 g, the supernatant is discarded and the RNA pellet is washed carefully with 300 mL of 70% ethanol, dried in a Speedvac apparatus, dissolved in 20 mL of 1 annealing buffer (see Note 11), and stored at –20C. 3.3. Helicase and Streptavidin Displacement Assays

1. A 7.5% polyacrylamide gel containing 0.3% SDS (see Section 3.2.1.4) is prepared and installed in advance into an electrophoretic unit (set the power at 130 V and perform a pre-run for at least 15 min). To collect reaction aliquots, eppendorf tubes (one tube per time point) containing 4.5 mL of quench buffer are also prepared in advance and stored at room temperature. The helicase (or streptavidin displacement) reaction mixture is then prepared by mixing 0.15 pmole of RNA– DNA hybrid (or biotinylated RNA-streptavidin complex) and 0.6 pmole of Rho hexamers in 27 mL of 1 helicase buffer (see Note 13) and incubated for 3 min at 30C. The helicase (or streptavidin displacement) reaction is initiated by addition of 3 mL of initiation mix before further incubation at 30C (see Note 14). At defined incubation times, reaction aliquots (1.5 mL) are withdrawn, mixed with quench buffer in the preset tubes, and immediately loaded on the running gel (see Note 15). After 3 h of electrophoresis at 130 V, one of the gel plates is removed and replaced by a sheet of Whatman paper. The second plate is replaced by a piece of saran wrap and the gel is dried in a vacuum-gel dryer (e.g., HydroTech gel drying system from Biorad) for 30 min at 80C. The gel is then placed into an exposure cassette and exposed overnight to a phosphorimager screen for subsequent detection and analysis (Fig. 10.3c) with a phosphorimager system and related software (e.g., Storm-860 imager and ImageQuant software from GE-Healthcare).

4. Notes 1. This salt is highly hygroscopic. Make sure to use a powder batch than has been stored in a dry cabinet. Whenever necessary, the concentration of solutions made from hygroscopic salts can also be verified using density tables such as the ones found in the Handbook of Chemistry and Physics (CRC Press). 2. Solutions and buffers for FPLC should be filtered and degassed by vacuum filtration on a glass filter holder (Millipore) equipped with a 0.22-mm filter. They are best prepared and cooled to 4C in advance.

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3. Use only RNA-grade water to prepare buffers and solutions. We routinely obtain RNA-grade water by filtering ultrapure MilliQ (Millipore) water with 0.2-mM bottle-top sterile filter units (Nalgene). We usually avoid DEPC-treated water because contaminants, such as rust particles (a source of metal ions harmful for RNA), are often introduced during autoclaving (which is necessary to remove excess DEPC). 4. To eliminate bacteria, the major source of RNase contamination, solutions and buffers for RNA preparation, storage, and assays should be prepared in small amounts (<50 mL) and sterilized with a 0.22-mm filter unit. 5. Manipulation of 32P-containing materials should be performed exclusively by individuals who have received proper training and permission from a radiation safety officer. 6. In the absence of RNA and ATP, the oligomeric state of Rho can depend significantly on parameters such as temperature, protein concentration, nature, and concentration of salts (19). We have observed that the Rho protein is best stored in storage buffer at concentrations larger than 0.5 mM and at –20C. When required, dilutions of active Rho can be prepared ex tempo with 0.5 storage buffer and kept on ice for a few minutes. 7. It is important to assess the enzymatic activity of every fresh Rho preparation and to verify that it does not decay upon storage at –20C for extended period of times. The simplest test for Rho activity is the determination of the steady-state ATPase rate, kATPase, in the presence of an excess of poly[rC]. This is easily done with a thin layer chromatography or colorimetric assay (detailed procedures can be found in ref. (18)) or by using a commercial phosphate assay kit (e.g., EnzChek kit from Molecular Probes). In our standard conditions (50 mM KCl, 40 mM HEPES, pH 8.0, 1 mM ATP, 1 mM MgCl2, 25 nM Rho, 0.1 mg/mL poly[rC], 37C), kATPase ranges between 65 and 85 ATP hydrolyzed/(hexamers) for wildtype Rho. We found that kATPase is not much affected by changes of the buffer composition provided that the substrate is present in the form of long (>300 nts) poly[rC] fragments and in large excess (>103 cytosine residues per Rho hexamer). 8. To improve the homogeneity of the transcript 30 ends, one can use DNA templates that have been amplified by PCR with a downstream primer containing 20 -O-methyl residues at its 50 end (20). 9. If transcription has been successful, the band corresponding to the full-length transcript should be clearly seen as one strong UV-shadow. Bands of lesser intensity and faster migration, corresponding to RNA degradation and/or abortive

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transcription products, are also sometimes visible. If these side-products are present in large amounts (> 50% total products), one may consider repeating the transcription/ purification procedure with fresh solutions and new batches of reactants. 10. If using a classical quartz cuvette, clean it thoroughly with water and ethanol to avoid introducing RNases or other contaminants in the RNA solution. We usually prefer to lose a few microliters of solution in measurements with a mLspectrophotometer (e.g., Nanodrop ND-1000). 11. Assuming that the 32P-labelling of RNA transcripts is quantitative, the concentrations of 32P-labeled species can be estimated by a standard calibration procedure in which aliquots of the corresponding solutions and of dilutions of 32P-gATP (of known concentrations) are spotted on a TLC plate. The plates are then exposed to a phosphorimager screen for the quantification of the radioactivity in each spot. The concentrations of 32P-labeled species are deduced by comparing their phosphorimager signal to the ones of the 32P-gATP dilutions (standard calibration curve). 12. The presence of streptavidin in the elution buffer limits the rearrangement of the complexes into unwanted species, notably ones containing several biotinylated RNA molecules per streptavidin tetramer (Fig. 10.3b). 13. Although unnecessary in most cases, an inhibitor of RNases such as Superasin (5–10 units) may be added to the mixture without affecting the reaction outcome (14). 14. Mechanistic information is usually more easily extracted from single-run experiments. An excess of poly[C] oligomers can be added to the initiation mix to trap unbound Rho and establish a single-run regimen. However, poly[C] oligomers can also have peculiar inhibitory or exhilarating effects on Rho function (14, 16, 17). This may be due to the composite nature of the Rho-transcript interaction (Fig. 10.1a) and the theoretically possible binding of several RNA chains per Rho hexamer which may be favored in some specific conditions (at low ionic strength, for instance). We thus caution against any definitive conclusion that would be drawn from single-run experiments alone (14). 15. For helicase reactions, the aliquots can also be stored on ice and loaded only at the end of the experiment. Note that a small precipitate of potassium dodecyl sulfate may appear in the reaction aliquots. We usually do not eliminate this precipitate before loading the samples on the gel as it does not affect the electrophoresis outcome.

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Acknowledgments We apologize to our colleagues, especially past and present members of the Platt, Richardson, Roberts, and von Hippel labs, whose fantastic work on Rho cannot be exhaustively cited here. We gratefully acknowledge J.M. Berger, J.P. Richardson, and P.H. von Hippel for the gift of materials. Special thanks to A.R. Rahmouni, head of the ‘RNP interactions and therapeutic applications’ lab at Centre de Biophysique Mole´culaire, for his constant support and stimulating discussions. This work was supported by grants from the Agence Nationale de la Recherche (PCV06135253), the Conseil Re´gional du Centre (CPR 07.09.24), the Association pour la Recherche sur le Cancer (grant #3639), and the Ligue contre le Cancer (re´gion Centre). References 1. Roberts J. W. (1969) Termination factor for RNA synthesis. Nature 224, 1168–1174. 2. Ciampi M. S. (2006) Rho-dependent terminators and transcription termination. Microbiology 152, 2515–2528. 3. Richardson J. P. (2002) Rho-dependent termination and ATPases in transcript termination. Biochim. Biophys. Acta. 1577, 251–2560. 4. Skordalakes E. and Berger J. M. (2003) Structure of the rho transcription terminator. Mechanism of mRNA recognition and helicase loading. Cell 114, 135–146. 5. Skordalakes E. and Berger J. M. (2006) Structural insights into RNA-dependent ring closure and ATPase activation by the Rho termination factor. Cell 127, 553–564. 6. Lowery-Goldhammer C. and Richardson J. P. (1974) An RNA-dependent nucleoside triphosphate phosphohydrolase (ATPase) associated with rho termination factor. Proc. Natl. Acad. Sci. USA 71, 2003–2007. 7. Brennan C. A., Dombroski A. J., and Platt T. (1987) Transcription termination factor rho is an RNA-DNA helicase. Cell 48, 945–952. 8. Guerin M., Robichon N., Geiselmann J., and Rahmouni A. R. (1998) A simple polypyrimidine repeat acts as an artificial Rhodependent terminator in vivo and in vitro. Nucleic Acids Res. 26, 4895–4900. 9. Richardson J. P. (1982) Activation of rho protein ATPase requires simultaneous interaction at two kinds of nucleic acid-binding sites. J. Biol. Chem. 257, 5760–5766.

10. Zalatan F., Galloway-Salvo J., and Platt T. (1993) Deletion analysis of the Escherichia coli rho-dependent transcription terminator trp t0 . J. Biol. Chem. 268, 17051–17056. 11. Chen C. Y. and Richardson J. P. (1987) Sequence elements essential for rho-dependent transcription termination at lambda tR1. J. Biol. Chem. 262, 11292–11299. 12. Bogden C. E., Fass D., Bergman N., Nichols M. D., and Berger J. M. (1999) The structural basis for terminator recognition by the Rho transcription termination factor. Mol. Cell 3, 487–493. 13. Wei R. R., and Richardson J. P. (2001) Identification of an RNA-binding site in the ATP binding domain of Escherichia coli Rho by H2O2/Fe-EDTA cleavage protection studies. J. Biol. Chem. 276, 28380–28387. 14. Walmacq C., Rahmouni A. R., and Boudvillain M. (2004) Influence of substrate composition on the helicase activity of transcription termination factor Rho: reduced processivity of Rho hexamers during unwinding of RNA-DNA hybrid regions. J. Mol. Biol. 342, 403–420. 15. Walmacq C., Rahmouni A. R., and Boudvillain M. (2006) Testing the steric exclusion model for hexameric helicases: substrate features that alter RNA-DNA unwinding by the transcription termination factor Rho. Biochemistry 45, 5885–5895. 16. Schwartz A., Walmacq C., Rahmouni A. R., and Boudvillain M. (2007) Noncanonical

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interactions in the management of RNA structural blocks by the transcription termination rho helicase. Biochemistry 46, 9366–9379. 17. Schwartz A., Margeat E., Rahmouni A. R., and Boudvillain M. (2007) Transcription termination factor Rho can displace streptavidin from biotinylated RNA. J. Biol. Chem. 282, 31469–31476. 18. Nowatzke W., Richardson L., and Richardson J. P. (1996) Purification of transcription termination factor Rho from Escherichia coli

and Micrococcus luteus. Methods Enzymol. 274, 353–363. 19. Geiselmann J., Yager T., Gill S., Calmettes P., and von Hippel P. (1992) Physical properties of the Escherichia coli transcription termination factor rho. 1. Association states and geometry of the rho hexamer. Biochemistry 31, 111–121. 20. Kao C., Rudisser S., and Zheng M. (2001) A simple and efficient method to transcribe RNAs with reduced 30 heterogeneity. Methods 23, 201–205.

Chapter 11 Single-Molecule Studies of RecBCD Thomas T. Perkins and Hung-Wen Li Abstract RecBCD is a processive molecular motor composed of two independent helicase domains and a nuclease domain. Understanding the molecular mechanism of its motor activity involves, in part, determining RecBCD’s translocation properties (e.g., velocity, propensity to pause, pause duration). Single-molecule techniques, in general, and optical trapping, specifically, provide for measuring the translocation of individual molecules along DNA. We developed a high-spatial resolution optical-trapping assay for RecBCD. The RecBCD is anchored to the surface via a genetically engineered biotin. This RecBCD-bio exhibited native activity, as measured by biochemical assays. Motion is continuous down to a detection limit of 2 nm, implying a unitary step size below 6 base pairs. Unexpectedly, the catalytic rate changes abruptly and persists at different values for tens of seconds. This technically demanding, high-resolution optical-trapping assay is complemented by a simpler single-molecule assay—the tethered particle motion assay. Key words: Single molecule, optical trap, optical tweezers, RecBCD, helicase, tethered particle motion.

1. Introduction Escherichia coli RecBCD is one of the fastest helicases known. It can unwind tens of thousands of base pairs (bp) at rates up to 500 bp/s at 23C (1, 2). RecBCD also exhibits nuclease activity. To promote DNA repair, this nuclease activity is modulated by enzymatic recognition of the site (5’-GCTGGTGG-3’). This sequence-induced change in RecBCD alters both its velocity (3) and its interaction with RecA, a strand-exchange protein (4). To study the complex properties of RecBCD, we developed a highresolution assay to measure the effects of mechanical forces that may act on molecules during function in vivo (5). M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_11, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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In the last decade, single-molecule techniques have revolutionized biophysics (6–8). The study of individual DNA-based molecular motors by both optical trapping (9–13) and fluorescence techniques (14, 15) is flourishing. One strength of optical traps is that they allow researchers to go beyond watching molecules by exerting precisely controlled loads deduce the effects of force on enzymatic activity, including catalytic rate (9) and force-induced pausing (5). Pioneering single-molecule RecBCD experiments (1, 16) were conducted in the absence of mechanical forces. Our optical-trapping assay for RecBCD allows for controlled application of force and increased spatial precision (5). To increase the reproducibility of the in vitro assay, we use RecBCD-bio, a fully functional derivative that is biotinylated on a carboxy-terminal extension of the RecD subunit (16). RecBCD-bio, bound to a blunt end of double-stranded (ds) DNA, is attached to a streptavidin-coated cover slip via its biotin moiety (Fig. 11.1a). A micron-sized polystyrene bead, attached to the opposite end of this DNA, is ‘‘tethered’’ to the surface by the DNA molecule (Fig. 11.1b). The bead is captured and held under tension with an optical trap, and its position is monitored by a second, detection laser. In the presence of ATP, the enzyme unwinds its DNA substrate, leading to an increase in the tension within the DNA. Our instrument incorporates a stage-based force clamp to maintain a constant force (5, 17). Such feedback allows for controlled application of force and real-time monitoring of enzyme position along the DNA at sub-nanometer spatial precision. High-resolution measurements with optical traps require attention to numerous optical details that are outside the scope of this chapter. The virtues and limitations of various optical-trapping geometries are well addressed in a pair of seminal reviews (18, 19) as well as in reviews discussing the application of optical trapping (8, 20, 21); these reviews are crucial guides to those entering the field.

Fig. 11.1. Illustration of the experiment. (a) Biotinylated-RecBCD (RecBCD-bio) is coupled to streptavidin that is, in turn, anchored to the surface via biotinylated-BSA (bio-BSA). Bio-BSA is passively absorbed onto clean glass at random orientations where biotin is depicted as a dark sphere. DNA bound to the enzyme is held taut by an optical trap. (b) Optical-trapping assay (not to scale).

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2. Materials 2.1. Overexpression and Purification of RecBCD-Bio

1. Minimal plates: 2YT medim containing 0.1 mM biotin. 2. 10 mg/mL lysozyme is dissolved in 10 mM Tris–HCl, pH 7.5 (see Note 1). 3. Dialysis tubing with MWCO 12-14000 (Spectra/Por). 4. RecBCD Reaction buffer: 25 mM Tris–HCl, pH 7.5, 1 mM magnesium acetate [Mg(OAc)2], 1 mM dithiothreitol (DTT). 5. RecBCD Storage buffer: 20 mM Tris–HCl, pH 7.5, 0.1 mM EDTA, pH 7.5, 0.1 mM DTT. 6. Monomeric avidin column (Pierce). 7. Buffer A: 50 mM Tris–HCl, pH 7.5, 0.1 mM PMSF, 10% (w/v) sucrose. 8. Centrifugal concentrator: 50,000 NMWL, 0.5 mL maximum volume (Millipore). 9. Biotin.

2.2. Biochemical Characterization of RecBCD-Bio

1. Native DNA electrophoresis gel: 0.9% (w/v) agarose, 40 mM Tris–acetate, 1 mM EDTA, pH 8. 2. Denaturing DNA gel: 4% (w/v) polyacrylamide, 7 M urea, 45 mM Tris–borate, 1 mM EDTA, pH 8. 3. PhosphorImager (Amersham Biosciences). 4. RecBCD Reaction buffer: 25 mM Tris–HCl, pH 7.5, 1 mM Magnesium acetate [Mg(OAc)2], 1 mM dithiothreitol (DTT). 5. PK: 800 units (u)/mL pyruvate kinase in RecBCD reaction buffer with 50% glycerol, store frozen at 100 . 6. PEP: 50 mM phosphoenolpyruvate (PEP) in RecBCD Reaction buffer with 50% glycerol, stored frozen at 100 . 7. ATP regeneration system: 1 mM phosphoenolpyruvate and 4 u/mL pyruvate kinase. 8. Single-stranded binding (SSB) protein: 59 mM SSB (Promega). 9. ATP: 100 mM ATP, store frozen.

2.3. Coupling Antibodies to Polystyrene Beads

1. MES buffer: 100 mM MES (pH 5) þ 0.05% Tween 20. 2. Phos-Tween buffer: 100 mM sodium phosphate (pH 7.5) þ 0.05% Tween 20. 3. Carboxy/sulfate, 0.5-mm diameter polystyrene beads (Interfacial Dynamics) (see Note 2).

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4. Sulfo-NHS: 100 mM Sulfo-NHS (Thermo Scientific) in Phos-Tween buffer (21 mg/mL). 5. EDC: 1-ethyl-3-[3-dimethylaminopropyl]carbodiimide hydrochloride (Thermo Scientific). 6. b-mercaptoethanol. 7. Anti-digoxigenin from sheep (Roche). 8. Cup-horn sonicator (Sonics Vibracell). 9. 1 M glycine (75 mg glycine dissolved in 1 mL H2O, filtered through a 0.2-mm spin filter). 2.4. Cleaning Cover Slips for SingleMolecule Assays

1. KOH pellets. 2. Ethyl alcohol, Chemicals).

completely

denatured

(Mallinckrodt

3. Bath sonicator. 4. Microscope cover slips [22 40 mm2, (thickness ¼ #1.5 for trapping)]. 5. Squirt bottles with H2O and denatured ethyl alcohol. 6. Custom-made cover slip holder machined from Teflon (see Fig. 11.2a).

Fig. 11.2. (a) Teflon holder for cleaning cover slips. (b) Epoxy-stabilized flow cell. Coin is for scale and tape is used to label samples, useful if varying reaction conditions such as incubation time with bead-DNA complexes.

2.5. Making Bead-DNA Complexes

1. Dig-DNA: DNA made via PCR using a single 50 -digoxegininlabeled primer, typically DNA length is 3–10 kbp. Use at a final concentration of 100 pM. 2. AD beads: Anti-digoxigenin (AD) beads from Section 3.3. Use at a final concentration of 500 pM. 3. Phos-Tween buffer: 100 mM sodium phosphate (pH 7.5) þ 0.05% (v/v) Tween 20. 4. WB buffer: 25 mM Tris–acetate (pH 7.5), 1 mM Mg(OAc)2, 1 mM NaCl, 1 mM DTT, 0.4% (v/v) Tween 20, and 3 mg/ mL bovine serum albumin (BSA). 5. Cup-horn sonicator (Sonics Vibracell).

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1. Cleaned cover slips as described in Section 3.4. 2. Double-stick tape. 3. Five-minute epoxy. 4. WB buffer: 25 mM Tris–acetate (pH 7.5), 1 mM Mg (OAc)2, 1 mM NaCl, 1 mM DTT, 0.4% Tween 20, and 3 mg/mL BSA. 5. Phosphate buffer: 100 mM sodium phosphate (pH 7.5). 6. Bio-BSA (Vector labs): biotinylated BSA (10 mg/mL in phosphate buffer) (see Note 3). 7. SA: streptavidin (5 mg/mL) in WB. 8. Bead-DNA complexes as described in Section 3.5. 9. RecBCD-bio: biotinylated RecBCD as described in Section 2.1, dilute to 15 nM in WB.

2.7. Single-Molecule Optical-Trapping Assay

1. Flow cells with RecBCD as described in Section 3.6. 2. Single-stranded binding (SSB) protein: 59 mM SSB (Promega). 3. Catalase: 1.3 103 u/mL. Stock concentration is 1.3 106 u/mL made by suspending 250 mg catalase (6500 u/ mg) in 12.5 mL H2O. Aliquot at 1:1000 in H2O and store at 4C. 4. Glucose oxidase (GOD): 25 mM GOD. 20 mg GOD (Roche) þ 5 mL H2O. Aliquot into 5 1 mL aliquots and store at 4C. 5. Glucose: 100 mg/mL in H2O, store frozen. 6. ATP: 100 mM ATP, store frozen. 7. WB buffer: 25 mM Tris–acetate (pH 7.5), 1 mM Mg(OAc)2, 1 mM NaCl, 1 mM dithiothreitol (DTT), 0.4% (v/v) Tween 20, and 3 mg/mL BSA (see Note 4). 8. RXN buffer: 5 mM ATP, 1.1 mM SSB, 30 u/mL catalase, 1.2 mM GOD, 6 mg/mL glucose in WB. Made by mixing 79.8 mL WB, 5 mL ATP, 1.9 mL SSB, 2.3 mL catalase, 5 mL GOD, 6 mL glucose. Use immediately. 9. High-resolution optical-trapping microscope.

2.8. Video-Based Tethered Particle Assay

1. Cleaned cover slips as described in Section 3.4. 2. Microscope slides. 3. Double-stick tape. 4. Five-minute epoxy. 5. Anti-digoxigenin: 20 mg/mL in 100 mM phosphate buffer, pH 7.5. 6. TPM RXN buffer: 25 mM Tris–HCl, pH 7.5, 1 mM Mg(OAc)2, 1 mM DTT, and 10 mg/mL BSA.

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7. Dig-DNA: 1 nM in TPM RXN buffer. DNA made via PCR using a single 5’-digoxeginin-labeled primer, typically DNA length is 1.5 kbp. 8. RecBCD-bio: biotinylated RecBCD as described in Section 2.1; dilute to 0.08 nM in TPM RNX buffer. 9. Streptavidin beads: 0.2-mm diameter streptavidin beads (Bang laboratory) at 0.1 nM in TPM RXN buffer. 10. Single-stranded binding (SSB) protein: 59 mM SSB (Promega). 11. Glucose: 100 mg/mL in H2O, store frozen. 12. ATP: 100 mM ATP, store frozen. 13. PK: 800 u/mL pyruvate kinase in RecBCD reaction buffer with 50% glycerol, store frozen at 100 . 14. PEP: 50 mM phosphoenolpyruvate in RecBCD reaction buffer with 50% glycerol, store frozen at 100 . 15. Video-enhanced differential interference contrast (DIC) microscope with high-sensitivity camera (Newvicon).

3. Methods Successful single-molecule, optical-trapping experiments require both the instrumentation and the single-molecule assay to work at the same time. The assay, called a motility assay for molecular motors, is often the most difficult and variable part of singlemolecule biophysics. For instance, there are many ways to anchor proteins to a surface. The challenge is to maintain the biochemical activity. The ‘‘gold’’ standard in single-molecule experiments is that the average of the single-molecule rates should agree with biochemical rates measured by traditional assays. As the single-molecule-biophysics community is becoming more sophisticated, researchers are adopting genetically cloneable handles for anchoring enzymes to the surface (22). We use biotin carboxyl carrier protein (BCCP)-tagged RecBCD (‘‘RecBCDbio’’) (16). This specific anchoring increases the fraction of active enzymes over the simpler passive absorption of proteins onto glass used in the pioneering optical-trapping studies of a DNA-based molecular motor (9, 23). 3.1. Overexpression and Purification of RecBCD-Bio

1. Streak out E. coli strain encoding the biotin carboxyl carrier protein (BCCP)-tagged RecBCD (‘‘RecBCD-bio’’) onto minimal plates (ampr/camr) and grow for 48 h at 37C. 2. Inoculate 10 mL of 2YT media containing 100 mg/mL ampicillin and 10 mg chloramphenicol with one colony from the plate and grow overnight in 37C shaker.

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3. Inoculate 1 L of 2YT in a 4-L erlenmeyer flask with the 10mL overnight culture (1:100 dilution) and grow in 37C shaker until the absorbance at 650 nm ¼ 0.4 (4.5 h). 4. Once absorbance reaches 0.4, induce with 1190 mL of 0.84 M IPTG (1 mM final conc.) and replenish the culture with ampicillin to 100 mg/mL. 5. Allow the culture to grow for an additional 4 h. 6. Harvest cells by spinning down at 6250 g for 15 min at 4C. 7. Remove supernatant and wash the pellets in 10 mL buffer A. 8. Pool the suspensions and spin again at 6250 g for 15 min at 4C. 9. Add the 10 mg/mL lysozyme while stirring to a final concentration of 0.2 mg/mM (1:50). 10. Incubate 5 min at 4C. 11. Add 4 M NaCl to a final concentration of 0.1 M (1:40), and 0.4 M spermidine to a final concentration of 0.01 M (1:40). Incubate 25 min at 4C. 12. Warm suspension to 20C in a 37C water bath. 13. Chill rapidly to 4C. 14. Spin the lysate at 14,500 g at 4C for 20 min. Collect the supernatant. 15. Spin this supernatant at 136,500 g at 4C for 30 min. Collect the supernatant. 16. Dialyze against RecBCD Reaction buffer to remove excess biotin. Repeat four times. 17. Carefully remove the post-dialysis sample from tubing and spin post-dialysis sample at 14,500 g at 4C for 20 min. Collect the supernatant. 18. Load this supernatant onto a pre-blocked 5 mL monomeric avidin column that has been pre-equilibrated with RecBCD Storage buffer. Collect the flow through at the rate of 100 mL/min in one tube. 19. Stop the flow completely for 15 min to allow RecBCD-bio to bind. 20. Rinse with 30 mL RecBCD Storage buffer at the rate of 100 mL/min and collect this flow through in a second tube. 21. Elute protein with 10 mL of 0.2-mm-filtered 5 mM biotin in RecBCD Storage buffer. Collect 1 mL fractions at the flow rate of 500 mL/min. 22. Run a 10.5% (w/v) SDS-PAGE of the collected flow-through and fractions to determine the protein peak.

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23. Pool the appropriate fractions and rapidly dialyze out the excess biotin to prevent interference in the single-molecule assay. This requires 6–8 buffer changes depending on volume of fractions pooled. 24. Concentrate the RecBCD-bio with centrifugal concentrator. 25. Aliquot 5-mL of purified proteins into tubes and store in –80C. 26. Determine the protein concentration by Bradford Assay and by absorbance at 280 nm (e = 40,000 M–1 cm–1). 3.2. Biochemical Characterization of RecBCD-Bio

1. A 2.6-kbp DNA lacking -sequences is prepared by PCR including [a-32P]-dATP. 2. The RecBCD-unwinding reaction mixture (final volume ¼ 40 mL) is made by mixing the constituents with RecBCD Reaction buffer to achieve final concentrations of 1.4 nM dsDNA ends, 0.14 nM RecBCD-bio, 1 mM SSB, and ATP regeneration system in RecBCD Reaction buffer. Let it sit for 5 min at room temperature. 3. The reaction is initiated by adding ATP to a final concentration of 15 mM ATP. 4. The reaction mixtures are quenched at the specified times with 10 mL of 0.1 M EDTA, 3% (w/v) sodium dodecyl sulfate, and 50% glycerol. 5. The samples are then extracted with phenol/chloroform/ isoamyl alcohol (25:24:1) and split into two aliquots for native and denaturing DNA gels. 6. One aliquot of 12 mL is loaded on a native electrophoresis gel to analyze the RecBCD-catalyzed unwinding. 7. A second aliquot of 12 mL is mixed 1:1 (v/v) with formamide, denatured for 5 min at 96C, and 24 mL is loaded on a 4% (w/v) polyacrylamide-denaturing gel to analyze the nuclease activity. 8. Radioactivity is measured using a PhosphorImager. 9. Concentrations are calculated by integrating the specified areas of the gel lanes and normalized to the full-length DNA band at zero time. 10. Helicase activity was calculated by the decrease of full-length duplex DNA in the time-coursed native DNA gel, and the nuclease activity was determined by the decrease of full-length ssDNA in the denaturing DNA gel.

3.3. Coupling Antibodies to Polystyrene Beads

1. Add 165 mL stock carboxy/sulfate 0.5-mm diameter beads (3.4% solids) to 150 mL MES buffer. Vortex. 2. Mix and wash twice by pelleting in a microcentrifuge at 18,000 g for 3 min. Remove supernatant. Resuspend in 200 mL MES buffer (see Note 5).

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3. Add 600 more mL MES buffer and 100 mL of Sulpho-NHS (21 mg/mL). Briefly vortex. 4. Add 5 mg EDC. Briefly vortex. Put on rotator for 30–45 min at room temperature. 5. Add 1000 mL Phos-Tween buffer to manufacturer’s vial containing 200 mg of anti-digoxigenin antibody. Mix gently by occasionally rotating vial by hand at room temperature over 20 min (see Note 6). 6. Add 1 mL b-mercaptoethanol to bead solution. Briefly vortex. Let it rotate for 10 min. 7. Pellet beads for 5 min at 18,000 g. Remove as much of the supernatant as possible while retaining the beads in the pellet. 8. Resuspend pellet by adding 200 mL of Phos-Tween buffer, being careful not to introduce bubbles. 9. Wash beads twice more by pelleting and resuspending into Phos-Tween buffer. 10. Add 300 mL Phos-Tween buffer to bring the final volume to 500 mL. 11. Add beads to 1000 mL anti-digoxigenin solution, and quickly mix with pipette followed by a gentle vortex (2 s). Place on rotator for > 2 h at room temperature. 12. Quench reaction by adding 30 mL of 1 M glycine. Let it sit for 10 min at room temperature on rotator. 13. Wash beads five times in Phos-Tween buffer to remove unbound antibody, resuspending into 200 mL. First spin is 6 min. 14. Add 800 mL Phos-Tween buffer to bring final volume to 1000 mL. 15. Take 20 mL diluting into 1 mL Phos-Tween buffer. Measure OD at 500 nm and compare to similar dilution of stock beads to determine concentration (see Note 7). 16. Store at 4C (see Note 8). 3.4. Cleaning Cover Slips for SingleMolecule Assay

Single-molecule assays are very sensitive to surface conditions. We always clean cover slips prior to use; otherwise the success of the assay is highly variable (see Note 9). 1. Put 250 mL ethyl alcohol (completely denatured) and 80 g potassium hydroxide pellets into a 1-L beaker. Put in medium stir bar. Cover with parafilm. Place on stir plate. Stir until dissolved (see Note 10). 2. Fill bath sonicator half full with water. 3. Place two 1-L beakers half filled with H2O in sonicator. 4. Load cover slips into cover-slip holder (see Fig. 11.2a).

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5. Place cover slips into the ethanolic KOH that is sitting in a bath sonicator. Cover with parafilm. 6. Sonicate cover slips in ethanolic KOH for 3 min, occasionally agitating the solution by hand. 7. Rinse cover slips with H2O squirt bottle. 8. Sequentially rinse cover slips in the two H2O-filled 1-L beakers for 3 min each, while sonicating. Cover beakers with parafilm. 9. Rinse with H2O using squirt bottle. 10. Rinse with ethyl alcohol (completely denatured). 11. Microwave dry 2 min. 12. Place holder (with cover slips) in container to isolate from ambient dust (see Note 11). 3.5. Making Bead-DNA Complexes

Bead-DNA complexes are made fresh each day. Anti-digoxigenin (AD) beads clump over time. For optical-trapping applications, we start with 7.5 fmol (15 mL 500 pM) beads per slide. We typically use 8–12 slides per day. Bead are made concurrent with Section 3.6. 1. Briefly vortex AD bead stock. 2. Add 7.5 fmol AD beads per slide to a new tube. If volume is less than 200 mL, add Phos-Tween buffer to 200 mL. 3. Centrifuge for 5 min at 9000 g. Remove supernatant. 4. Resuspend beads in 200 mL Phos-Tween buffer, centrifuge 5 min, and remove supernatant. Repeat two more times. 5. Resuspend in 15 mL WB per slide (e.g., 120 mL for eight slides). 6. Seal top of tube by wrapping with parafilm. Sonicate beads for 10–20 min in cup sonicator with either a recirculating bath or a cup pre-chilled with some ice. Remove 1 mL beads, dilute into 100 mL WB, and check these diluted beads under microscope for monodispersity. For each clump of beads there should be 8–10 single beads. If not, repeat sonication (see Note 12). 7. Dilute digoxigenin-DNA (1.5 fmol per slide) into 15 mL WB per slide. 8. Add DNA to bead solution. Mix quickly by pipetting a few times and then a gentle, rapid (1 s) vortex. Incubate for 20 min at room temperature.

3.6. Flow Cell Assembly of RecBCD-DNA-Bead Complex for OpticalTrapping Assay

The goal is to maintain activity of the helicase while minimizing non-specific sticking of the beads and DNA to the surface. BSA is used as a sacrificial protein to bind to any ‘‘sticky’’ spots. In addition, low concentrations of a weak nonionic detergent (e.g., Tween 20) significantly reduce unwanted sticking.

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1. Place slide on clean bench surface. 2. Carefully cut 5 cm of double-stick tape in half lengthwise with razor blade. Place each half perpendicular to the long axis of the microscope slide but centered on the slide. The interior gap between the tape should be 5–8 mm. 3. Place the clean cover slip on the tape so there is an overhang on each side (see Fig. 11.2b). Gently press down on cover slip with Teflon tweezers to seal the tape to the cover slip. 4. Repeat steps 1–3 until the appropriate number of slides is made (typically 8–12 per day). 5. Mix 5-min epoxy, apply to the long edges first, and wait for the epoxy to flow into the gap between the tape and slide. Then apply small dots of epoxy on the short edges. 6. Pipette in 50 mL of bio-BSA (10 mg/mL). Wait 20–60 min at room temperature (see Notes 13 and 14). 7. Wash 200 mL WB through flow cell two times. Wait 10 min. Wash with an additional 200 mL WB (see Note 15). 8. Flow through 50 mL SA (10 mg/mL). Let it sit for 20 min at room temperature (see Note 16). 9. Wash 200 mL WB through flow cell two times. Wait 10 min. Wash with an additional 200 mL WB. 10. Flow through 20 mL RecBCD-bio (15 nM in WB). Wait 2 h at room temperature. 11. Wash 200 mL WB through flow cell two times. Wait 10 min. Wash with an additional 200 mL WB. 12. Flow through 30 mL bead-DNA complexes. Wait 1 h (see Note 17). 13. Wash 200 mL WB through flow cell two times. 3.7. Optical-Trapping Assay

1. Turn on optical-trapping apparatus (Fig. 11.3a). Mount flow cell with 10 fM beads in WB and align for Kohler illumination. Let warm up for > 2 h with trapping laser at a nominal trapping stiffness (ktrap = 0.08 pN/nm) (see Note 18). 2. Turn off trapping laser. Mount flow cell from Section 3.6. 3. Verify through the Bertrand lens (rotate thumbwheel in the eye piece) that there are no bubbles in the immersion oil. Turn on trapping laser. Let it sit for 5 min. 4. Use a coarse translation stage to find a field of view with approximately six tethers (see Note 19). 5. Mix up RXN buffer. Flow it through the flow cell (see Note 20). 6. Turn off trapping laser. Using the 3D PZT stage, approximately position a tethered particle under the stationary trap. Turn trap on, capturing a tethered bead, and then pretension

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Fig. 11.3. (a) Optics diagram for high-resolution optical trapping. A detection laser (DL, l ¼ 785 nm) measures the position of the bead in the trapping laser (TL, l ¼ 1064 nm) using back-focal-plane detection (28, 29). The two lasers are combined with dichroics to couple them into the objective (Obj). The piezo-electric (PZT) mirror is imaged onto the back focal plane of the objective, enabling independent beam steering of the trapping beam relative to the detection beam. As the bead position increases due to enzymatic motion, a software-based feedback loop stabilizes the force by moving the stage (5, 17). Acronyms represent the following: OI (optical isolator), PBS (polarizing beamsplitter), quadrant photodiode (QPD), and l/2 (half-wave plate). (b) Experimental trapping geometry. RecBCD, bound to DNA, is anchored to the surface. The distal end of the DNA is attached to a bead held in an optical trap. Pretensioning of the DNA moves the bead out of the trap. As the experiment proceeds, the stage (xstage) is moved to keep the bead displacement relative to the trap center (xbd) constant. Force is modulated by changing the trap stiffness (not shown). The vertical center of the trap (ztrap) is positioned about (300 nm þ rbd) above the surface to keep the bead from touching the cover slip as it is pulled out of the trap.

the DNA to 1 pN. Measure force as a function of time, where a linearly increasing force indicates an active helicase. If after 20 s, the tension does not increase by at least 0.2 pN, the tether is inactive. Turn off trap and move to a different tethered bead. Repeat until an active tether is found or until the RecBCD’s motion is expected to decrease the DNA length to less than 700 nm. 7. To establish the proper trapping geometry (Fig. 11.3b), determine the vertical position by tapping the trapped bead off the glass (Fig. 11.4a). Set the height to 300–400 nm. Next, determine lateral position of the anchor point by measuring the force-extension curve of DNA (Fig. 11.4b). Repeat both procedures another time to refine the measurement of the anchor point. 8. Tension the DNA to the desired force using a stage-based force clamp (17). Change the set force by changing the trap stiffness (ktrap). Maintain this force by changing the lateral offset between the trap center and the anchor point (xstage). The bead displacement (xbd) is held constant (see Note 21). Record ktrap, xstage, ztrap, and measurements of xbd at 10 kHz, where ztrap is the vertical distance from the trap center to the cover slip [ztrap ¼ height þ rbd] (see Figs. 11.3b and 11.4a).

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Fig. 11.4. Establishing the three-dimensional trapping geometry. (a) The symmetry in the force versus extension curve determines the anchor point relative to the optical trap in each axis. The elasticity data is quantitatively fit by the wormlike chain theory (black line), yielding the persistence length (p = 46 nm), a measure of DNA-bending stiffness. Tethers anchored by multiple DNA are identified by a low persistence length (p < 25 nm) during this measurement and not further measured. (b) To determine the vertical location of the trapped bead over the surface, the sample is moved vertically while monitoring the sum of all the light on the quadrant photodiode (QPD). When the bead touches the cover slip surface (height, h = 0 when ztrap = rbd), there is a discontinuity in the signal, allowing for the vertical trapping geometry to be well controlled to 20 nm.

9. Analyze data based on the two-dimensional trapping geometry (Fig. 11.3b), using the established formulism (24). 3.8. Video-Based Tethered-Particle Assay

To validate RecBCD-unwinding activity at the single-molecule level, we use a video-based tethered particle motion (TPM) assay to monitor the translocation of RecBCD as it unwinds the duplex DNA molecules. Binding, unwinding, and ultimately release can be directly visualized by video-enhanced differential interference contrast (DIC) microscopy (16). The TPM assay is biochemically identical to the optical-trapping assays except that the dsDNA is anchored to the surface of a cover slip, and the RecBCD-bio is bound to commercially available streptavidin-coated polystyrene beads. 1. Flow chambers are assembled as detailed in Section 3.6. 2. Pipette 30 mL of anti-digoxigenin antibody into flow cell. Wait for 1 h at room temperature. 3. Wash 100 mL TPM RXN buffer through flow cell two times. Wait for 30 min. Wash with an additional 100 mL TPM RXN. 4. Flow through 50 mL of 1 nM dig-DNA. Wait 1 h at room temperature. 5. In parallel with step 4 above, incubate 0.08 nM RecBCD-bio with 0.1 nM streptavidin-labeled beads in a total volume of 300 mL on ice for at least 30 min.

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6. Wash flow cell with 100 mL TPM RXN buffer through flow cell two times. Wait for 5 min. Wash with additional 100 mL TPM RXN. 7. Mount slide on microscope. Wait 5 min to settle. 8. Immediately before use, prepare 50 mL of final reaction mixture [0.05 nM RecBCD-bead complex (based on bead concentration), 10 mM ATP, 16 u/mL PK, 1 mM PEP, 1 mM SSB in TPM RXN buffer (all final concentrations)]. 9. Using differential interference contrast (DIC) imaging, record the next 20 min of video (using custom software written in LabView). RecBCD-bead complexes bound to the DNA do not leave the focal plane. Rather, they appear as tethered particles that undergo vigorous Brownian motion within a restricted range (25). As RecBCD unwinds the DNA, the DNA tether length decreases, reducing the range of the Brownian motion (see Notes 22 and 23). 10. Recordings are analyzed using LabView-based program. Based on prior calibrations (26), this analysis returns the time-course of the RecBCD position along duplex DNA and the rate unwinding rate of individual RecBCD helicase.

4. Notes 1. All solutions were prepared with 18.2 M -cm water. 2. For high precision, it is important to use beads with a low 3 coefficient of variation (CV) in size (3%) because ktrap rbd . 3. When binding proteins passively to glass, do not add Tween 20. Tween 20 is added to the solution to suppress nonspecific sticking (as is the high concentration of BSA). Also, high levels of divalent cations (e.g., >3 mM MgCl2) should be avoided, since they also lead to excessive sticking of the DNA to the surface. 4. For trapping applications, buffers are filtered with 0.2-mm filters to remove particles, including clumps of protein, which can fall into the optical trap. 5. Optimum centrifugation time and speed depends on bead size. For example, spin 320-nm-dia. Beads at 13,000 g for 2 min; 400-nm-dia. Beads at 9000 g for 5 min; and 180nm-dia. Beads at 18,000 g for 5 min. Spinning too long can lead to irreversible clumping.

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6. To prevent covalent coupling of two beads by the same protein, it is important to have an excess of protein to completely cover the surface of the bead. You can assume each protein covers 3-nm diameter surface area. With that, you can divide the surface area per bead by this ratio and then multiply by the number of beads per liter to get molarity. 7. To account for loss of beads during repeated washing, it is best to monitor bead concentration by measuring the optical density at 500 nm. Different dilutions from the stock concentration are used to establish calibration curve. 8. The activity of the AD beads can be tested by a DNA-binding assay. Incubate beads and DNA at a molar ratio of 1:1 with a total of 50 ng of DNA. Spin down, collect the supernatant, and run on gel. If the DNA is bound to the beads, it will not be in the supernatant and thus not in the gel. 9. If available, a quick way to clean cover slips is in a plasma cleaner with either pure oxygen or atmosphere. Alternatively, you can use an acid etch [sulfuric acid and hydrogen peroxide (8:1 (v/v), 80C)] for 10 min. 10. KOH solution needs to be clear not cloudy and the approximate color of apple juice. This is very sensitive to the source and the exact denatured ethyl alcohol used. We use catalog #7018. 11. Clean cover slips are useable for 2 weeks if properly stored. Passive absorption, as is used here, is sensitive to surface cleanliness, but not nearly as sensitive as PEG-silanization of cover slips. 12. AD beads tend to clump over time. It is best to sonicate them immediately prior to use. Use a standard optical microscope to assure that the beads are monodisperse (typically > 90%) before adding DNA. Some batches are irreversibly clumpy. If this continues to occur, decrease the EDC so that there is one EDC molecule per ten carboxylate groups on the bead. 13. We always flow fluid through flow cells in one direction. Buffer is dripped in with a micropipette onto one side of the overhanging cover slip and gently aspirated out the other side using a gel loading tip. In the case of a bubble near the input, we add a drop of buffer to the output (‘‘dirty’’) side to try to force the bubble out the input (‘‘clean’’) side. Bubbles that are pulled though the central region of the flow cell will inactive the RecBCD. Such flow cells are discarded. 14. During incubation, flow cells are placed in a humidity chamber to prevent evaporation. These are made from used pipette tip boxes, a plastic pipette (5 mL) broken into two short sections (10 cm) and held onto the pipette tip rack with double-stick tape. Fill the bottom with water. Pipette tips

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inserted into the pipette tip rack holes provide restraint to the flow cells, preventing sliding. Each box holds two flow chambers. 15. The multiple rinses assure quantitative removal of various reactants. While this may seem excessive, the fluid flow is parabolic in the channel, with the boundary condition of fluid velocity equal to zero at the surface. So while the reactions take place at the surface, the vast majority of fluid goes through the center of the channel. The wait periods allow for some diffusion to promote removal of reactants. 16. Proper tether formation is dependent on streptavidin. For properly formed tethers, leaving out the streptavidin leads to a dramatic reduction in tether density (>50-fold). 17. Tether density can be controlled by changing the incubation time with the bead-DNA complex. Typical density is 20–30 tethers per 100 100 mm2. It can also be controlled by increasing the bead-DNA complex concentration. Higher concentrations lead to more stuck beads, which can get in the way of optical-trapping assays, and use beads at an accelerated rate. 18. DNA labeled at opposite ends with digoxigenin and biotin is recommended to test the optical trapping set up, to test the ability to make tethers, and to verify the trap calibrations. Elasticity curves should return a reasonable persistence length [40–50 nm depending on length (27)] for dsDNA and the correct contour length [L ¼ 0.34 nm/bp DNA length (in bp) to – 10 nm]. 19. Typically, 30% of tethers have a RecBCD molecule that moves. So, by having several tethers in the field of view, we increase the probability of finding a moving enzyme before the enzyme translocates too far along the DNA [L < 700 nm (2100 bp)]. 20. Be delicate when flowing buffer through on the microscope. Small mechanical agitations caused by pressing too hard with the pipette tip can lead to significant drifts (> 10 nm) during the measurement. 21. Substantial changes in laser power will lead to significant vertical drift (> 1 nm/min), which can alter experimental geometry. If the height changes significantly, then the bead touches the cover slip and the assumed trapping geometry is not longer valid. This can also happen if using very short tethers (L < 500 nm). 22. On average, there are about 10–20 tethers that appear within the field of view (40 60 mm2). When things are working well, about 30–60% of the tethers unwind. The majority of unwinding tethers detached from the surface at the end of the unwinding.

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23. For the TPM assay, each tether forms at a unique time (e.g., different time stamp), so measurements can be made in parallel on several molecules. The ATP regeneration system (PK/PEP) in the buffer assures that the ATP concentration is uniform over at least 30 min. At longer times, the tethering rate decreases, most likely due to degradation of the surfaceanchored DNA molecules by prior RecBCD molecules.

Acknowledgments The authors would like to thank Ashley Carter for critical reading of the manuscript, and Ashley Carter and D. Hern Paik for figure preparation. This work was supported by a Burroughs Wellcome Fund Career Award in the Biomedical Sciences (TTP), the National Science Foundation (NSF Phys-0404286 to TTP), National Research Council (Taiwan; to H.-W.L.), and National Institute of Standards and Technology (NIST). Mention of commercial products is for information only; it does not imply NIST recommendation or endorsement, nor does it imply that the products mentioned are necessarily the best available for the purpose. TTP is a staff member of NIST’s Quantum Physics Division. References 1. 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. 2. 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. 3. Spies M., Bianco P. R., Dillingham M. S., Handa N., Baskin R. J., and Kowalczykowski S. C. (2003) A molecular throttle: the recombination hotspot chi controls DNA translocation by the RecBCD helicase. Cell 114, 647–654. 4. Kowalczykowski S. C., Dixon D. A., Eggleston A. K., Lauder S. D., and Rehrauer W. M. (1994) Biochemistry of homologous recombination in Escherichia coli. Microbiol. Rev. 58, 401–465. 5. Perkins T. T., Li H. W., Dalal R. V., Gelles J., and Block S. M. (2004) Forward and

6.

7.

8.

9.

10.

11.

reverse motion of single RecBCD molecules on DNA. Biophys. J. 86, 1640–1648. Weiss S. (1999) Fluorescence spectroscopy of single biomolecules. Science 283, 1676–1683. Mehta A. D., Rief M., Spudich J. A., Smith D. A., and Simmons R. M. (1999) Singlemolecule biomechanics with optical methods. Science 283, 1689–1695. Bustamante C., Bryant Z., and Smith S. B. (2003) Ten years of tension: single-molecule DNA mechanics. Nature 421, 423–427. Wang M. D., Schnitzer M. J., Yin H., Landick R., Gelles J., and Block S. M. (1998) Force and velocity measured for single molecules of RNA polymerase. Science 282, 902–907. Abbondanzieri E. A., Greenleaf W. J., Shaevitz J. W., Landick R., and Block S. M. (2005) Direct observation of base-pair stepping by RNA polymerase. Nature 438, 460–465. Wuite G. J., Smith S. B., Young M., Keller D., and Bustamante C. (2000) Single-molecule

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13.

14.

15.

16.

17.

18.

19. 20.

Perkins and Li studies of the effect of template tension on T7 DNA polymerase activity. Nature 404, 103–106. 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. Smith D. E., Tans S. J., Smith S. B., Grimes S., Anderson D. L., and Bustamante C. (2001) The bacteriophage straight phi29 portal motor can package DNA against a large internal force. Nature 413, 748–752. 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. 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. Dohoney K. M. and Gelles J. (2001) Chisequence recognition and DNA translocation by single RecBCD helicase/nuclease molecules. Nature 409, 370–374. Perkins T. T., Dalal R. V., Mitsis P. G., and Block S. M. (2003) Sequence-dependent pausing of single lambda exonuclease molecules. Science 301, 1914–1918. Svoboda K. and Block S. M. (1994) Biological Applications of Optical Forces. Annu. Rev. Biophys. Biomol. Struct. 23, 247–285. Neuman K. C. and Block S. M. (2004) Optical trapping. Rev. Sci. Instrum. 75, 2787–2809. Mehta A. D., Pullen K. A., and Spudich J. A. (1998) Single molecule biochemistry using optical tweezers. FEBS Lett. 430, 23–27.

21. Perkins T. T. (2009) Optical traps for single molecule biophysics: a primer. Laser Photonics Rev., 3, 203–220. 22. Noji H., Yasuda R., Yoshida M., and Kinosita K., Jr. (1997) Direct observation of the rotation of F1-ATPase. Nature 386, 299–302. 23. Yin H., Wang M. D., Svoboda K., Landick R., Block S. M., and Gelles J. (1995) Transcription against an applied force. Science 270, 1653–1657. 24. Wang M. D., Yin H., Landick R., Gelles J., and Block S. M. (1997) Stretching DNA with optical tweezers. Biophys. J. 72, 1335–1346. 25. Schafer D. A., Gelles J., Sheetz M. P., and Landick R. (1991) Transcription by single molecules of RNA polymerase observed by light microscopy. Nature 352, 444–448. 26. Yin H., Landick R., and Gelles J. (1994) Tethered particle motion method for studying transcript elongation by a single RNA polymerase molecule. Biophys. J. 67, 2468–2478. 27. Seol Y., Li J., Nelson P. C., Perkins T. T., and Betterton M. D. (2007) Elasticity of short DNA molecules: theory and experiment for contour lengths of 0.6–7 mm. Biophys. J. 93, 4360–4373. 28. Visscher K., Gross S. P., and Block S. M. (1996) Construction of multiple-beam optical traps with nanometer-resolution position sensing. IEEE J. Sel. Top. Quant. Electr. 2, 1066–1076. 29. Pralle A., Prummer M., Florin E.-L., Stelzer E. H. K., and Horber J. K. H. (1999) Threedimensional high-resolution particle tracking for optical tweezers by forward scattered light. Microsc. Res. Tech. 44, 378–386.

Chapter 12 Mutational Analysis of Bloom Helicase Xu Guang Xi Abstract DNA helicases are biomolecular motors that convert the chemical energy derived from the hydrolysis of nucleotide triphosphate (usually ATP) into mechanical energy to unwind double-stranded DNA. The unwinding of double-stranded DNA is an essential process for DNA replication, repair, recombination, and transcription. Mutations in human RecQ helicases result in inherent human disease including Bloom’s syndrome, Werner’s syndrome, and Rothmund-Thomson syndrome. Bloom’s syndrome (BS) is a rare human autosomal recessive disorder characterized by a strong predisposition to a wide range of cancers commonly affecting the general population. In order to understand the molecular basis of BS pathology and the mechanism underlying the function of Bloom helicase, we have analyzed BS-causing missense mutations by a combination of structural modeling, site-directed mutagenesis, and biochemical and biophysical approaches. Here, we describe the methods and protocols for measuring ATPase, ATP and DNA binding, DNA strand annealing, and DNA unwinding activities of Bloom protein and its mutant variants. These approaches should be applicable and useful for studying other helicases. Key words: Bloom’s syndrome, missense mutation, unwinding activity, ATPase activity, fluorescence resonance energy transfer (FRET), DNA binding, DNA annealing.

1. Introduction Bloom’s syndrome (BS) is a rare human autosomal recessive disorder characterized by a strong predisposition to a wide range of cancers commonly affecting the general population (1, 2). Cells from persons with BS display chromosomal instability characterized by elevated rates of chromatid gaps, breaks, sister chromatid exchanges (SCEs), and quadriradials. The BS protein (BLM) is 1417 amino acids in length (3). It shows strong homology to Escherichia coli RecQ family helicases in its central domain and has several catalytic activities. BLM is a DNA-dependent ATPase M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_12, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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and utilizes the energy derived from ATP binding or hydrolysis to unwind the canonical Watson-Crick duplex and alternative DNA structures including Holliday junction, double Holliday junction (crossing over during homologous recombination), and the highly stable G-quadruplex (4–6). In addition to its DNA unwinding activity, BLM displays a strand annealing activity, which is necessary for the correction of the genomic instability of BLM cells (7). More recently, it was shown that BLM disrupt the Rad51-ssDNA filament by dislodging human51 protein from ssDNA and stimulate DNA repair synthesis (8). Different disease-linked mutations have been identified in the BLM gene (9). The most frequent mutations include introduction of stop codons into the open reading frame, insertions and deletions that generate frameshift mutations, and missense mutations (9). By combination of several approaches including biochemical, biophysical, and molecular modeling methods, we have analyzed several disease-causing missense mutant proteins identified in BS patient (10–12). The analyses of these mutants not only enable us to reveal the molecular basis of BLM disease-causing mutation and the conserved mechanisms employed by the RecQ family helicases but also could allow us to interpret new mutations identified in the future.

2. Materials 2.1. Protein Purification 2.1.1. Equipment

1. French Press (Bioritech, France). 2. Sonicator 3000 (Misonix, Inc). ¨ KTApurifier (GE Health, France). 3. A 4. Ni-NTA resin (Novagen). 5. Hiload Superdex 200 prep grade chromatography column (GE Health, France).

2.1.2. Chemical Reagents and Solutions

1. EDTA-free protease inhibitors (Roche Molecular Biochemicals, France). 2. Binding buffer (8 ): 40 mM imidazole, 4 M NaCl, 160 mM Tris–HCl, pH 7.9. 3. Wash buffer (4 ): 480 mM imidazole, 4 M NaCl, 160 mM Tris–HCl, pH 7.9. 4. Elute buffer (4 ): 4 M imidazole, 2 M NaCl, 80 mM Tris– HCl, pH 7.9. 5. Gel filtration buffer: 0.5 M NaCl, 10% (v/v) glycerol, 20 mM Tris–HCl, pH 7.9.

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All the above solutions were filtrated through a 0.22 mm pore Millipore membrane (type GS) and were stored at 4C. 2.2. ATPase

1. Wallac 1414 Win Spectral Liquid Scintillation Counter (PerkinElmer).

2.2.1. Equipment

2. Water bath.

2.2.2. Reagents

1. Solution A: ammonium molybdate (Sigma) was dissolved at 8.1 mM in 0.8 N HCl and stored at room temperature (see Note 1). 2. Solution B: 25% (v/v) cyclohexane, 25% butanol-2, 5% acetone, and 1% solution A. Stored in dark at room temperature (see Note 2). 3. 20 mM phosphoric acid (Fluka). 4. [g-32P]ATP (New England Nuclear). 5. Ultima Gold Cocktails (PerkinElmer). 6. 0.5 M ATP solution (see Note 3). 7. ATPase buffer (10 ): 0.2 M Tris–HCl, pH 8.0, 30 mM MgCl2, 5 mM DTT.

2.3. Fluorescence Anisotropy Based DNA Unwinding Assay

1. Beacon 2000 polarization instrument (Panvera Corp). 2. High quality of PAGE-purified unlabeled and 50 - or 30 -fluorescein-labeled oligonucleotides (Eurogentec). 3. 0.5 M ATP solution (see Note 3). 4. Unwinding buffer (10 ): 250 mM Tris–HCl (pH 8.0), 300 mM NaCl, 30 mM magnesium acetate, and 1.0 mM DTT.

2.4. DNA Binding

1. Beacon 2000 polarization instrument (Panvera Corp). 2. 50 - or 30 -fluorescein-labeled oligonucleotides (Eurogentec). 3. DNA binding buffer: 20 mM Tris–HCl, 50 mM NaCl, 1 mM MgCl2, 0.1 mM DTT.

2.5. ATP Binding

1. Bio-Logic auto-titrator (TCU-250) and Bio-Logic optical system (MOS-450/AF-CD) (Bio-Logic, France). 2. N-methylisatoic anhydride and ATP (ACROS). 3. Sephadex LH-20 (Amersham) column (2.282 cm, packed in water). 4. 101040 mm3 quartz cuvette. 5. ATP binding buffer: 20 mM Tris–HCl, 50 mM NaCl, 0.1 mM DTT.

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2.6. Strand Annealing

1. Bio-Logic stopped-flow (SFM-400) and Bio-Logic optical system (MOS-450/AF-CD) (Bio-Logic, France). 2. 30 -fluorescein-labeled and 50 -hexachlorofluorescein-labeled oligonucleotides (Eurogentec). 3. DNA annealing buffer: 20 mM Tris–HCl, 50 mM NaCl, 1 mM MgCl2, 0.1 mM DTT.

3. Methods 3.1. Protein Purification

1. A plasmid for producing the BLM helicase core consisting of amino acid residues 642–1290 was generated by inserting the corresponding gene between the NdeI and XhoI sites of the expression plasmid pET15b (Novagen). All missense mutations were constructed by ‘‘splicing by overlap extension’’ as described (13) with the desired mutations in the internal mutagenic primers (see Note 4). The resulted plasmids were transformed into E. coli strain BL21-CodonPlus (Stratagene). 2. A single colony was grown overnight in 10 ml of LB containing 100 mg/ml ampicillin and 34 mg/ml chloramphenicol at 37C with an agitation speed of 250 rpm. 3. An aliquot of 0.1 ml of the E. coli cell culture was diluted into 1 l of pre-warmed LB containing 100 mg/ml ampicillin and 34 mg/ml chloramphenicol. The cells were grown to the midexponential phase (A600 of 0.5–0.6) at 37C. 4. Protein production was induced by addition of isopropyl-1thio--D-galactopyranoside to a final concentration of 0.25 mM, and the culture was incubated with shaking at 18C for 18 h. 5. The cells were harvested by centrifugation and suspended in a final volume of 2530 ml of 1 binding buffer. 6. Cells were lysed by passage through a French pressure cell and the samples were sonicated to reduce viscosity. To remove any insoluble materials, the cell lysate was centrifuged twice at 23,426g for 45 min. The soluble extract was filtrated through a 0.45 mm filter (see Note 5) and was then applied to a column containing 20 ml of Ni-NTA resin. 7. The subsequent purification procedures were performed with ¨ KTA Purifier) at 18C. The column was washed FPLC system (A with 1 binding buffer at a flow rate of 3.5 ml/min until the UV absorbance at 280 nm became stable. Bound proteins were eluted with 300 ml linear gradient of imidazole (0.02–0.4 M). Fractions containing the proteins were identified by SDS-polyacrylamide gel electrophoresis or ATPase assay.

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8. Pooled fractions were concentrated to 2 ml and were further purified by size exclusion chromatography (Superdex 200) at a flow rate of 0.4 ml/min. 9. The purified proteins were electrophoresed on SDSpolyacrylamide gels and visualized with Coomassie Brilliant Blue. The corresponding fraction containing the purified protein was concentrated and the concentration of the purified protein was determined by the Bio-Rad dye method using bovine serum albumin as the standard. 10. The protein was immediately frozen in single use aliquots (20–30 ml) and stored at –80C. 3.2. ATPase Assay

1. In a screw-capped microcentrifuge tube, mix the following: 20 ml 10 ATPase buffer; 2 ml 50-mer ssDNA of 200 nM; 50 ml ATP solution (8 mM) containing 0.25 mCi [g-32P]ATP (see Note 6); Y ml ddH2O to make the final volume 200 ml. 2. Prepare two glass tubes (see Note 7), each containing the same concentration ATP in a total of 200 ml 1 reaction buffer. One tube is referenced as ‘‘reaction tube’’ and other one as ‘‘control tube.’’ 3. Add 25-ml solution of the enzyme at an appropriate concentration to the reaction tube. As a blank, add 25 ml 1 ATPase buffer to the control tube. 4. Mix and incubate at 37C for 10 min. 5. Transfer 150 ml of the above reaction solution into a vial containing 1.5 ml pre-cold solution A and vortex briefly to mix, and further incubate on ice for 2 min. 6. Add 15 ml H3PO4 (20 mM) into each tube and mix. 7. Add 3 ml solution B and vigorously vortex for 30 s. 8. Centrifuge the tubes at 8000g for 2 min to separate the solution into two phases. 9. Take 1 ml of the upper organic phase solution to a vial and add 8 ml AQUASOL solution. 10. Mix well and count the c.p.m. value. 11. Deduct the c.p.m. value of the control tube from that of the reaction tube and determine the specific activity of BLM helicase with the help of the mmol value (see Note 6).

3.3. DNA Unwinding Assay

We developed an assay for monitoring simultaneously the DNA binding and subsequent DNA unwinding in the same sample (14). It is based on the observation that steady-state fluorescence anisotropy (r) is sensitive to the rotational diffusion of the fluorescent molecules. The

Xi

low anisotropy, due to the fast tumbling of a free fluorescein-labeled oligonucleotide in solution, increases abruptly upon binding of the helicase to form a helicase-DNA complex. After initiation of the helicase-catalyzed DNA unwinding, the high anisotropy will return to its low level again as the fluorescein-labeled oligonucleotide is released from the complex. Schematic illustration of the fluorescence anisotropy based unwinding assay and an example for such experiments are presented in Fig. 12.1. A

1) Δr DNA binding

n t io

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Fig. 12.1. (a) Schematic illustration of the unwinding assay based on fluorescence anisotropy. A 30 fluoresecein-labeled oligonucleotide was annealed to the proximal end of an ssDNA molecule. This substrate alone gives an anisotropy value under steady-state condition. Upon helicase binding to the DNA molecule, steady-state anisotropy (r ) increase. The binding amplitude r can be determined from the direct observation of increase in r-value. Upon ATP binding and hydrolysis, helicase unwinds dsDNA and releases the 30 fluorescein-labeled ssDNA. This reaction leads to a decrease in r. The amplitude r2 represent the differences in anisotropy values of ssDNA and dsDNA. (b) DNA unwinding followed by fluorescence anisotropy. Fluorescein-labeled 25-bp duplex DNA (2 nM) was incubated in the unwinding buffer. The anisotropy value increased and then decreased upon addition of 30 nM BLM helicase and 1 mM ATP, respectively.

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1. Set up the sample chamber of Beacon 2000 to 25C (or desired temperature). 2. Prepare a serial dilution of the enzyme such that 1 ml will contain the desired amount of the protein and keep everything on ice. 3. In a test glass tube, make up the required amount of unwinding reaction mix as given below: 15 ml 10 unwinding buffer; X ml partial double-stranded DNA annealed with one strand of 30 - or 50 -fluorescein-labeled oligonucleotide (1–3 nM); Y ml ddH2O to make the final volume 150 ml. 4. Incubate the glass tube in a 25C water bath or other incubation device for 3–5 min. 5. Transfer the test tube to sample chamber (25C) and measure the anisotropy value successively until it is stabilized. 6. Add 1 ml enzyme (see Note 8) to the tube and mix well. The anisotropy value should increase significantly. 7. When the anisotropy value becomes stable again, start the helicase reaction by the addition of 1 ml of 150 mM ATP and record the anisotropy value every 8–10s. 3.4. DNA Binding

1. Put an appropriate quantity of fluorescein-labeled oligonucleotide (for example, 2–5 nM) into the temperaturecontrolled cuvette at 25C, containing 150 ml DNA binding buffer. 2. Titrate the fluorescein-labeled DNA oligonucleotide with increasing protein concentration, ranging from low to high. For example, add 1 ml of an appropriate concentration of protein to the cuvette. 3. Mix well and measure the fluorescence polarization anisotropy every 15 s. 4. When the anisotropy becomes stable, put another microliter of the protein into the tube, mix well, and measure the anisotropy value. 5. Continue the above procedure until the anisotropy value does not increase anymore with increasing protein concentration. 6. Plot the anisotropy values as a function of protein concentration (see Note 9). 7. Determine the equilibrium dissociation constant by fitting the data to an appropriate equation such as the Michaelis– Menten or Hill equations.

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3.5. ATP Binding

I. mantATP preparation 15: 1. Dissolve 1 mmol ATP in 15 ml ddH2O with a magnetic stirring bar and adjust the pH to 9.6 with NaOH. 2. Add 1.5 mmol methylisatoic anhydride to the above solution with continuous stirring. 3. The pH was maintained at 9.6 by titration with 2 N NaOH for 2 h. 4. After adjusting the pH of the reaction mixture to 7.0, the reaction product was applied to a Sephadex LH-20 column and the column was eluted with water at a flow rate of 0.65 ml/min. Fractions of 5 ml were collected. 5. The elution of mantATP can be monitored under an ultraviolet lamp (366 nm) in the dark-room (see Note 10). The corresponding fractions were pooled and were liophylized, then re-dissolved in an appropriate quantity of distilled water (see Note 11). II. mantATP binding measurement 16 1. Add 300 ml of 0.1 mM mantATP in binding buffer in the auto-titrator (TCU-250) and give a titration sequence in the program (for example, injecting 5 ml mantATP solution into the cuvette every 1.5 min for 50 times). 2. Add 1 ml of binding buffer containing 0.5 mM BLM protein to a quartz cuvette (101040 mm3) and put the cuvette into the measurement cell of the auto-titrator. The solution was kept at 25C and stirred continuously by a magnetic stir bar during the whole titration process. 3. Set the excitation wavelength at 280 nm (see Note 12). Start the auto-titration process. The fluorescence signal of mantATP at 440 nm and the absorption of the solution at 280 nm were simultaneously recorded by two detectors. (See Fig. 12.2 for typical excitation and emission spectra of the enzyme and mantATP. The fluorescence of the mantATP molecule will be enhanced upon binding to the enzyme due to the overlap of the emission spectrum of the enzyme and the excitation spectrum of mantATP.) 4. Determine the apparent dissociation constant Kd by fitting the fluorescence intensity at 440 nm (see Note 13) with equation [1]: F ¼ Fs cd þ fd xþ fc

ðx þ 0:5cd þ Kd Þ

qﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃ [1] ðx þ 0:5cd þ Kd Þ2 4 0:5cd x 2

where, Fs is the starting fluorescence of the reaction mixture, fd is the fluorescence coefficient of free mantATP, fc is the fluorescence coefficient of the complex formed, and x is the total concentration of mantATP. cd ¼ V0 =ðV0 þ Vi Þ 1 x=½mantATP is included to

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Fig. 12.2. (a) The overlap of the BLM emission spectrum (lex = 280 nm) and the mantATP excitation spectrum (lem = 440 nm). The excitation wavelength for the mantATP emission spectrum is lex = 345 nm. (b) Changes in fluorescence intensity when 0.5 mM BLM were titrated with increasing concentration of mantATP. Solid lines represent the best fit of the data to equation [1] and molecular structure of mantATP is shown in the insert.

correct accurately for the sample dilution effect, where V0 is the initial sample volume, Vi is the volume of titrant added, and [mantATP] is the mantATP concentration of the titrant. We have developed a new method to measure the helicasemediated DNA/RNA strand annealing activity (Fig. 12.3). This assay is based on the observation that fluorescence

3.6. DNA Strand Annealing Assay

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Fig. 12.3 (a) Principle of DNA-annealing assay based on fluorescence resonance energy transfer (FRET). The annealing was measured by monitoring the decrease in fluorescence at 525 nm caused by FRET between fluorescein and hexachlorofluorescein labels on the oligonucleotide substrates as indicated. Reaction contained 0.5 nM fluorescein-labeled SPFA DNA, 0.5 nM hexachlorofluorescein-labeled complementary SPFB DNA, and 30 nM RECQ5b. (b) Time course of DNA annealing by BLM proteins (30 nM) in the presence of 0.5 nM 30 -fluorescein-labeled ssDNA and 0.5 nM 50 -hexachlorofluorescein-labeled ss DNA. Fraction of annealed DNA was obtained by normalization of fluorescence signal F using the value Fmin as obtained from the calibration measurement that corresponded to 100% annealing (see Sections 2 and 3), i.e., fraction annealed ¼ (Fs–F)/(Fs–Fmin), where Fs is the fluorescence signal at the start of annealing.

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resonance energy transfer (FRET) is a distance-dependent interaction between the electronic excited states of two dye molecules, in which the excitation is transferred from the donor molecule to the acceptor molecule without emission of a photon from the donor. FRET should occur when donor (fluorescein) and acceptor (hexachloroflurorescein) covalently attached to the 30 and 50 ends of the complementary oligonucleotides, respectively, are within a distance of 10–50 A˚, resulting to a reduced fluorescence of the donor. Therefore, the helicase-catalyzed DNA strand annealing can be followed by monitoring the FRET increase (or the donor fluorescence decrease) in real time. The obtained kinetic parameters from such studies could be very useful for understanding molecular mechanism of helicase-mediated strand annealing.

1. The reaction was performed in a three-syringe-mixing mode, where 300 ml BLM protein (300 nM) in annealing buffer was in syringe #1, 300 ml 30 -fluorescein-labeled (6 nM) and 300 ml 50 -hexachlorofluorescein-labeled (6 nM) oligos in annealing buffer were in syringes #2 and #3, respectively. 2. Set the excitation wavelength at 492 nm. 3. Mix equal volumes (60 ml) of samples from three syringes to initiate the reaction and measure the fluorescein emission signal at 525 nm, which should decrease due to FRET. The assay was performed at 25C. 4. Six thousand data points were collected in 30 min for each kinetic time course. 5. For converting the output signal change from volts to percentage of annealing, a control reaction with a two-syringemixing mode was performed, where 300 ml BLM protein (200 nM) in annealing buffer was in syringe #1 and 300 ml pre-hybridized duplex (4 nM) in annealing buffer was in syringe #2. 6. Initiate the control reaction by mixing the samples from the two syringes under the same experimental condition as in the case for annealing reaction. 7. Data from the annealing reaction was normalized by using the voltage (corresponding to 100% annealing) obtained in the control measurement. 8. Fitting the curve with an appropriate equation (single or multiple exponentials) to determine the kinetic parameters.

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4. Notes 1. Ammonium molybdate is harmful if swallowed or inhaled. It is advised to handle the product in a ventilate area and wear appropriate personal protective equipment. To completely dissolve ammonium molybdate in 0.8 N HCl, the solution needs to be stirred overnight. 2. This mixture can be stored at room temperature for several months or even years without significant decline in quality if the liquid container bottle is closed well. 3. ATP dissolved in pure water is a very acidic solution. The pH should be carefully adjusted to 7.0–7.5 with NaOH. The final concentration of ATP can be determined spectrophotometrically using an extinction coefficient at 259 nm of 15.4103 M–1 cm–1. 4. To avoid introducing undesired mutations, the resulting plasmids should be sequenced. 5. When using the syringe driven filter (0.45 mm, Millex1) to filter the soluble extract, the solution must be sonicated carefully and centrifuged two times. If the above procedures were not well done, the filter can be blocked quickly and frequent change of the filter will be obligated, which is time- and material-consuming. 6. We usually prepare 500 ml of 8 mM ATP solution containing 2.5 mCi [g-32P] ATP to make a final concentration of 2 mM ATP in the reaction mixture. One unit of ATPase activity is defined as the amount of BLM protein required to hydrolyze one micromole ATP per minute at 37C. The c.p.m. value corresponding to 1 mmol ATP can be determined by counting 125 ml of 8 mM ATP solution containing 0.25 mCi [g-32P] ATP. 7. For the sake of achieving the temperature rapidly, it is most preferred to use glass tubes rather than eppendorf tubes. 8. The appropriate protein concentration should be determined experimentally. Protein at very high concentration will immediately bind the released fluorescein-labeled oligonucleotide (14) and the helicase activity can be sub-estimated. 9. When the fluorescence intensities of the free and bound sates are significantly different, the apparent anisotropy value must be corrected. Please consult the instruction manual of the Beacon 2000 (Panvera). 10. The mantATP has brilliant blue fluorescence, while N-methylanthranilic acid shows violet fluorescence.

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11. The concentration of mantATP can be determined spectrophotometrically using an extinction coefficient at 255 nm of 23103 M–1 cm–1. 12. The instrument is equipped with a 150-W Xe–Hg lamp and the monochromator was set with a slit width of 1 mm. 13. As the absorption of the sample for the excitation light (280 nm) may increase significantly with increasing mantATP concentration, the measured fluorescence intensity at 440 nm should be corrected for the inner filter effect by using the measured absorption at 280 nm for each data point. References 1. Mohaghegh P., and Hickson I. D. (2003) The Bloom’s syndrome helicase: keeping cancer at bay. Biologist (London) 50, 29–33. 2. Cheok C. F., Bachrati C. Z., Chan K. L., Ralf C., Wu L., and Hickson I. D. (2005) Roles of the Bloom’s syndrome helicase in the maintenance of genome stability. Biochem. Soc. Trans. 33, 1456–1459. 3. Ellis N. A., Groden J., Ye T. Z., Straughen J., Lennon D. J., Ciocci S., Proytcheva M., and German J. (1995) The Bloom’s syndrome gene product is homologous to RecQ helicases. Cell 83, 655–666. 4. Karow J. K., Chakraverty R. K., and Hickson I. D. (1997) The Bloom’s syndrome gene product is a 30 –50 DNA helicase. J. Biol. Chem. 272, 30611–30614. 5. Sun H., Karow J. K., Hickson I. D., and Maizels N. (1998) The Bloom’s syndrome helicase unwinds G4 DNA. J. Biol. Chem. 273, 27587–27592. 6. Wu L. and Hickson I. D. (2003) The Bloom’s syndrome helicase suppresses crossing over during homologous recombination. Nature 426, 870–874. 7. Cheok C. F., Wu L., Garcia P. L., Janscak P., and Hickson I. D. (2005) The Bloom’s syndrome helicase promotes the annealing of complementary single-stranded DNA. Nucleic Acids Res. 33, 3932–3941. 8. Bugreev D. V., Yu X., Egelman E. H., and Mazin A. V. (2007) Novel pro- and antirecombination activities of the Bloom’s syndrome helicase. Genes Dev. 21, 3085–3094. 9. German J., Sanz M. M., Ciocci S., Ye T. Z., and Ellis N. A. (2007) Syndrome-causing mutations of the BLM gene in persons in the Bloom’s Syndrome Registry. Hum. Mutat. 28, 743–753.

10. Guo R. B., Rigolet P., Zargarian L., Fermandjian S., and Xi X. G. (2005) Structural and functional characterizations reveal the importance of a zinc binding domain in Bloom’s syndrome helicase. Nucleic Acids Res. 33, 3109–3124. 11. Ren H., Dou S. X., Rigolet P., Yang Y., Wang P. Y., Amort-Gueret M., and Xi X. G. (2007) The arginine finger of the Bloom syndrome protein: its structural organization and its role in energy coupling. Nucleic Acids Res. 35, 6029–6041. 12. Guo R. B., Rigolet P., Ren H., Zhang B., Zhang X. D., Dou S. X., Wang P. Y., Amort-Gueret M., and Xi X. G. (2007) Structural and functional analyses of disease-causing missense mutations in Bloom syndrome protein. Nucleic Acids Res. 35, 6297–6310. 13. Horton R. M., Hunt H. D., Ho S. N., Pullen J. K., and Pease L. R. (1989) Engineering hybrid genes without the use of restriction enzymes: gene splicing by overlap extension. Gene 77, 61–68. 14. Xu H. Q., Zhang A. H., Auclair C., and Xi X. G. (2003) Simultaneously monitoring DNA binding and helicase-catalyzed DNA unwinding by fluorescence polarization. Nucleic Acids Res. 31, e70. 15. Hiratsuka T. (1983) New ribose-modified fluorescent analogs of adenine and guanine nucleotides available as substrates for various enzymes. Biochim. Biophys. Acta. 742, 496–508. 16. Liu J. L., Rigolet P., Dou S. X., Wang P. Y., and Xi X. G. (2004) The zinc finger motif of Escherichia coli RecQ is implicated in both DNA binding and protein folding. J. Biol. Chem. 279, 42794–42802.

Chapter 13 In Vivo Analysis of Drosophila BLM Helicase Function During DNA Double-Strand Gap Repair Mitch McVey Abstract The BLM helicase is a member of the RecQ DNA helicase family and is mutated in the cancer-prone disorder Bloom syndrome. BLM plays a role in a number of cellular processes including DNA doublestrand break repair, Holliday junction dissolution, and chromosome segregation. In Drosophila melanogaster, the BLM ortholog (DmBlm) is encoded by the mus309 gene. To study the role of DmBlm in double-strand break repair, we utilized a genetic assay in which a targeted DNA double-strand gap is created through excision of a P transposable element. By recovering and molecularly analyzing individual repair products from wild-type and mus309 male pre-meiotic germline cells, we demonstrated that the DmBlm helicase is involved in homologous recombination downstream of strand invasion. This assay can be adapted to test the roles of numerous DNA metabolic factors in DNA double-strand gap repair. Key words: Bloom syndrome, DNA helicase, homologous recombination, P element, D-loop disruptase, double-strand break repair.

1. Introduction Bloom syndrome is an autosomal recessive disorder characterized by small stature, immunodeficiency, genome instability, and cancer (1). It is caused by mutation of BLM, which encodes a RecQ DNA helicase with 30 to 50 polarity (2). BLM has been implicated in multiple processes that promote genome stability, including Holliday junction dissolution (3), D-loop unwinding (4), and segregation of chromosomes during anaphase (5). Loss of BLM also increases the incidence of spontaneous mitotic crossovers and sister chromatid exchanges (6). Mutation of the Drosophila melanogaster BLM ortholog, DmBlm, results in female subfertility and M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_13, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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sensitivity to ionizing radiation (7). We have utilized an in vivo P element excision assay to demonstrate that mus309 mutant flies lacking DmBlm helicase activity are deficient in DNA doublestrand gap repair (8). This assay uses the P{wa} transposon, a 14-kilobase P element containing the wa allele (Fig. 13.1). Male flies possessing P{wa} and a transposase source are created through standard genetic crosses. Transposon excision in these males creates a double-strand gap that can be repaired by several repair pathways. Individual repair events occurring in pre-meiotic germline cells are recovered in the progeny of these males and assigned to a repair pathway based on eye color. Aberrant repair events are subsequently analyzed by PCR and DNA sequencing to determine the lengths of repair synthesis tracts. Using this assay, we have demonstrated that the DmBlm helicase is required for efficient gap repair by synthesis-dependent strand annealing, a particular mechanism of homologous recombination repair that does not result in the formation of crossovers. Repair products isolated from DmBlm mutant males have short repair synthesis tracts and are frequently accompanied by deletions into flanking genomic sequence. One interpretation of these

Fig. 13.1. The P{wa} assay. (a) Schematic of P{wa} inserted into scalloped (sd) on the X chromosome. Narrow black rectangles indicate the left and right ends of P{wa}. The white (w) gene (dark gray rectangles) is interrupted by a 4.6-kilobase copia retrotransposon (long black rectangle), flanked by 276-bp-long terminal repeats (LTRs, white boxes). (b) The broken DNA ends remaining after excision of P{wa} have noncomplementary, 17-nucleotide 30 overhangs that can prime repair synthesis from the sister chromatid in the somatic and germline cells of parental males. Different types of repair result in different eye colors in female progeny inheriting the repair event from their fathers and an intact copy of P{wa} from their mothers. (c) Complete repair by homologous recombination (HR) or a failure to excise results in apricot eyes. (d) HR repair with annealing at the long terminal repeats of copia results in flat red eyes, due to increased expression of the white gene. (e) Aberrant repair involving HR followed by end joining (or end joining without HR) results in yellow eyes. (Adapted from ref. 8 with permission from Science.)

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findings is that DmBlm is involved in disruption of D-loop intermediates formed during homologous recombination, and in its absence unidentified endonucleases and/or exonucleases act to process the D-loops, resulting in flanking deletions (9). This versatile assay can also be utilized to discern the in vivo functions of other proteins involved in gap repair and to identify proteins that may be involved in repair pathway choice.

2. Materials 2.1. Creation of Males for Gap Repair Studies and Scoring of Progeny and Assignment of Repair Events

1. The P{wa} stock was obtained from the lab of Steve Mount (10). Other fly stocks, including the transposase stock y[1] w[*]; CyO, H{w[+mC]=P2–3}HoP2.1/Bc[1], the mus309D2 and mus309D3 alleles, and balancer stocks, can be obtained from the Bloomington Drosophila Stock Center, Bloomington, Indiana. 2. Drosophila solid culture medium. Our recipe contains yellow cornmeal (SciMart Inc.), Drosophila yeast nutrient flake (SciMart, Inc.), soy flour, Karo light corn syrup, Drosophila agar (Genesee Scientific), propionic acid, and Tegosept (Genesee Scientific). The food is prepared according to the protocol published on the Bloomington Stock Center website (http://flystocks.bio.indiana.edu/).

2.2. Molecular Characterization of Aberrant Repair Events

1. Squishing buffer: 10 mM Tris-Cl, pH 8.2, 1 mM EDTA, 25 mM NaCl. Use 50 mL per fly; add 1 mL proteinase K from 10 mg/mL frozen stock (ThermoFisher Scientific) (11). 2. 10 PCR buffer (500 mM KCl, 100 mM Tris–HCl, pH 9.0, 1% Triton X-100, 25 mM MgCl2). 3. 10 mM deoxynucleotide triphosphate mix (Sigma-Aldrich). 4. Taq polymerase (Sigma-Aldrich). 5. Oligonucleotides: 50 mM in ddH2O(Integrated DNA Technologies). SdF: 50 CCCTGTCTGAAGTTCCGTAG 30 SdR: 50 CCCTCGCAGCGTACTATTGAT 30 Pout: 50 CCGCGGCCGCGGACCACCTTATGTTATTTC 30 920R: 50 AGATGGGTGTTTGCTGCCTCCG 30 1487F: 50 CGTTGTTTGCACGTCTCGCTCG 30 2420R: 50 GAGCGAGATGGCCATATGGCTG 30 4287F: 50 GCAACGAGCGACACATACCG 30 4674R: 50 GGACTGGGCCCATAACCTGTTG 30

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6. Agarose HS, molecular biology grade (Denville). 7. 50 TAE: 4 M Tris, 11.4% acetic acid, 0.1 M EDTA. 8. Ethidium bromide: 1% solution (Fisher Biotech). 9. QIAquick gel extraction kit (Qiagen).

3. Methods The P{wa} double-strand gap repair assay involves excision of a P element containing wa, an allele of the white gene with a copia retrotransposon inserted in an intron. This allele results in apricot eye color in hemizygous males and homozygous females and yellow eye color in hemizygous females, due to decreased expression of white (Fig. 13.1). In the version of the assay described here, P{wa} is inserted in an intron of the essential scalloped (sd) gene (12), located at cytological position 13F on the X chromosome. Transposase-catalyzed excision results in a 14-kilobase gap that is preferentially repaired by homologous recombination, using the sister chromatid as a template. Due to the extreme requirement for repair synthesis, homologous recombination often aborts and repair is frequently completed by an end-joining pathway (13). The type of repair event (homologous recombination vs. end joining) is scored based on eye color (Fig. 13.1). Repair events from germlines of individual males are totaled and percentages of repair pathways utilized are calculated on a per vial basis, thereby allowing for direct comparison between wild-type and mutant backgrounds. 3.1. Creation of Males for Gap Repair Studies

The first step of the assay involves the creation of males possessing one copy of both the P{wa} transposon and the 2–3 transposase through standard genetic crosses (Fig. 13.2a, also see Note 1). 1. Generate a stock of flies that is homozygous for the P{wa} transposon and heterozygous for the mutation that you are testing. In the case of mus309, the appropriate genotype is P{wa}; mus309D2/Balancer. 2. Collect 20–25 virgin females from this stock and mate them with 10–15 males of the genotype w; +/CyO, H{w+, 2–3}; mus309D3/Balancer in bottles containing 25 mL of food (see Note 2). Incubate these crosses at 25C for 3–4 days. Incubation at lower temperatures results in decreased excision rates. 3. On the same day, mate 20–25 P{wa} virgin females with 10– 15 males of the genotype w; +/CyO, H{w+, 2–3}. This cross serves as the wild-type control.

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Fig. 13.2. Cross schemes for the P{wa} assay when testing the role of the DmBlm helicase in double-strand gap repair. (a) Females that are homozygous for P{wa} and heterozygous for one allele of mus309 are mated with males that possess one copy of P transposase and are heterozygous for a different allele of mus309. Male progeny transheterozygous for mus309 mutations that also inherit P{wa} and the P transposase are backcrossed to homozygous P{wa} females. Eye color (and therefore proportions of different repair events) is scored in the F2 female progeny that do not inherit transposase. (* represents the P{wa} repair event) (b) To recover independent repair events for molecular analysis, one yellow-eyed female obtained from each F1 cross in (a) is crossed to 2–3 males with an X balancer chromosome (FM7w in this cross). Whiteeyed male progeny possess a single copy of the repair event and can be processed for molecular analysis.

4. The parents can be transferred into new bottles every 3–4 days up to three additional times to increase the yield of F1 progeny. 5. Collect F1 males of the desired genotype possessing both P{wa} and the P transposase. These males will have mosaic eyes (with red and yellow patches on a background of apricot) due to excision of P{wa} and repair in somatic cells (including those of the developing eye). 6. Hold these males for 1–3 days at 25C. 7. To sample repair events from the pre-meiotic germline of these males, mate individual males with 3–5 homozygous P{wa} females (see Note 3) in vials containing 10 mL of food. Incubate these crosses at 25C. 8. Remove the parents 4–6 days after the initiation of egg laying, once third instar wandering larvae on the sides of the vials are observed (see Note 4).

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3.2. Scoring of Progeny and Assignment of Repair Events

1. Begin scoring the phenotypes of the F2 female progeny 12 days after setting up the cross and continue scoring every other day until day 20, at which point you should discard the vial to prevent inclusion of F3 progeny in the data set. 2. Score eye color in females lacking the transposase (as determined by absence of the linked dominant marker). These females will inherit one copy of P{wa} from their mothers and the repaired copy of P{wa} from their fathers. Three distinct eye colors will be observed (see Note 5), representing three distinct repair mechanisms (Fig. 13.1c–e): l Apricot, which results from accurate repair of the doublestrand gap by homologous recombination (or failure of P{wa} to excise). l

Flat red, which results from homologous recombination repair involving synthesis of the white coding sequence and annealing of newly synthesized copia LTR sequences.

Yellow, which results from incomplete synthesis of the white coding sequence followed by end joining repair. 3. The collective eye color data can be reported in two ways: l As an unweighted average, in which the overall percentages of red- and yellow-eyed female progeny are calculated relative to the total number of females counted, or l

As a weighted average, in which the individual percentages of red and yellow-eyed females are calculated relative to the total number of females lacking transposase in each vial. This method allows for the calculation of mean and standard error for each repair class. A Kruskal-Wallis (non-parametric ANOVA) test can then be performed to determine whether significant differences exist between the weighted averages of red- and yellow-eyed females obtained from the wild-type vs. mutant crosses. 4. Score wing phenotype in yellow-eyed F2 females lacking the transposase. Females in which repair resulted in a large flanking deletion that removes scalloped coding sequence will have scalloped wings (Fig. 13.3). l

5. Female F2 progeny with yellow eyes can be further characterized to determine the exact nature of the repair event, as described below (Fig. 13.2b). 6. Mate one yellow-eyed female from each progeny vial to 3–4 males with an X chromosome balancer, such as FM7w (see Note 6). 7. Remove the parents 6–8 days after the initiation of egg laying, once third instar wandering larvae on the sides of the vials are observed.

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Fig. 13.3. The scalloped wing phenotype indicates deletion into sequences flanking P{wa} during repair. (a) Wild-type wing structure in a female fly. (b) A mild scalloped-wing phenotype resulting from a small deletion of sd (the female is heterozygous for this mutation). (c) A severe scalloped-wing phenotype resulting from a large deletion of sd (again, the female is heterozygous).

8. Desiccation of vials at this stage is common; adding 100– 250 ml of distilled water to each vial after removal of the parents can alleviate this problem. 9. Three possible results will be obtained from these crosses: l Orange- and white-eyed male progeny with wild-type wings will be recovered in approximately equal numbers, indicating that the repair event does not involve a flanking deletion into scalloped. l

White-eyed male progeny with scalloped wings will be recovered, indicating a repair event involving a non-lethal deletion into scalloped.

No white-eyed male progeny will be recovered (only orange-eyed males will survive), indicating a repair event involving a lethal deletion into scalloped. This frequently occurs when the female parent had scalloped wings. 10. In vials where white-eyed male progeny are observed, collect one white-eyed male from each vial in a 0.5-mL eppendorf tube and freeze at –20C for at least 1 h (see Note 7). l

3.3. Molecular Characterization of Aberrant Repair Events

1. Prepare genomic DNA from each white-eyed male. Place one frozen fly in a 0.5-mL tube and mash the fly with a pipette tip containing 50 mL of squishing buffer + proteinase K, but do not expel any liquid until after squishing (see Note 8). 2. Incubate at 37C (or room temperature) for 30 min. This step and step 3 are conveniently done in a thermal cycler.

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3. Inactivate the proteinase K by heating to 95C for 1–2 min (see Note 9). 4. Perform PCR on genomic DNA isolated from individual males to determine amount of DNA synthesis in each repair event. PCR is done in a 20-mL reaction volume, with 2 mL of 10 PCR buffer, 1 mL of genomic DNA, 0.5 mL dNTPs, 0.5 mL of each primer, and 0.2 mL of Taq polymerase. Primer pairs will amplify products 120 bp to 1 kb in length (see Note 10). 5. Prepare a 1% TAE-agarose gel by dissolving 1 g of agarose in 100 mL of 1 TAE in a microwave. Add 2 mL of 1% ethidium bromide once the gel has cooled to 70C but before it has polymerized. 6. Load 10 mL of PCR product into each well of the gel. Connect to a power supply and run for approximately 1 h at 90 V. Visualize the PCR products under ultraviolet light. 7. The presence of a band of the expected size indicates that the repair event involved DNA synthesis through the end of the reverse primer. Synthesis tract lengths for each repair event can be estimated by compiling the results of each set of PCR reactions. Examples of synthesis tract lengths for repair events isolated from wild-type and mus309 mutant flies are shown in Fig. 13.4. 8. In cases where synthesis tract lengths from both sides of P{wa} are short (<1 kb), the exact amount of repair DNA synthesis can be determined by performing PCR using primers SdF and SdR. After electrophoresis of these PCR products, excise the

Fig. 13.4. Repair synthesis tract lengths in wild-type and mus309 mutant flies. Repair synthesis from the right end of P{wa} was analyzed for 71 independent aberrant repair events from wild-type flies and 147 aberrant events from mus309 mutants. The percentages that displayed synthesis tracts of at least the lengths indicated are shown in the histogram. The primer pairs corresponding to each tract length (see Section 2) and the expected size of each PCR product are indicated. (Adapted from ref. 8 with permission from Science.)

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bands from the gel while under UV illumination using a razor blade (wearing goggles and a face shield). Place each gel slice in a 1.5-mL eppendorf tube and store at 4C. 9. Purify the DNA from each gel slice using a gel extraction kit. Resuspend in 30 mL of distilled water. 10. Sequence 20 mL of the purified DNA using either of the two primers used to generate the PCR product. Using alignment software, compare the sequence to the wild-type P{wa} sequence to determine the exact amount of repair synthesis in the repair event.

4. Notes 1. P{wa} insertions on autosomes can also be used. In this case, homologous recombination repair utilizing the homolog as a template can occur. Similar to end-joining repair, homologous recombination from the homolog results in the generation of yellow-eyed F1 female progeny. These two classes of repair products can be differentiated using PCR-based assays. Furthermore, the homolog can be distinguished from the P{wa} insertion chromosome by including one or more flanking dominant markers on the insertion chromosome. 2. If desired, an alternate transposase source, such as P{w+, 2–3 99B} on chromosome 3, can be used. We have found that this transposase source is more active in somatic cells but less active in the male pre-meiotic germline. 3. Alternatively, you can cross the males to attached-X females (C(1)DX). This allows the male progeny to inherit the repair event directly from their fathers and allows for direct molecular analysis of the repair product. However, any repair events that delete into the essential scalloped gene will not be recovered, as they are lethal in males. 4. If desiccation of the food in the vials occurs, add a small amount (100–250 mL) of distilled water to the vials after removing the parents. If the food liquefies due to excessive larval chewing action, a piece of Kimwipe can be added to the vial to absorb excess liquid. 5. Occasionally, progeny with dark orange/light red eyes are observed. These are likely due to transposition events in which P{wa} transposes to a new genomic location and are not counted as repair events in the final analysis.

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6. The yellow-eyed females need not be virgins. Their male progeny will inherit one of their two X chromosomes, containing either P{wa} or P{wa}* (the repair event). They are crossed to FM7w males so that the repair events can be subsequently maintained in females over an X chromosome balancer. 7. These white-eyed males should lack the transposase source to prevent the initial P{wa} repair event from being subjected to additional rounds of excision and repair. 8. If working with large numbers of flies, the genomic DNA isolation can also be done in a 96-well plate. 9. The genomic DNA preparation can be stored at 4C for months with no substantial loss of DNA integrity. 10. If desired, additional primer pairs can be designed within the wa sequence to test for repair synthesis at other regions of P{wa}. References 1. German J. (1993) Bloom syndrome: a mendelian prototype of somatic mutational disease. Medicine 72, 393–406. 2. Ellis N. A. et al. (1995) The Bloom’s syndrome gene product is homologous to RecQ helicases. Cell 83, 655–666. 3. Wu L. and Hickson I. D. (2003) The Bloom’s syndrome helicase suppresses crossing over during homologous recombination. Nature 426, 870–874. 4. van Brabant A. J. et al. (2000) Binding and melting of D-loops by the Bloom syndrome helicase. Biochemistry 39, 14617–14625. 5. Chan K. L., North P. S., and Hickson I. D. (2007) BLM is required for faithful chromosome segregation and its localization defines a class of ultrafine anaphase bridges. EMBO J. 26, 3397–3409. 6. German J., et al. (1977) Bloom’s syndrome. IV. Sister-chromatid exchanges in lymphocytes. Am. J. Hum. Genet. 29, 248–255. 7. McVey M., et al. (2007) Multiple functions of Drosophila BLM helicase in maintenance of genome stability. Genetics 176, 1979–1992.

8. Adams M. D., McVey M., and Sekelsky J. J. (2003) Drosophila BLM in double-strand break repair by synthesis-dependent strand annealing. Science 299, 265–267. 9. McVey M., et al. (2004) Formation of deletions during double-strand break repair in Drosophila DmBlm mutants occurs after strand invasion. Proc. Natl. Acad. Sci. U.S.A. 101, 15694–15699. 10. Kurkulos M., et al. (1994) P elementmediated in vivo deletion analysis of white-apricot: deletions between direct repeats are strongly favored. Genetics 136, 1001–1011. 11. Gloor G. B., et al. (1993) Type I repressors of P element mobility. Genetics 135, 81–95. 12. Campbell S. D., et al. (1991) Cloning and characterization of the scalloped region of Drosophila melanogaster. Genetics 127, 367–380. 13. McVey M., et al. (2004) Evidence for multiple cycles of strand invasion during repair of double-strand gaps in Drosophila. Genetics 167, 699–705.

Chapter 14 Purification and Characterization of RecQ Helicases of Plants Daniela Kobbe, Manfred Focke, and Holger Puchta Abstract Helicases are essential for DNA metabolism. Different helicases have different properties tailored to fulfill their specific tasks. RecQ-helicases are known to be important in DNA repair and DNA recombination. In higher organisms several RecQ homologues can be identified. For instance, seven RecQ homologues were identified in the model plant Arabidopsis thaliana. Specialization of those proteins can possibly be reflected by differences in their biochemical substrate spectrum. Moreover, a helicase of interest might be defined by its biochemical properties as a functional ortholog of a RecQ helicase in other organisms. In this chapter the initial steps that will provide the basis for a proper biochemical characterization are given. After the description of the expression of the helicase of interest in the heterologous host Escherichia coli, its purification with the help of two affinity tags and the preparation of a model DNA substrate for the strand displacement assay are described. Finally, it is shown how this model substrate can be used to ensure the purity of the enzymatic preparation of interest. Key words: RecQ, helicase, Walker A motif, Arabidopsis thaliana, Ni-IMAC, His-tag, overexpression, calmodulin binding peptide, calmodulin affinity chromatography, strand displacement assay.

1. Introduction RecQ helicases play an important role in the maintenance of genomic stability and are conserved in all kingdoms of life (1–5). Whereas Escherichia coli possesses just one RecQ homologue, higher eukaryotes usually possess several RecQ homologues. In the model plant Arabidopsis thaliana, seven RecQ homologues were identified (6). Several lines of evidence indicate a functional specialization of RecQ helicases (e.g., (7)). To some extent this specialization will also be reflected in different biochemical properties (e.g., (8)). M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_14, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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Analyzing substrate preferences can help to assign a defined helicase to a specific DNA repair or DNA recombination pathway. In addition, comparing the biochemical properties of RecQ helicases helps to distinguish them and to define interspecies functional homologs. In order to analyze the biochemical properties, first a protein preparation has to be obtained that is enzymatically active but free from contaminant activities. This chapter focuses on how this can be achieved, exemplified with the helicase activity of AtRECQ2 from A. thaliana (9). This protocol has already been successfully applied to other plant helicases. Contaminating activities can be co-purified due to interactions of unwanted proteins with the column matrix, the affinity material, the target protein or by indirect interactions mediated, for instance, by DNA. Therefore the best control is to purify a protein that is as similar as possible to the protein of interest but does not show intrinsic activity, as it is the case for RecQ helicases with a specific amino acid substitution in the helicase motif I, which knocks out ATPase and thereby helicase activity (e.g., (10, 11)). Both protein preparations should be analyzed with all substrates as it is possible that another quite specific helicase or nuclease has been co-purified. A flow diagram is shown in Fig. 14.1. The method described here exploits the T7 RNA polymerase expression system (12) and BL21-CodonPlus1(DE3)-RIPL. The expression vector used here allows the attachment of an

Fig. 14.1. Flow diagram of the basic steps toward the characterization of RecQ helicases of plants.

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N-terminal calmodulin-binding peptide (for calmodulin affinity chromatography (13)), thrombin and enterokinase recognition sites, and a FLAG epitope (for detection in the Western blot) and at the C-terminus, a thrombin recognition site followed by an hexahistidine tag (for Ni-IMAC (14)). During expression, the correct folding of the target protein can be supported by reducing the temperature of expression (also see Note 3). The combination of the two affinity chromatography steps for purification selects full-length proteins and reduces the concentrations of contaminant proteins. A washing step with the detergent Triton-X-100 as well as a washing step of the Ni-IMAC with an optimized imidazole concentration remove contaminant proteins. To determine if the helicase is functional and the preparation is free from contaminants, a DNA substrate providing a 30 and a 50 flap is used and a strand displacement assay is conducted (also see (15, 16)). If the helicase is active the duplex DNA region will be unwound, setting free the composing single-stranded DNA. The different DNA species can be separated via native gel electrophoresis. To provide a high sensitivity the substrate is labeled with P-32. For the strand displacement assay equal amounts of the helicase preparation (as well as dilutions of the presumable active enzyme) and the helicase with helicase motif I mutation are incubated with the substrate in a suitable buffer both in the presence and in the absence of ATP. The controls without ATP are well suited to judge contamination by nucleases. In case the RecQ preparation is enzymatically pure the characterization can be extended to more specialized DNA substrates, such as Holliday junctions, D-loops, or replication forks.

2. Materials 2.1. Expression of the Recombinant Plant Helicase in E. Coli

1. pCAL-n-FLAG (Stratagene) or another suitable vector 2. BL21-CodonPlus1(DE3)-RIPL (Stratagene) or another suitable expression strain 3. LB-medium: 10 g Trypton, 5 g yeast extract, 5 g NaCl, dissolve in ddH2O, adjusted to 1 1, and autoclave 4. LB plates with ampicillin: LB medium with 1.5% (w/v) micro agar, autoclave, then cool down to approximately 50C before adding ampicillin stock solution to a final concentration of 100 mg/ml, pour into petri dishes and store at 4C 5. Antibiotic stocks: ampicillin (50 mg/ml in ddH2O), carbenicillin (50 mg/ml in ddH2O), chloramphenicol (50 mg/ml in ethanol)

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6. 0.4 M isopropyl-ß-D-thiogalactopyranoside (Duchefa) in ddH2O, filter to sterilize (0.22-mm PVDF filter) 2.2. Purification of the Recombinant Helicase by Double Affinity Purification 2.2.1. Preparation of the Soluble Protein Fraction for Affinity Chromatography

1. Buffer A: 20 mM Tris–HCl (pH 7.5), 200 mM NaCl, 20 mM imidazole, 5% glycerol, 10 mM 3-mercapto-1,2-propanediol (thioglycerol). Prepare all buffers for the purification without thioglycerol and filter through a 0.45-mm cellulose acetate filter (0.45-mm Sartorius AG) at 300 mbar with a vacuum pump. Additionally apply 300 mbar for at least 10 min to degas the solution. Store at 4C. 2. Lysozyme (Roche Diagnostics). 3. Sonopuls Ultraschall-Homogenisator HD 2070 (Bandelin). 4. GF/PET filter (Roth).

2.2.2. Ni-IMAC

1. Buffer A as described in Section 2.2.1. 2. Buffer B: components as buffer A, plus 400 mM imidazole. 3. HiTrap chelating HP column (1 ml) (GE Healthcare). 4. Low pressure liquid chromatography system BioLogic LP (BioRad Laboratories). 5. 0.1 M NiSO4 solution (in ddH2O). Caution: toxicity. 6. PD-10 columns, filled with Sephadex G25-M (GE Healthcare). 7. Buffer C: 50 mM Tris–HCl (pH 7.5), 500 mM NaCl, 2 mM CaCl2, 1 mM Mg(CH3COO)2, 1 mM imidazole, 10 mM thioglycerol.

2.2.3. Calmodulin Affinity Chromatography

1. Calmodulin affinity resin for purification of CBP-tagged proteins (Stratagene). 2. PolyPrep1 chromatography column (0.8 4 cm) (BioRad Laboratories). 3. Buffer D: 50 mM Tris–HCl (pH 7.5), 500 mM NaCl, 2 mM EGTA, 10 mM thioglycerol. 4. Buffer E: 50 mM Tris–HCl (pH 7.5), 300 mM NaCl, 10% glycerol, 10 mM thioglycerol. 5. Buffer F: 50 mM Tris–HCl (pH 7.5), 1000 mM NaCl, 2 mM EGTA. 6. Buffer G: 0.1 M NaHCO3, 2 mM EGTA (pH 8.6). 7. Buffer H: 1 M NaCl, 2 mM CaCl2. 8. Buffer I: 0.1 M sodium acetate buffer, 2 mM CaCl2 (adjust pH to 4.4. with acetic acid). 9. Servapor1 dialysis tubing with 16 mm diameter, MWCO 12–14 kDa (Serva). Cut approximately 15 cm and put in ddH2O with EDTA. Heat for at least 20 min at 70C. Store at 4C.

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10. PD-10 columns, (filled with Sephadex G25-M) (GE Healthcare). 2.3. Preparation of the Radioactively Labeled DNA Substrate

1. Oligonucleotide 50 -AAAAAAAAAA GTC GAC TCT AGA GGA TC AAAAAAAAAA-30 ; best results are obtained with PAGE purified oligonucleotides. The middle part of this oligonucleotide is complementary to nucleotides (nt) 6252– 6268 of M13mp18. 2. [g-32P]ATP (3000 Ci/mmol) (GE Healthcare), GE Healthcare no longer manufactures this radioactive material. We now use Hartmann Analytic instead. A license is required to order this radioactive material as well as shielding. 3. T4 PNK and the T4 PNK buffer (New England Biolabs). 4. MicroSpinTM G-25 Columns (GE Healthcare). 5. M13mp18 single-stranded DNA (New England Biolabs). 6. LumaSafe Plus (Lumac LSC) and scintillation vials. 7. Liquid scintillation analyzer Tri-Carb 2100 TR (Packard Instrument Company). 8. MicroSpinTM S-400 HR columns (GE Healthcare) or alternatively SephacrylTM S-400 high resolution resin (GE Healthcare).

2.4. Detection of the Helicase Activity via the Strand Displacement Assay

1. TBE buffer (10 ): 890 mM Tris-base, 890 mM boric acid, 20 mM EDTA, pH 8.0 2. Acrylamide (30% T, 2.67% C) (Roth) 3. N,N,N’,N’-tetramethylethylenediamine (TEMED) 4. 10% ammonium persulfate (APS) (w/v); dissolve in water, aliquot, and store at –20C 5. Whatman Multigel-Long (Biometra) and connected cooling system 6. Reaction buffer (5 ): 200 mM Tris–acetate, 250 mM potassium acetate (pH 8.0) 7. 120 mM DTT, dissolve in ddH2O, aliquot and store at –20C 8. 36 mM ATP (Fluka), calculate the necessary weight according the certificate of analysis (due to water in the chemical), dissolve in ddH2O, aliquot, and store at –20C 9. 1 M stock solution of MgCl2 (Applichem) due to the hygroscopic behavior of MgCl2. Dilute to 36 mM with ddH2O 10. BSA (New England Biolabs) (10 mg/ml), dilute 1:10 with ddH2O, aliquot, and store at –20C 11. PCR chiller 12. Multichannel pipette 13. Heating block for PCR tubes

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14. Stop solution (3 ): 50 mM EDTA, 0.6% SDS, 20% glycerol, 0.1% xylene cyanol, 0.1% bromophenol blue 15. Saran Wrap (Roth) 16. Instant Imager (Canberra Packard Company)

3. Methods 3.1. Expression of the Recombinant Plant Helicase in E. Coli

1. Starting from the coding sequence information of the plant helicase of interest, identify (on the protein level) the helicase domain with its helicase motifs, e.g., by means of multiple sequence alignment with homologous helicases. Introduce a point mutation leading to an amino acid substitution for the conserved lysine of helicase motif I (also called Walker A) to methionine. We usually do this via overlap extension mutagenesis with the appropriately designed primers (17). Clone the ORF of interest and the ORF with the introduced point mutation in an expression vector for heterologous expression in E. coli and verify the construct by sequencing. Choose a vector that exploits the expression by T7 RNA polymerase system and allows the attachment of an N-terminal calmodulin binding peptide tag and a C-terminal His-tag to the protein of interest. For this purpose we modified the 30 end of the MCS of the pCAL-n-FLAG vector (9). 2. Transform the constructs in competent BL21-CodonPlus1(DE3)-RIPL cells by heat shock transformation and plate on LB-plates containing ampicillin. Incubate overnight at 37C. The following day, remove from the 37C incubation and store at room temperature until the evening. 3. Inoculate 500 ml of LB with 75 mg carbenicillin/ml and 34 mg chloramphenicol/ml in a 1 1 Erlenmayer flask with 50 colonies and incubate overnight at 37C by means of shaking (200 rpm) (also see Note 1). 4. The next morning inoculate the appropriate number of 1 l Erlenmeyer flasks containing 500 ml of LB without antibiotics (see Note 2) with 50 ml of the cultures (helicase construct and helicase construct bearing point mutation, respectively) of step 3 and incubate at 28C at 200 rpm. Monitor the optical density at 600 nm of the culture. If the optical density is between 0.6 and 0.9, add IPTG to a final concentration of 0.2 mM (this corresponds to 275 ml of a 0.4 M IPTG stock solution). Reduce the temperature to 16C and continue to incubate by means of shaking at 200 rpm for approximately 20 h (see Note 3).

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5. Harvest the E. coli cells by centrifuging at 2700 g for 10 min and discard supernatant. It may be necessary to repeat the spinning process. At the end, transfer the pellet corresponding to 550 ml culture into one 50 ml polypropylene tube (by resuspending and centrifuging) and store it at –20C. 3.2. Purification of the Recombinant Helicase by Double Affinity Purification 3.2.1. Preparation of the Soluble Protein Fraction for Affinity Chromatography

All purification steps are performed at 4C or on ice.

1. Slowly thaw two identical cell pellets (from step 5 Section 3.1) on ice. Add 10 mM of the reducing agent 3mercapto-1,2-propanediol (thioglycerol) (corresponds to 8.68 ml of thioglycerol per 10 ml of buffer) to the buffers and mix carefully. Carefully resuspend the cell pellets in 5 ml of buffer A, by pipetting over the pellet again and again. When this step has been completed, adjust the volume in each tube to 25 ml with buffer A making use of the scale on the tube. 2. Freshly prepare a lysozyme stock solution of 10 mg/ml in buffer A and add lysozyme to a final concentration of 0.1 mg/ ml (corresponds to 250 ml of the stock solution). Incubate the cell suspension for 30 min on ice by means of shaking. 3. Break up the cells and disrupt the DNA in order to reduce viscosity by sonication. Use six 10-second cycles with 53% power und 50% duty cycle and between cycles cool down the extract on ice for 1 min. 4. Adjust the weight in ultracentrifugation tubes and ultra-centrifuge at 40,000 g for 30 min at 4C. 5. Combine the supernatants from the two pellets and filter through a GF/PET filter (see Note 4).

3.2.2. Ni-IMAC

Each chromatography column should be used for one construct only to prevent cross contamination. The buffers should all be filtered and degassed (approximately 10 min at 300 mbar) and thioglycerol added just before use. The flow rate used, except during loading the extract on the column, is 1 ml/min. 1. Wash the column with ddH2O for at least 5 min to get rid of the ethanol. 2. Charge the column with 0.5 ml of 0.1 M NiSO4 solution (see Note 5). 3. Wash the column with ddH2O for at least 5 min. 4. Wash the column with buffer A for 10 min. 5. Wash the column with buffer B for 10 min.

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6. Equilibrate the column with buffer A for 10 min. 7. Load the filtered supernatant of the ultra-centrifugation step (described in Section 3.2.1) at a flow rate of 0.5 ml/min. 8. Wash the column for 45 min with buffer A containing 0.5% Triton-X-100 9. Wash the column for 15 min with buffer A 10. Wash the column with 31% buffer B for 20–35 min (see Note 6). 11. Elute the protein with 90% of buffer B for 15 min (see Note 7). 12. The column is treated for 20 min with buffer B, subsequently with ddH2O and then with 20% of ethanol for storage. 13. Equilibrate a PD-10 column with 25 ml of buffer C. Then apply 2.5 ml of the fractions containing the target protein. Elute the proteins with 3.5 ml of buffer C. Then adjust the volume of the eluate to 10 ml. 3.2.3. Calmodulin Affinity Chromatography

1. Carefully resuspend the Calmodulin (CaM) affinity resin and fill 2 ml in a PolyPrep chromatography column for gravity flow. This yields 1 ml of bed volume. Drain the liquid. 2. Equilibrate the column with at least 10 ml of buffer C. 3. Apply the diluted eluate of the PD-10 column. 4. Wash the column with 10 ml of buffer C plus 0.5% Triton-X100. 5. Wash the column with 10 ml of buffer C. 6. Elute the proteins in five 1-ml aliquots with buffer D (see Note 8). For this purpose it is best to put the column on a 1.5-ml reaction tube and add 1 ml. Wait until the column has run dry, exchange the tube, and repeat the procedure. The fractions containing the protein are usually mainly the first three with the highest concentration in the second one. They can easily be identified with a Bradford protein assay (micro method) with 30 ml of the fractions. 7. Equilibrate a PD-10 column with buffer E and exchange the buffer of the protein containing fractions to buffer E as described in step 12 of Section 3.2.2. 8. Rinse a prepared dialysis tubing and pipette the eluate of the PD-10 column into it. Close the tubing using suitable clips and put it on a petri dish filled with sucrose. Also, add sucrose on top of the tubing. Incubate at 4C for at least 2.5 h. Exchange the sucrose from time to time. When the volume has been reduced to 300–600 ml, pipette it into a 1.5-ml reaction tube. Mix this concentrated protein solution carefully but thoroughly with the same volume of 100% glycerol

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and store at –20C. Also, prepare a buffer used as a negative control in the strand displacement assay and for dilutions of the finalized enzyme preparation by mixing 500 ml of buffer E with 500 ml of glycerol and store at –20C. 9. Regenerate the CaM column by washing with at least 10 ml of the following buffers each: buffer F, G, H, I, C. Then equilibrate the CaM column in buffer C containing 20% of ethanol for storage at 4C. The PD-10 columns are stored in ddH2O with sodium azide. The purification procedure has to be performed the same way for the helicase of interest and with the helicase with the amino acid substitution introduced in the helicase motif I. The concentration of the two preparations needs to be checked, best via Coomassie staining of an SDS-Gel with BSA as a standard. This information is important to ensure the utilization of the same amount of protein (and therefore also possible contaminants) in the activity assay. Once the purification method is well established, several subsequent purifications with the same protein can be performed and the final preparations can be pooled. Then small aliquots can be made and stored at –80C until usage. It is best to repeat the determination of the concentration of the pooled fractions after aliquoting. 3.3. Preparation of the Radioactively Labeled DNA Substrate

Make sure to comply with the law when working with radioactivity. All steps should be performed behind plexiglas shielding without lead, consisting of a thickness of at least 0.8 cm, to insulate the ß-particles from P-32. It is best to use filter tips for the pipette to prevent contamination. In case you use [g-32P]ATP with a specific activity of 3000 Ci/ mmol and 10 mCi/ml, this corresponds on the reference date to a total concentration of 3.3 pmol ATP/ml made up of approximately 1 pmol/ml [g-32P]ATP and 2.3 pmol/ml ATP without [g-32P]label. 1. Pipette the appropriate volume of ddH2O, 5 ml of 10 PNK buffer, and 15 pmol of the oligonucleotide (e.g., 15 ml of a 1:100 dilution of a 100 pmol/ml stock) into a 1.5-ml reaction tube. Then add 2 ml of the T4 PNK and subsequently, with shielding, 20 pmol of ATP. On the reference date this corresponds to approximately 6 ml. Mix well and incubate at 37C for 1 h. 2. Heat-inactivate the T4 PNK for 20 min at 67C, then let the sample cool down to room temperature and pulse down. 3. Prepare a G-25 spin column to remove the ATP excess. First mix the bead material of the column. Then break it open and slightly unscrew. Centrifuge for 1 min according to the manual; with our special centrifuge which allowing centrifugation

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behind the plexiglas screen from Neolab we centrifuge at 4000 rpm. Then put the column into a new tube, unscrew, and immediately pipette the labeling mixture on it. Close the column gently and centrifuge again. We centrifuge twice for 1 min at 4000 rpm. Discard the column. 4. Mix the eluate well and determine the volume with the help of a pipette. Then pipette 1 ml of the eluate to 4 ml of scintillation cocktail in a scintillation vial together with the tip. Vortex the scintillation vial and let it sit for at least 20 min. Then measure the activity via liquid scintillation counting. Use the direct DPM mode and measure twice for 10 min to ensure a correct value. The specific activity can be calculated, assuming that 100% of the oligonucleotide left the spin column, by the following formula: (activity/ml determined volume of the eluate)/15 pmol. 5. To this tube add 7.5 pmol of M13mp18 single-stranded DNA and T4 PNK buffer to obtain a 1 concentration. In case you use M13 mp18 DNA from New England Biolabs with a concentration of 250 mg/ml, this corresponds to 71 and 13.5 ml of buffer. Incubate at 95C for 5 min then turn off the heating block and let the mixture cool down slowly (approximately 3 h) to room temperature. Then spin for a short time to assemble the whole content on the bottom of the tube. 6. Perform a gel filtration with Sephacryl S-400 spin columns to remove the excess of the labeled oligonucleotide as described in step 3 of this subsection. Do not pipette more than 50 ml on each column. Determine the activity of the eluate as described in step 4 of this subsection (see Note 9). 7. The concentration of the substrate can be calculated dividing the activity/ml (determined in step 6) by the specific activity (determined in step 4). Calculate which volume corresponds to 3 fmol of substrate (also see Note 10). The success of the purification can be verified by mixing 1 ml of the eluate with stop solution and running a native 12% TBEPAGE-Gel as described in the following subsection. For comparison also load the annealed substrate before purification, or heat denature 1 ml of the substrate with stop solution by heating 5 min at 95C before shock cooling on ice. 3.4. Detection of the Helicase Activity via the Strand Displacement Assay

1. Pour two 12% TBE-PAGE gels in the Whatman MultigelLong system. For this purpose assemble the glass plates and the insulation with the help of clamps and mix a solution of 16 ml ddH2O, 3.2 ml 10 TBE buffer, 12.8 ml of acrylamide (30% T, 2.67% C), 15 ml of TEMED, and 224 ml of 10% APS that is poured subsequently between the glass plates, then insert the comb. It is best to use a 16-teethed comb.

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Once the gels are polymerized, remove the insulation and install them in the running chamber with 1 TBE buffer. Prevent air bubbles trapped between the glass plates. Cool the chamber to 4C. 2. Dilute your enzymatic preparations with the dilution buffer prepared in step 8 of Section 3.2.3, yielding equal (maximal) concentrations for the presumably active helicase and the helicase with amino acid substitution. Further dilute the presumably active helicase preparation by factors 2 and 4. Prepare at least 7 ml of each dilution. 3. Prepare two mastermixes (MM), one with ATP and one in which the ATP is omitted and replaced by ddH2O for 18 reactions each. The volumes/amounts for one reaction are the following: 4 ml of reaction buffer (5 ), 1 ml of ATP (20 ), 1 ml of DTT (20 ), 1 ml of BSA (20 ), 1 ml of MgCl2 (20 ), 3 fmol of DNA substrate, ddH2O up to 19 ml. Pipette in the following order: water, buffer, everything except substrate, and with shielding add the substrate and mix very well. Prepare one PCR stripe with five wells and pipette 66.5 ml of the MM with ATP in it and repeat the same for the MM without ATP (see Fig. 14.2). 4. Pipette 1 ml of each enzyme dilution and the dilution buffer twice in triplicates in a PCR-plate pre-cooled to –20C on a PCR chiller (see Fig. 14.2).

Fig. 14.2. Schematic drawing of the procedure for carrying out the strand displacement assay.

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5. Bring the substrate and the enzyme together by adding 19 ml of the respective mastermix to the enzyme on a PCR chiller and mix well with a multichannel pipette. 6. Start the incubation by putting the plate on a 37C block and incubate for 20 min. In the meantime, pipette 1 ml of dilution buffer and 19 ml of the respective leftover mastermix as well as 10 ml of stop solution into two 1.5-ml reaction tubes. Place the tubes in a block pre-warmed to 95C for 5 min. Then place the tubes on ice immediately. Pulse down after cool-down. Now the heat denatured samples are ready for analysis on the gel. 7. Once the incubation time is over, put the reaction plate on a PCR chiller of about 4C. Add 10 ml of stop solution with the multichannel plate in the same order as used for starting the reaction and mix well. When this step is over, place the plate at room temperature. 8. Load 10 ml into each well of the pre-cooled gel and run at 200 V for 30 min (see Note 11). 9. Remove the gels from the electrophoresis chamber. Take off the upper glass plate. Wrap the gel in plastic foil to prevent leakage of radioactivity (see Note 12). 10. Place the gel in the Instant Imager and monitor the appearance of the bands. Wait long enough to be able to detect potential minor bands indicative of nucleolytic degradation.

4. Notes 1. It may be possible to start the overnight culture from a glycerol stock (however we have experienced cases of unstable plasmid in the procedure described here for some constructs). Prepare the glycerol stock by cultivating the transformed bacteria in LB medium containing 0.1% or 1% of glucose at 37C until an OD at 600 nm of approximately 0.3–0.6 is reached. Then add glycerol to a final percentage of 20%, mix, make aliquots and shock freeze in liquid nitrogen. The glycerol stocks are kept at –80C. Inoculate the overnight culture with 0.1% of the glycerol stock. 2. Several lines of evidence have indicated that the addition of antibiotics is not favorable if the method is performed as described. Either the same amount or even less target protein was obtained when antibiotics were added. 3. The optimal incubation temperature and time have to be determined experimentally for each protein of interest. For the approach presented here, it is important to increase the

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concentration of the target protein in the soluble fraction. Therefore, in order to investigate the best conditions for expression, the soluble protein fraction has to be prepared as described in Section 3.2.1. Usually the RecQ helicases are not very well expressed in the soluble fraction but accumulate in inclusion bodies. Therefore, a Coomassie stained gel is generally not sufficient and a Western Blot has to be performed. In order to conclude correctly about proteolytic degradation or incomplete translation, one should also analyze the soluble protein fraction of E. coli cells transformed with the expression vector without the target ORF (sometimes called ‘‘mock control’’). Bands visible in this control are proteins that are detected ‘‘naturally’’ with the antibodies. We usually increase the incubation time when decreasing the incubation temperature and use temperatures of 28C, 21C, or 16C. We could also show that sometimes the addition of 1% glucose to the LB medium greatly improves the yield of the target protein in the soluble fraction. This is sometimes not correlated to the repression of the target protein expression when not induced by IPTG, which is the usual way to explain this phenomenon (18, 19). For the addition of glucose we did not see differences for either autoclaving the LB medium with the glucose or adding the glucose afterwards. The glucose solutions added were either sterilized by autoclaving or by filtration. 4. This step removes cell debris that was not removed by the centrifugation step. The filter, composed of a prefilter of fiber glass and then a 0.45-mm polyester filter, has proved to be useful because otherwise the filters clog fast. 5. For repeated use of the same column the initial conditions can be re-established by stripping off the Ni2+ and recharging the column with Ni2+. In practice after the washing step detailed in 1, the Ni2+ is stripped of for approximately 15 min with stripping buffer (20 mM NaH2PO4, 50 mM EDTA, pH 7.2). Afterwards the column is washed with water until the conductance is again at the water level to make sure that all EDTA is washed out. Then step 2 is performed. 6. 31% of buffer B corresponds to the optimal concentration of imidazole, which does not elute significant amounts of AtRECQ2 but as many E. coli proteins as possible that have bound to the column with less affinity. This imidazole concentration will have to be determined for each target protein. For this purpose perform all steps up to step 9 and then run a linear gradient of buffer B instead of the two step elution protocol. Calculate the optimal concentration of imidazole by analyzing the elution profile on a Coomassie stained SDSPAGE gel and taking into account the volumes of the tubing.

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The purification success is greater this way than via washing with a gradient, where the more suitable imidazole concentration is applied just in a very limited volume and often superimposed with the elution of the target protein. 7. The protein will be eluted in a concentrated form at the beginning; the concentration is much higher with this kind of elution than with a gradient. 8. We could show that a minimal concentration of salt, such as NaCl, is necessary for the elution of some proteins of interest. However, some contaminant proteins are eluted without NaCl. Therefore, an additional purification step can be designed on this basis. 9. If the purity after this step is not sufficient, the selectivity for the annealed substrate can be optimized by running a self-made gravity flow Sephacryl S-400 column with a height of approximately 6 cm in an empty PD-10 column with 10 mM Tris–HCl, pH 7.5, 100 mM NaCl buffer. 10. It is possible to use the prepared substrate for a longer period of time if you adjust the volume needed. In this respect it is not important to always use the same amount of radioactivity but the same amount of substrate, which includes both labeled and unlabeled substrate molecules. Over time the concentration of the labeled substrate will decrease following the exponential law with a half life of 14.3 days, but the concentration of unlabeled substrate will not (assuming that the radioactive decay only destroys the molecule in which the decay was taking place). This has to be considered in the calculation. 11. The M13mp18 DNA will stay close to the wells, whereas the separated oligonucleotide behaves in a similar way as the bromophenol blue. The optimal separation length should provide information on the appearance of products smaller than the oligonucleotide due to nucleases possibly co-purified with the helicases. 12. The resolution of the image improves, if the gel is dried before imaging. For this purpose pre-wet two cellophane sheets in ddH2O and mantle the gel. Put filter papers on the bottom and the top and place the sandwich on a gel dryer for at least 1 h at 80C with 300 mbar. Then remove the filter papers and wrap the gel mantled with the cellophane sheets in plastic wrap. This is necessary as the gel is not completely dry after the incubation and additional unequal drying will cause the gel to form cracks. The cellophane efficiently keeps the radioactive label in the sandwich.

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Acknowledgments We thank Katharina Demand and Sandra Blanck for active involvement in the processes described. Furthermore, we thank Helena Plchova, Jasmin Duerr, Verena Geuting, and Sandra Thies for sharing their experiences on the different methods and Sabrina Hettinger and Carina Moock for skillful technical assistance. This work has been supported by the DFG with grant number Pu 137/8. References 1. Sharma S., Doherty K. M., and Brosh R. M., Jr. (2006) Mechanisms of RecQ helicases in pathways of DNA metabolism and maintenance of genomic stability. Biochem. J. 398, 319–337. 2. Bachrati C. Z. and Hickson I. D. (2003) RecQ helicases: suppressors of tumorigenesis and premature aging. Biochem. J. 374, 577–606. 3. Hanada K. and Hickson I. D. (2007) Molecular genetics of RecQ helicase disorders. Cell Mol. Life. Sci. 64, 2306–2322. 4. HicksonI.D.(2003)RecQhelicases:caretakers of the genome. Nat. Rev. Cancer 3, 169–178. 5. Opresko P. L., Cheng W. H., and Bohr V. A. (2004) Junction of RecQ helicase biochemistry and human disease. J. Biol. Chem. 279, 18099–18102. 6. Hartung F., Plchova H., and Puchta H. (2000) Molecular characterisation of RecQ homologues in Arabidopsis thaliana. Nucleic Acids Res. 28, 4275–4282. 7. Hartung F., Suer S., and Puchta H. (2007) Two closely related RecQ helicases have antagonistic roles in homologous recombination and DNA repair in Arabidopsis thaliana. Proc. Natl. Acad. Sci. U.S.A. 104, 18836–18841. 8. Popuri V., Bachrati C. Z., Muzzolini L., et al. (2008) The Human RecQ helicases, BLM and RECQ1, display distinct DNA substrate specificities. J. Biol. Chem. 283, 17766–17776. 9. Kobbe D., Blanck S., Demand K., Focke M., andPuchtaH.(2008)AtRECQ2,aRecQ-helicase homologue from Arabidopsis thaliana, is able to disrupt different recombinogenic DNA-structuresinvitro.PlantJ.55,397–405. 10. Tuteja N. and Tuteja R. (2004) Unraveling DNA helicases. Motif, structure, mechanism andfunction.Eur.J.Biochem.271,1849–1863. 11. Brosh R. M., Jr., Orren D. K., Nehlin J. O., et al. (1999) Functional and physical interaction between WRN helicase and human

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replication protein A. J. Biol. Chem. 274, 18341–18350. Studier F. W. and Moffatt B. A. (1986) Use of bacteriophage T7 RNA polymerase to direct selective high-level expression of cloned genes. J. Mol. Biol. 189, 113–130. Stofko-Hahn R. E., Carr D. W., and Scott J. D. (1992) A single step purification for recombinant proteins. Characterization of a microtubule associated protein (MAP 2) fragment which associates with the type II cAMP-dependent protein kinase. FEBS Lett. 302, 274–278. Chaga G. S. (2001) Twenty-five years of immobilized metal ion affinity chromatography: past, present and future. J. Biochem. Biophys. Methods 49, 313–334. Brosh R. M., Jr., Opresko P. L., and Bohr V. A. (2006) Enzymatic mechanism of the WRN helicase/nuclease. Methods Enzymol. 409, 52–85. Bachrati C. Z. and Hickson I. D. (2006) Analysis of the DNA unwinding activity of RecQ family helicases. Methods Enzymol. 409, 86–100. Sambrook J. and Russell D. W. (2001) Molecular Cloning: A Laboratory Manual. 2nd ed. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory. Terpe K. (2006) Overview of bacterial expression systems for heterologous protein production: from molecular and biochemical fundamentals to commercial systems. Appl. Microbiol. Biotechnol. 72, 211–222. Grossman T. H., Kawasaki E. S., Punreddy S. R., and Osburne M. S. (1998) Spontaneous cAMP-dependent derepression of gene expression in stationary phase plays a role in recombinant expression instability. Gene 209, 95–103.

Chapter 15 Fluorometric Assay of Hepatitis C Virus NS3 Helicase Activity Mariusz Krawczyk, Anna Stankiewicz-Drogon´, Anne-Lise Haenni, and Anna Boguszewska-Chachulska Abstract The development of techniques based on fluorescence has made it possible to create new types of assays that represent an advantageous alternative to old tests relying on radioactivity. Such a novel approach has been applied to develop a high-throughput assay to measure the helicase activity of the hepatitis C virus (HCV) NS3 protein and the inhibitory potential of several classes of compounds. The NS3 helicase is one of the most promising targets of anti-HCV-oriented screening of compounds due to the urgent need for more effective and tolerable drugs. The 96- or 384-well microplate assay that we developed is based on the use of a quenched double-stranded DNA substrate labeled with a fluorophore (Cy3 or FAM) and with a Black Hole Quencher 1 or 2. It allows for direct (real-time) measurements of substrate unwinding and inhibition of unwinding by anti-helicase compounds. After a few modifications of buffers and assay conditions this method can be applied to various variants of HCV helicase and other proteins with helicase activities. Key words: Helicase, hepatitis C virus, NS3 protein, fluorometric assay, inhibitor screening.

1. Introduction Amplification of viral genomes, composed of either DNA or RNA strands, requires unwinding of base-paired structures that precede the synthesis of complementary strands. This phenomenon calls for specific enzymes, helicases, and energy supplied in the form of an NTP. In the case of RNA viruses, these enzymes are necessary to disrupt long as well as short secondary structures corresponding to RNA–RNA interactions, including full-length copies of genomes formed during RNA replication.

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The genome of RNA viruses codes for a helicase that together with the virus-encoded RNA-dependent RNA polymerase direct replication of the genome. These two proteins, which are the key enzymes for viral replication, are considered potential targets for anti-viral therapy. Due to the world-wide epidemiological threat and lack of fully efficient treatment or vaccine against hepatitis C virus (HCV), a monopartite single-stranded (þ) RNA virus of the Flaviviridae family (1, 2), its helicase has become an object of numerous studies that aim at hindering or abolishing its activity thereby preventing viral replication (3). A major advantage of this approach is that this protein has no close homologs among cellular enzymes (4). Various methods have been used to determine the activity of the HCV helicase encoded by the C-terminal part of the NS3 region of the viral polyprotein. The most widely applied method has been a radioactive test, adapted for the NS3 helicase by Kim et al. (5) and since then used with some modifications by other groups (6–8). The helicase moiety forms a well-defined domain (separated from the NS3 protease domain by a flexible linker) whose activity can be studied separately (5). This facilitates expression and purification of the enzyme for biochemical studies. NS3 has a broad substrate specificity; however, it unwinds doublestranded (ds) DNA more efficiently than dsRNA or heteroduplexes (3). To efficiently unwind dsRNA, NS3 requires the NS4A cofactor (9). Moreover, dsRNA substrates are difficult to prepare, unstable and expensive. Therefore, to develop a largescale, high-throughput screening assay for the HCV helicase that could be applied to test its inhibitors, the helicase domain has most frequently been used with a dsDNA substrate. In spite of its high sensitivity, the radioactive test has some disadvantages, especially the tedious preparation and instability of the radiolabeled substrate (10). Moreover, it is not a simple direct measurement of enzyme activity since the reaction products must be separated by gel electrophoresis and then submitted to autoradiography, scanning, and densitometry analyses. To partially overcome these problems, three types of high-throughput assays were developed, two tests involving radioactivity and signal amplification by scintillation [the scintillation proximity assay (11) and the FlashplateTM assay (12)] and one ELISA-based assay (13). These methods are still rather complicated and slow; none of them allows to directly follow the helicase activity. To develop a fast non-radioactive high-throughput assay to easily and directly assess enzyme activity, and to measure the inhibitory potential of libraries of potential HCV helicase inhibitors in a microplate format (14, 15), we made use of the F¨orster resonance energy transfer (FRET) phenomenon and of fluorescent dyes as well as of newly introduced fluorescence quenching molecules (Black Hole Quenchers, Biosearch Technologies) (16).

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In this chapter we present the full protocol used to purify the HCV helicase, measure its activity, and test its prospective inhibitors, based on the high-throughput assay that we have recently developed (14, 15). After a few modifications this method could be applied to test other proteins with helicase activities.

2. Materials 2.1. Helicase Purification

1. Express Five SFM medium (Gibco): 1 L medium is supplemented with 40 mL L-glutamine 100 , 1 mL gentamycin 50 mg/mL, and 1 mL amphotericin 0.25 mg/mL (Gibco).

2.1.1. Cell Culture and Lysis

2. Buffer A: 20 mM Tris–HCl, pH 7.5, 500 mM NaCl, 5 mM imidazole. 3. Protease inhibitor cocktail (Complete, EDTA-free, Roche): dissolve 1 tablet of the protease inhibitor cocktail in 2 mL of buffer A, aliquot (250 or 500 mL), and store at 20C.

2.1.2. Talon Affinity Purification

1. Talon metal affinity resin (Clontech). 2. Buffer B: buffer A supplemented with 10% glycerol and 0.05% CHAPS (AppliChem). 3. Buffer E: 20 mM Tris–HCl, pH 7.5, 200 mM NaCl, 200 mM imidazole. 4. Glycerol, CHAPS (AppliChem).

2.1.3. Heparin Purification

1. Low salt buffer: 100 mM NaCl, 20 mM Tris–HCl, pH 7.5; filtered and degassed prior to use. 2. High salt buffer: 1 M NaCl, 20 mM Tris–HCl, pH 7.5; filtered and degassed prior to use. 3. 1 mL heparin HP column (Amersham Biosciences). 4. OD280 buffer: 6 M guanidinium hydrochloride, 20 mM sodium phosphate buffer, pH 6.5. All buffers should be kept at 4C.

2.2. Fluorometric Assay of the Helicase Activity 2.2.1. Preparation of a Quenched Fluorescent Substrate

1. Annealing buffer: 20 mM Tris–HCl, pH 7.6. 2. Donor strand (FAM-TAGTACCGCCACCCTCAGAACCT TTTTTTTTTTTT, HPLC purified and lyophilized) (metabion) is dissolved to 200 mM concentration in 20 mM Tris– HCl, pH 7.6, 0.01% Tween 20, aliquoted (10 mL/aliquot), and stored at –80C. 3. Acceptor (quencher) strand (GGTTCTGAGGGTGGCGGT ACTA-BHQ1, HPLC purified and lyophilized) (metabion) prepared as for the donor strand, aliquoted (12 mL/aliquot), and stored at –80C.

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2.2.2. Assay Conditions

1. Reaction buffer: 30 mM Tris–HCl, pH 7.6, 6 mM MnCl2, 0.075% Triton X-100, 0.05% sodium azide (see Note 1). 2. ATP is brought to 0.25 M in 20 mM Tris–HCl and the pH is adjusted to 7.5 with a saturated solution of NaOH. Store aliquots at –20C. 3. Capture strand (TAGTACCGCCACCCTCAGAACC, Institute of Biochemistry and Biophysics) is diluted to 50 mM in water. Store aliquots at –20C. 4. Black 384-well plate (NUNC). 5. Multi-detection Microplate Reader (Synergy HT, Biotek) with excitation filter 485/20 nm and emission filter 528/20 nm (see Note 2).

3. Methods 3.1. NS3 Helicase Purification 3.1.1. Cell Culture and Lysis

1. To express the NS3 helicase, whose cloning as well as construction of the recombinant baculovirus was previously described (14, for technical details consult 17), Hf insect cells growing in suspension in Express five SFM medium are used. For passaging, the cells should reach 2–5 106 cells/mL and should be diluted to 2 105 cells/mL. A culture of 2 106 cells/mL is used for infection with the recombinant baculovirus at a MOI ¼ 2.5 (multiplicity of infection: number of virus particles per cell) (17). The infected cells are harvested 44–48 hpi (hours post infection), cooled on ice, centrifuged for 10 min at 720 g at 4C, and stored at –20C (see Note 3). 2. For lysis thaw the cells from 50 mL of culture at room temperature and suspend in 5 mL of buffer A. Add 250 mL of protease inhibitor cocktail and vortex thoroughly. Freeze in liquid nitrogen, then thaw quickly in a water bath at 37C (see Note 4), and vortex thoroughly. Repeat the freezing/ thawing procedure an additional four times. 3. Centrifuge the cell lysate twice for 15 min at 20,000 g at 4C, add glycerol to 10% and CHAPS to 0.05% to the supernatant, and proceed with this sample to the Talon affinity purification step.

3.1.2. Talon Affinity Purification

1. Equilibration of Talon affinity resin: centrifuge 500 mL of Talon affinity resin for 1 min at 720 g at room temperature. Gently remove the supernatant and add 5 mL of buffer B to wash the resin. Centrifuge for 1 min at 720 g at room temperature and discard the supernatant (see Note 5).

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2. Binding of the NS3 helicase: add the sample to the washed Talon affinity resin and incubate for 1–16 h at 4C on a rocking platform. 3. Purification of the NS3 helicase: discard the flow-through and wash the resin three times with 5 mL buffer B. Add 500 mL buffer E to the resin and elute the helicase by gently tapping the tube for 1 min. Centrifuge for 1 min at 720 g at 4C and collect the eluate. Repeat 3–5 times (see Note 6). 4. Run an SDS-12% PAGE to verify the purity and amount of NS3 helicase in each eluate, loading 10 mL of eluate into each well (Fig. 15.1, lane 1).

Fig. 15.1. Purification of HCV helicase, analyzed by SDS-PAGE and detected by Coomassie staining. Lane 1—combined eluates from the Talon column; lanes 2–4—eluates from the heparin column.

3.1.3. Heparin Purification

1. Prepare the 1 mL Heparin HP column by washing the column with 5–10 column volumes of water, then with high salt buffer, and finally with low salt buffer, at a speed of 0.5 mL/min. 2. For purification on the column use 2–3 eluates from the Talon affinity purification step of the highest concentrations based on the SDS-PAGE. A Biologic chromatography system (Bio-rad) is used for this step of protein purification. All steps should be performed at 4C and with buffers cooled to 4C. 3. Apply the eluates at a speed of 0.2 mL/min, wash the column with 5 column volumes of low salt buffer followed by 5 column volumes of high salt buffer at a speed of 0.5 mL/min. Collect fractions of 500 mL during the entire run. The NS3 helicase should be eluted in the unbound fraction, in the low salt buffer (see Note 7).

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4. Run an SDS-12% PAGE to verify the amount and purity of the NS3 helicase, loading 10 mL of eluate into each well (Fig. 15.1, lanes 2–4). 5. Collect the fraction with the highest concentration based on the SDS-PAGE, add glycerol to 20%, aliquot the protein (50–100 mL), and store at –80C. 6. Calculate the final concentration of the NS3 helicase: dilute the NS3 helicase 20–100 times in OD280 buffer to a final volume of 100 mL and measure the absorbance at 280 nm (A280). Calculate the molar concentration (CM) from the equation: CM = A280 n m/, where n is the cuvette pathlength, m is the dilution of the sample, and is the molar extinction coefficient of 51,760, calculated on the basis of protein composition by the ProtParam program from the Expasy website (http://us.expasy.org). 3.2. Fluorometric Helicase Activity Assay

The microplate assay presented below is based on the use of a quenched dsDNA substrate with a 30 single-stranded (ss)DNA tail, obtained by annealing of a 36-mer, which is 50 labeled with a fluorophore FAM, with a 22-mer that is 30 labeled with a Black Hole Quencher (BHQ) 1 (see Note 8). The helicase binds to the ssDNA tail and the unwinding reaction is started by the addition of ATP (Fig. 15.2). In this assay excess capture strand, complementary to the oligonucleotide labeled with the quencher, is added to prevent re-annealing of unwound duplex and to increase reaction efficiency.

Fig. 15.2. Scheme of the helicase reaction. The donor strand is labeled with FAM and the acceptor strand is labeled with BHQ1.

3.2.1. Preparation of the Quenched Fluorescent Substrate

1. Mix the following: 10 mL (1 aliquot) of donor strand, 12 mL (1 aliquot) of acceptor strand, and 78 mL of annealing buffer. 2. Denature for 2 min at 90C and cool slowly to room temperature. 3. Add 400 mL of annealing buffer. The final concentration of dsDNA substrate is 4 mM. 4. Store aliquots (50 or 100 mL) at –80C, in the dark.

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1. Prepare two eppendorf tubes for each reaction. One is for the reaction and the other is for the control reaction without enzyme (background). 2. Add 500 mL of reaction buffer to the reaction tube along with 1.25 mL of dsDNA substrate (final concentration: 10 nM) and 1.25 mL capture strand solution (final concentration: 125 nM) and mix well. 3. Transfer 200 mL of the mixture to the background tube. 4. Add the helicase, usually to 10 or 20 nM to the reaction tube. 5. Preincubate both tubes at room temperature for 15 min in the dark. 6. Add 2.25 mL of ATP to the reaction tube and 1.5 mL to the background tube and mix well avoiding air bubbles. Final ATP concentration is 1.5 mM (see Note 9). 7. Add 60 mL of the mixture from the reaction tube to each of four wells of a 384 well plate. Similarly add 60 mL of the mixture from the background tube, but only to two wells (see Note 10). 8. The plate is incubated for 1 h at 30C in a microplate fluorescence reader with scans every 2 min.

3.4. Evaluation of Inhibitory Potential of Compounds

1. Prepare two eppendorf tubes for each reaction. One is for the reaction and the other is for the control reaction without enzyme (background). 2. Prepare a reaction mix with 10 nM dsDNA substrate and 125 nM capture strand in the reaction buffer (for each reaction to be performed add 500 mL of reaction buffer, 1.25 mL of dsDNA substrate, and 1.25 mL capture strand solution) and mix well. 3. Transfer 450 mL of the mixture to the reaction tube. 4. Add the inhibitor to be tested (see Note 11) and mix well. 5. Transfer 150 mL of the mixture from the reaction tube to the background tube. 6. Subsequent steps are identical to steps 4–8 of Section 3.3.

3.5. Data Processing 3.5.1. Calculations of Helicase Activity

Mean background fluorescence values are calculated and subtracted from the values obtained for the helicase reactions (see Note 12). A progress curve is drawn for each reaction replica, a linear equation Y = A + BX is fitted to the experimental data, and the enzyme activity is calculated as the initial reaction velocity from the linear part of the progress curve (Fig. 15.3) using the linear regression method, where Y is the enzyme activity expressed in fluorescence units (FU), X is the reaction time, B is the slope or the initial velocity (mean of the increase in fluorescence per time unit), and A is the background level (fluorescence at the beginning of progress curve). The simplest way of calculating initial velocity is by using the ‘slope’

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Fig. 15.3. (a) Typical time courses of the helicase reaction at various helicase concentrations. (b) Effect of HCV helicase concentration on the rate of unwinding. The data are presented as the mean – standard error.

function of the Microsoft Excel program for each separate helicase reaction, followed by calculation of the mean value. Each experiment is repeated at least three times to obtain reliable data. 3.5.2. Quantification of Inhibitory Potential

Initial velocities of reactions carried out with various concentrations of inhibitors are calculated as described above. The helicase activity is expressed as percent of initial velocity of the control reaction without the inhibitor plotted against inhibitor concentration using the Origin 6.0 program (OriginLab Corporation); the most appropriate function is then fitted to the data set. The concentration of inhibitor necessary to reduce the enzyme activity to 50% (IC50) is calculated on the basis of a function fitted by the program. For example, to calculate IC50 based on the exponential

Fig. 15.4. Inhibition of the helicase activity by an inhibitor. The exponential decay function was fitted to the data that are mean of three independent assays – standard error.

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decay function (Y = A0 + Ae (–X/t)) fitted, X is calculated from the equation: X = –t (loge (Y–A0)/A), where X is IC50 and Y = 50, and values of parameters A and t are given by the program, while e corresponds to the base of the natural logarithm (Fig. 15.4).

4. Notes 1. Mn2+ in the reaction buffer oxidizes with time resulting in the buffer turning brown. Generally the reaction buffer is quite stable, but when there is an increase in fluorescence during the reaction in the control wells, a fresh buffer should be prepared. If a fresh buffer still causes an increase in fluorescence during the reaction, one of the buffer ingredients is most likely contaminated and must be replaced (usually Tris, Mg2+, or Mn2+ solution). 2. It is also possible to use other microplate fluorometers as well as cuvette fluorometers. 3. Do not freeze more cells than from 50 mL of culture in one tube; otherwise during the thawing procedure, the sample would be unevenly thawed and protein degradation could occur. 4. Thawing should take no more than 7 min to avoid a local increase of temperature that would favor protease activity and possible degradation of the NS3 helicase. 5. Do not add DTT to the buffers—it reduces Co2+ and prevents the protein from binding to the resin. The same occurs when the buffer pH is too low. In either case the resin changes color from pink to brown. 6. Avoid collecting any resin beads as this could block the heparin column in the following purification step. 7. For purification of the NS3 helicase of more than 200 mL culture, a 5 mL Heparin HP column can be used. Proceed with lysis and the Talon purification steps using proportionally more buffers. For heparin purification load the sample at a speed of 0.5 mL/min and perform other steps at a speed of 1 mL/min. Collect 1 mL fractions during the entire run. 8. A Cy3-labeled 36-mer together with a 22-mer 30 labeled with BHQ2 can be used instead of the FAM- and BHQ1-labeled pair of oligonucleotides. 9. Optimal ATP concentration as well as type of divalent metal ion (Mn2+ or Mg2+) should be determined for each helicase separately. Keep in mind that the concentration ratio of ATP: metal ions should be approximately 1:4.

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10. At every step try to avoid too many bubbles, especially while distributing samples into a microplate. If bubbles still form, try to pierce them with a needle. 11. Inhibitors are usually solubilized in DMSO, which does not appear to affect the helicase activity up to 2% concentration. 12. This is really necessary only in the presence of a compound (e.g., inhibitor) whose fluorescence changes during the reaction.

Acknowledgment This work was supported by the Ministry of Science and Higher Education grant 2P05A 038 29. References 1. Moradpour D., Penin F., and Rice C. M. (2007) Replication of hepatitis C virus. Nat. Rev. Microbiol. 5, 453–463. 2. Hayashi N. and Takehara T. (2006) Antiviral therapy for chronic hepatitis C: past, present, and future. J. Gastroenterol. 41, 17–27. 3. Frick D. N. (2007) The hepatitis C virus NS3 protein: a model RNA helicase and potential drug target. Curr. Issues Mol. Biol. 9, 1–20. 4. Lam A. M., Rypma R. S., and Frick D. N. (2004) Enhanced nucleic acid binding to ATP-bound hepatitis C virus NS3 helicase at low pH activates RNA unwinding. Nucleic Acids Res. 32, 4060–4070. 5. Kim D. W., Gwack Y., Han J. H., and Choe. J. (1995) C-terminal domain of the hepatitis C virus NS3 protein contains an RNA helicase activity. Biochem. Biophys. Res. Commun. 215, 160–166. 6. Hong Z., Ferrari E., Wright-Minogue J., Chase R., Risano C., Seelig G., Lee C. G., and Kwong A. D. (1996) Enzymatic characterization of hepatitis C virus NS3/ 4A complexes expressed in mammalian cells by using the herpes simplex virus amplicon system. J. Virol. 70, 4261–4268. 7. Gwack Y., Kim D. W., Han J. H., and Choe J. (1997) DNA helicase activity of the hepatitis C virus nonstructural protein 3. Eur. J. Biochem. 250, 47–54.

8. Borowski P., Deinert J., Schalinski S., Bretner M., Ginalski K., Kulikowski T. and Shugar D. (2003) Eur. J. Biochem. 270, 1645–1653. 9. Frick D. N., Rypma R. S., Lam A. M. and Gu B. (2004) The non-structural protein 3 protease/helicase requires an intact protease domain to unwind duplex RNA efficiently. J. Biol. Chem. 279, 1269–1280. 10. Tai C. L., Chi W. K., Chen D. S., and Hwang L. H. (1996) The helicase activity associated with hepatitis C virus nonstructural protein 3 (NS3). J. Virol. 70, 8477–8484 11. Kyono K., Miyashiro M., and Taguchi I. (1998) Detection of hepatitis C virus helicase activity using the scintillation proximity assay system. Anal. Biochem. 257, 120–126. 12. Hicham Alaoui-Ismaili M., Gervais C., Brunette S., Gouin G., Hamel M., Rando R. F., and Bedard J. (2000) A novel high throughput screening assay for HCV NS3 helicase activity. Antiviral Res. 46, 181–193. 13. Artsaenko O., Tessmann K., Sack M., Haussinger D., and Heintges T. (2003) Abrogation of hepatitis C virus NS3 helicase enzymatic activity by recombinant human antibodies. J. Gen. Virol. 84, 2323–2332. 14. Boguszewska-Chachulska A. M., Krawczyk M., Stankiewicz A., Gozdek A., Haenni A.L., and Strokovskaya L. (2004) Direct fluorometric measurement of hepatitis C virus helicase activity. FEBS Lett. 567, 253–258.

Hepatitis C Virus NS3 Helicase Activity 15. Boguszewska-Chachulska A. M., Krawczyk M., Najda A., Kopanska K., Stankiewicz-Drogon A., Zagorski-Ostoja W., and Bretner M. (2006) Searching for a new anti-HCV therapy: synthesis and properties of tropolone derivatives. Biochem. Biophys. Res. Commun. 341, 641–647.

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16. Johansson M. K., Fidder H., Dick D., and Cook R. M. (2002) Intramolecular dimers: a new strategy to fluorescence quenching in dual-labeled oligonucleotide probes. J. Am. Chem. Soc. 124, 6950–6956. 17. King L. A. and Possee R. D. (1992) The Baculovirus Expression System. A Laboratory Guide. Chapman and Hall, London.

Chapter 16 A Method to Simultaneously Monitor Hepatitis C Virus NS3 Helicase and Protease Activities David N. Frick, Olya Ginzburg, and Angela M.I. Lam Abstract The hepatitis C virus NS3 protein contains an N-terminal serine protease and a C-terminal helicase that unwinds RNA or DNA duplexes. The HCV NS3 protein is the target for several antiviral drugs in clinical trials, which inhibit the protease function. A method is reported to simultaneously monitor the helicase and protease function of the NS3 protein in a single reaction using fluorescence spectroscopy and a single chain recombinant protein where NS3 is fused to its protease activator NS4A. The method monitors both activities together in real time and is amenable to high-throughput screening. This new procedure could be used to identify compounds that inhibit both the helicase and protease activity of NS3. Key words: Helicase, protease, ATPase, high-throughput screening, antiviral agents.

1. Introduction The hepatitis C virus (HCV) causes a common liver disease affecting 2–3% of the world’s population and is frequently called a ‘‘silent’’ killer because patients show few signs of infection (1). As the disease progresses over a period of decades, when patients might unknowingly transmit the blood-borne positive sense RNA virus to others, hepatitis C patients frequently develop fibrosis, cirrhosis, or liver cancer. At this late stage of the disease, a liver transplant is the only option for survival, and as a result, HCV infection is presently the most common cause for liver transplantation in many parts of the world. Current HCV therapies, which combine pegylated interferon and ribavirin, are quite effective for patients infected with certain HCV strains, but they are costly and produce debilitating side effects that are usually worse than the M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_16, ª Humana Press, a part of Springer Science+Business Media, LLC 2010

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symptoms caused by the virus itself. In addition, patients infected with HCV genotypes most common in North America, patients with advanced disease, transplant recipients, and patients co-infected with HIV frequently do not respond to current therapies. Compounds that inhibit HCV-encoded enzymes are being developed as new antivirals to treat HCV infection. Two of the leading candidates in clinical trials are the protease inhibitors telaprevir (VX-950, Vertex) and boceprevir (SCH 503034, ScheringPlough). Telaprevir and boceprevir inhibit a protease formed from two of ten proteins encoded by the 9,600 nucleotide long HCV RNA genome—nonstructural protein 3 (NS3) and nonstructural protein 4A (NS4A). The HCV genome encodes one main open reading frame, which is translated into a single 3,000 amino acid long polyprotein. The NS3/NS4A protease, one other viral protease, and cellular proteins process the polyprotein into the mature peptides needed for replication. The catalytic triad of this serine protease resides near the N-terminus of NS3, and NS4A activates the enzyme by binding NS3. In addition to being a protease, NS3 is also an ATP-fueled helicase capable of separating duplex RNA or DNA. The two catalytic activities can be expressed separately as recombinant proteins containing truncated NS3 fragments, with the protease residing in the N-terminal NS3 fragment and the helicase residing in the C-terminal fragment. Mutants lacking helicase function still cleave the polyprotein but the virus is no longer viable (2). There are three lines of evidence that the two activities are linked. First, without the protease, NS3 unwinds RNA less efficiently. Second, RNA binds more tightly to NS3 when the protease is present possibly because the protease adds positively charged surfaces that may stabilize RNA (3). Third, mutations on the helicase domain alter binding of various protease inhibitors (4). It is therefore reasonable to believe that small molecules can be discovered that simultaneously inhibit both NS3 helicase and protease activities, and as proof of this concept, some RNA aptamers have already been designed with such properties (5). To aid in the discovery and analysis of dual NS3 helicase/ protease inhibitors, we have developed a method to simultaneously monitor both peptide cleavage and DNA unwinding catalyzed by NS3 in a single reaction. The simultaneous measurement of NS3 protease and helicase is conceptually simple because continuous fluorescence-based assays have been developed to separately measure either NS3 protease (6) or NS3 helicase (7). Combining these assays is technically challenging for two reasons. First, purification of the full-length NS3 protein in combination with NS4A is difficult because of their highly insoluble nature; in cells NS4A transverses membranes. Second, helicase-catalyzed unwinding is only apparent at low ionic strength, and NS3

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protease assays must be performed in high salt concentrations because NS4A binds NS3 through hydrophobic interactions. The use of a protein in which a His-tagged truncated NS4A peptide is covalently fused with full-length NS3 (8) essentially solved these problems. This single chain complex, called here ‘‘scNS34A,’’ differs from the native complex because NS4A, which is normally attached via a cleavable linker to the NS3 C-terminus, is instead attached to the N-terminus of NS3 with a stable peptide. We have found that under conditions where NS3 helicase is most active (without salt at slightly acidic pH), the protease of scNS34A can be monitored if glycerol, dithiothreitol (DTT), and the detergent b-octyl glucoside (b-OG) are added to the reactions. These components do not drastically alter detection sensitivity in helicase assays. In the assay below, both the helicase and protease substrates are designed to directly mimic parts of the HCV genome (Fig. 16.1a). The depsipeptide used to measure protease is based on the junction between the NS4A and NS4B proteins, and the molecular beacon helicase substrate is based on a stem loop found at the end of NS5B (Fig. 16.1a). While the protease substrate is widely used and now part of an assay Kit (SensoLyteTM 490 HCV Protease Assay Kit, Anaspec, San Jose, CA), our new molecular beacon based helicase assay is noteworthy because it differs from other fluorescent helicase assays in three critical ways. First, all modifications are made on only one strand of the helicase substrate. Second, the assay monitors a decrease in fluorescence, which is noteworthy because data can be easily converted to percentage duplex remaining simply by calculating fractional fluorescence (F/Fo) after subtracting background fluorescence. Third, the products form hairpins, making the reaction essentially irreversible. Intramolecular hairpin formation is thermodynamically favored even at relatively high DNA concentrations. Only at DNA concentrations above 50 nM is the intermolecular reaction favored. This last point is a key advantage over helicase assays described elsewhere in this volume because the molecular beacon assay does not require high concentrations of substrate traps, which could influence observed reaction rates. The simultaneous assay can be performed either in cuvettes or in 96- or 384-well microplates. Numerous potent specific inhibitors for NS3 protease are commercially available, like the NS3 product-based inhibitor Ac-Asp-D-Gla-Leu-Ile-b-cyclohexyl-Ala-cys-OH (Bachem, Torrance, CA), which gives an IC50 of 8.9 nM in this assay. However, only a few HCV helicase inhibitors are available at this time. Typical non-hydrolyzable nucleotide analogs are weak binders, and only a few nucleotide-like compounds display any potent inhibitory activity. Examples with IC50s in the micromolar range include a nucleoside analog with a diaminodihydrotriazine substituent replacing the 5-amino group of the

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Fig. 16.1. The simultaneous HCV protease/helicase assay. (a) Schematic diagram of the assay. The protease assay (6) and the helicase assay (7) have been previously reported. (b) Assay performed as described in Section 3. At the indicated times, protease substrate (P. S., 2 mM final concentration), scNS3-4A (10 nM) and ATP (1 mM) were added to the solution. The reaction was monitored at the excitation wavelength of 355 nm and the emission wavelength of 485 nm (protease substrate) and at excitation 550 nm and emission 570 nm (helicase substrate) using a Varian Cary Eclipse Fluorescence spectrophotometer. (c) Initial rates (arbitrary units/minute) of protease reactions as a function of scNS3-4A concentration. (d) Initial rates of helicase reactions as a function of scNS3-4A concentration.

nucleoside metabolite 5-aminoimidazole-4-carboxamide-1-bD-ribofuranoside (9), a nucleotide mimic called QU663 (10), 4,5,6,7-tetrabromobenzotriazole (TBBT), 5,6-dichloro-1(b-D-ribofuranosyl)benzotriazole (DRBT) (11), and a short peptide (HCVpep) that mimics a conserved helicase motif

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(12). Of these compounds, only NS3pep (12), DRBT, and TBBT (13) have been confirmed to have antiviral activity in assays using HCV replicons. We have used the assay below to evaluate the potency of many of these available HCV helicase inhibitors (7), but we have yet to find any inhibitors that bind more tightly than the helicase substrate, which binds with nanomolar affinity. Compared with various known inhibitors, the oligonucleotide, dT20, a representative compound that binds in place of the RNA or DNA substrate, is still the most potent inhibitor we know with an IC50 of 30 nM. Only NS3pep (12) is comparable with an IC50 of 100 nM. By comparison, non-hydrolyzable ATP analogs bind much weaker. For example, the IC50 of ,b-methylene-ATP is 140 mM, and b,g-imido-ATP binds even weaker with an IC50 of 1.8 mM (7). It is tempting to speculate that compounds could be found that influence both the helicase and protease activities of NS3, and this assay was designed to help discover such compounds. The many atomic structures available for NS3, in particular, the structure of scNS3-4A (14) and the structure of HCV helicase bound to an oligonucleotide (15) support this contention. In the full-length NS3-NS4A complex, two domains compose the protease core and three other domains form the helicase core. If the helicase is viewed like a ‘‘Y,’’ one strand of RNA binds below two RecA-like domains, and ATP likely binds between the two upper RecA-like domains. The protease domains pack with the helicase domains to bury the catalytic triad in a cleft formed behind the ATP and DNA-binding clefts. Based on electrostatic analyses of NS3 structures, our laboratory has suggested that in addition to binding below the RecA-like domains, RNA might also bind in the cleft separating the helicase from the protease. If RNA binds near the protease active site, then compounds binding to this region could affect both the helicase and protease activities.

2. Materials 2.1. Cloning scNS3-4A

1. A recombinant DNA clone of HCV containing the entire NS3 region. The primers below have been used successfully with HCV genotype 1b strains J4 and con1. Primer sequences may need to be adjusted if a distantly related HCV genotype is used as the source of the NS3 protein (see Note 1). 2. Plasmid vector pET28b (Novagen, Madison, WI).

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3. An upstream PCR primer, which encodes an NdeI restriction enzyme site, NS4A residues 21–32, and a gly-ser-gly-ser linker. 50 -GC GAT ATA CAT ATG GGT TCT GTT GTT ATT GTT GGT AGA ATT ATT TTA TCT GGT AGT GGT AGT ATC ACG GCC TAC TCC CAA-30 . 4. A downstream primer, which contains an EcoRI site. (50 -GCG CGC GAA TTC GGT CAA GTG ACG ACC TCC AGG TCA GCC GAC ATG C-30 ). 5. Molecular biology reagents including Nde1 and EcoR1 (New England Biolabs, Ipswich, MA), PCR reagents, thermocycler, DNA agarose gel electrophoresis apparatus, gel extraction kit (Qiagen), and a DNA ligation kit (Novagen). 2.2. Purification of scNS3-4A

1. Buffer A: 25 mM HEPES, 500 mM NaCl, 10 mM b-mercaptoethanol, 20% Glycerol, pH 8. 2. Sonifier cell disruptor (Branson, Danbury, CT). 3. Pre-charged nickel-nitrilotriacetic acid (NiNTA) resin preequilibrated with buffer A. 4. An 200 ml Sephacryl S-300 HR column (GE Healthcare Bio-Sciences, Piscataway, NJ) pre-equilibrated with buffer A. 5. Polyacrylamide gel electrophoresis apparatus.

2.3. The Simultaneous Helicase/Protease Assay

1. 2X reaction buffer: 50 mM MOPS, pH 6.5, 3 mM MgCl2, 60% glycerol, 20 mM DTT, 0.4% b-octyl glucoside, prepare fresh. 2. HCV Protease substrate (Anaspec, San Jose, CA). 3. Helicase substrate short strand: 50 -Cy3-GCTCC CCAAT CGATG AACGG GGAGC-IBQ-30 . IBQ is Iowa Black quencher from Integrated DNA Technologies (Coralville, IA). 4. Helicase substrate long strand 50 -GCTCC CCGTT CATCG ATTGG GGAGC TTTTT TTTTT TTTTT TTTTT-20 .

3. Methods 3.1. Cloning and Expression of scNS3-4A

1. Set up and perform four 25 ml PCRs using the HCV primers and templates using standard protocols (see Note 1). 2. Combine PCRs, add 20 ml of 6X agarose gel loading buffer, and load into one large well of a 1% DNA agarose gel. After electrophoresis, visualize DNA with ethidium bromide staining, excise the 1,964-bp band, and extract DNA from the gel with the Qiagen Kit.

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3. Digest purified amplicon with NdeI and EcoRI and in a second tube digest 2 mg of pET28. After 2 h at 37C, purify digested DNA again using gel electrophoresis. 4. Ligate purified vector and insert using DNA ligase, and transform competent cells that lack the T7 RNA polymerase (e.g., Novablue or BL21). 5. Purify DNA from transformed colonies and confirm the presence of the inserted DNA using restriction enzyme analysis. Confirm the sequence of clones bearing the insert using DNA sequencing. 6. After sequence verification, use the resulting plasmid (p28scNS3-4A) to transform a strain carrying a functional T7 polymerase (e.g., BL21(DE3)). 7. Pick a single kanamycin resistant colony from a fresh plate and inoculate 2 ml of LB media with 50 mg/ml kanamycin. 8. When the 2 ml culture is slightly turbid (OD600 0.1), transfer to a 4 l flask containing 1 l of LB-Kan. Incubate with vigorous shaking at 37C. Monitor OD600. 9. When OD600 reaches 1.0, transfer the flask to a room temperature (23C) shaker. 10. Add 5 ml of 200 mM IPTG, and continue shaking for 3 h at room temperature. Harvest cells using centrifugation. Store pellet at –80C. 3.2. Purification of scNS3-4A

1. Thaw frozen cells and suspend in 23.9 ml of buffer A. Add 0.25 ml of 10% b-OG (0.1% final concentration) and 1.0 ml of 500 mM imidazole (20 mM final concentration). All subsequent purification steps should be performed at 4C. 2. Sonicate cell suspension 30 s, rest 1 min, and repeat twice. 3. Spin at 14,000 rpm for 15 min and filter supernatant through glass fiber pre-filter. 4. Take 1 ml of NiNTA beads, place in 25 ml column, and wash with 10 ml of buffer A. 5. Suspend beads in 1 ml of buffer A and transfer to the crude extract. Mix gently for 30 min at 4C and pour solution into a small column (25 ml). Discard the flow-through. 6. Wash NiNTA column with 10 ml of buffer A supplemented with an additional 0.5 M NaCl. 7. Wash NiNTA column with 10 ml of buffer A supplemented with 40 mM imidazole. 8. Elute scNS3-4A with 1.5 ml of buffer A supplemented with 500 mM imidazole (see Note 2). Load eluent on Sephacryl S-300 HR column. Wash column with buffer A at 0.2 ml/min.

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9. Collect 4 ml fractions, while monitoring absorbance at 280 nm. 10. Analyze 10 ml aliquots of peak A280 fractions using 10% SDS PAGE (see Note 3). 11. Combine fractions containing the 71 kDa scNS3-4A protein (see Note 4). 12. Dialyze into 50 mM Tris, 0.1 M NaCl, 1 mM EDTA, 30% Glycerol, 0.1 mM DTT (pH 7.4). Calculate concentration from A280 of using an extinction coefficient of 68.4 mM–1 cm–1 (see Note 5). Concentrate if desired (see Note 6) and store aliquots at –80C. 3.3. The Simultaneous Helicase/Protease Assay

1. Prepare helicase substrate by diluting molecular beacon and long oligonucleotide to 20 mM in 20 mM TrisCl, pH 7. Heat to 95C, and cool slowly to room temperature. Although not absolutely necessary, the best results are obtained if the annealed substrate is purified from the free oligonucleotides using non-denaturing polyacrylamide gel electrophoresis. The free Cy3 molecular beacon and annealed helicase substrate are clearly visible without staining. 2. Reactions can be performed in microplates, or cuvettes at temperatures up to 45C. For a 100-ml reaction, add appropriate volumes of diluted helicase and protease substrate to 50 ml of 2X reaction buffer and water such that the final concentration of helicase substrate is 5 nM and the protease substrate is 2 mM. 3. Initiate protease reactions were by adding scNS3-4A protein (1–250 nM). 4. Initiate helicase reactions by adding ATP to a final concentration of 1 mM. 5. Continuously monitor fluorescence using a fluorescence spectrophotometer at the excitation wavelength of 355 nm and the emission wavelength of 485 nm (protease substrate) and at excitation 550 nm and emission 570 nm (helicase substrate) (Fig. 16.1). 6. Inhibitors of NS3 protease are commercially available and can be used as positive controls in high-throughput screens. For the helicase, simple oligonucleotides (e.g., dT20: 50 -TTTTT TTTTT TTTTT TTTTT TTTTT-30 ) are potent inhibitors with IC50 values in the nanomolar range (see Note 7).

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4. Notes 1. Upstream and downstream primers may need to be adjusted if they do not amplify DNA from the desired HCV isolate. Use appropriate computer software to align the primer to the desired sequence. If the 30 ends of the primer sequences do not completely match the desired HCV sequence, adjust them accordingly. The desired NS3 sequence should also be checked for internal NdeI and EcoRI sites, and if they are present other restriction sites should be incorporated into primers. An alternative upstream site is NheI, which is immediately downstream from the NdeI site of pET28. Alternate downstream sites are BamHI, SacI, SalI, HindIII, EagI, NotI, and XhoI. 2. Some investigators regularly use NS3 proteins purified in a single step using immobilize metal affinity chromatography, but in our experience such preparations are typically contaminated with RNase and other nucleases, which can confound helicase assays. We recommend assaying all stages of the purification with a commercial RNAse assay, such as RNaseAlert1 Kit (Ambion, Austin, TX). 3. Further purification of scNS3-4A might be necessary, but is challenging because the protein tends to aggregate in the absence of high salt. Affinity chromatography using poly(U)agarose (Sigma, St. Louis, MO) is recommended. 4. The His-tag can be removed after purification if desired using Thrombin. However, we have not yet found any impact of the His-tag in any assays. 5. The extinction coefficient was calculated based on the sequence of the scNS3-4A protein from the HCV genotype 1b(con1) strain using the program Sequence Analysis (http://informagen.com/SA/). Adjust if another HCV strain is used as the source. 6. Care should be taken with concentration because the protein tends to aggregate. Ultrafiltration and centrifugal concentrators are not recommended. We typically concentrate the protein by surrounding a dialysis bag with high molecular weight polyethylene glycol until a desired amount of liquid is removed. 7. Many compounds in libraries quench one of the two fluorophore used here. To assay such compounds other fluorescently labeled protease substrates are commercially available

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(Anaspec). Similarly a different molecular beacon helicase substrate in which Cy3 is substituted with a different fluorophore could be employed.

Acknowledgments This study was supported by a grant from the National Institutes of Health (AI052395). References 1. McHutchison J. G. (2004) Understanding hepatitis C. Am. J. Manag. Care. 10, S21–9. 2. Lam A. M. and Frick D. N. (2006) Hepatitis C virus subgenomic replicon requires an active NS3 RNA helicase. J. Virol. 80, 404–411. 3. Frick D. N. (2007) The hepatitis C virus NS3 protein: a model RNA helicase and potential drug target. Curr. Issues Mol. Biol. 9, 1–20. 4. Dahl G., Sandstrom A., Akerblom E., and Danielson U. H. (2007) Effects on protease inhibition by modifying of helicase residues in hepatitis C virus nonstructural protein 3. FEBS J. 274, 5979–5986. 5. Umehara T., Fukuda K., Nishikawa F., Kohara M., Hasegawa T., and Nishikawa S. (2005) Rational design of dual-functional aptamers that inhibit the protease and helicase activities of HCV NS3. J. Biochem. (Tokyo) 137, 339–347. 6. Taliani M., Bianchi E., Narjes F., Fossatelli M., Urbani A., Steinkuhler C., De Francesco R., and Pessi A. (1996) A continuous assay of hepatitis C virus protease based on resonance energy transfer depsipeptide substrates. Anal. Biochem. 240, 60–67. 7. Belon C. A., and Frick D. N. (2008) Monitoring helicase activity with molecular beacons. BioTechniques 45, 433–440. 8. Howe A. Y., Chase R., Taremi S. S., Risano C., Beyer B., Malcolm B., and Lau J. Y. (1999) A novel recombinant single-chain hepatitis C virus NS3-NS4A protein with improved helicase activity. Protein Sci. 8, 1332–1341. 9. Ujjinamatada R. K., Baier A., Borowski P., and Hosmane R. S. (2007) An analogue of AICAR with dual inhibitory activity against WNV and HCV NTPase/helicase: synthesis

10.

11.

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and in vitro screening of 4-carbamoyl-5(4,6-diamino-2,5-dihydro-1,3,5-triazin2-yl)imidazole-1-beta -D-ribofuranoside. Bioorg. Med. Chem. Lett. 17, 2285–2288. Maga G., Gemma S., Fattorusso C., Locatelli G. A., Butini S., Persico M., Kukreja G., Romano M. P., Chiasserini L., Savini L., Novellino E., Nacci V., Spadari S., and Campiani G. (2005) Specific targeting of hepatitis C virus NS3 RNA helicase. discovery of the potent and selective competitive nucleotidemimicking inhibitor QU663. Biochemistry 44, 9637–9644. Borowski P., Deinert J., Schalinski S., Bretner M., Ginalski K., Kulikowski T., and Shugar D. (2003) Halogenated benzimidazoles and benzotriazoles as inhibitors of the NTPase/ helicase activities of hepatitis C and related viruses. Eur. J. Biochem. 270, 1645–1653. Gozdek A., Zhukov I., Polkowska A., Poznanski J., Stankiewicz-Drogon A., Pawlowicz J. M., Zagorski-Ostoja W., Borowski P., and BoguszewskaChachulska A. M. (2008) NS3 peptide, a novel potent Hepatitis C virus NS3 helicase inhibitor, its mechanism of action and antiviral activity in the replicon system. Antimicrob. Agents Chemother. 52, 393–401. Paeshuyse J., Vliegen I., Coelmont L., Leyssen P., Tabarrini O., Herdewijn P., Mittendorfer H., Easmon J., Cecchetti V., Bartenschlager R., Puerstinger G., and Neyts J. (2008) Comparative in vitro antihepatitis C virus activities of a selected series of polymerase, protease, and helicase inhibitors. Antimicrob. Agents Chemother. 52, 3433–3437. Yao N., Reichert P., Taremi S. S., Prosise W. W., and Weber P. C. (1999) Molecular

Simultaneous NS3 Helicase/Protease Assay views of viral polyprotein processing revealed by the crystal structure of the hepatitis C virus bifunctional protease-helicase. Struct. Fold. Des. 7, 1353–1363. 15. Kim J. L., Morgenstern K. A., Griffith J. P., Dwyer M. D., Thomson J. A., Murcko M. A.,

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Chapter 17 Computer Modeling of Helicases Using Elastic Network Model Wenjun Zheng Abstract In this chapter, we will demonstrate the usage of a suite of computational techniques based on a coarsegrained elastic network model by applying them to a monomeric helicase—the NS3 helicase of hepatitis C virus. These techniques allow us to predict and visualize collective domain motions encoded in the protein structures, probe allosteric couplings between key functional sites (such as ATP-binding site and DNA/ RNA-binding site of a helicase), and simulate ATP-binding-induced global conformational changes. These general techniques are not only applicable to NS3 but also to other multi-domain protein structures. Key words: Correlation, elastic network model, helicase, normal mode analysis, NS3.

1. Introduction Helicases are a variety of enzymes that can translocate along singlestranded DNA/RNA and unwind double-stranded DNA/RNA in an ATP-dependent reaction. They are required for gene replication, transcription, translation, recombination, and repair (1–3). An increasing number of crystal structures have been solved for various helicases, which form the basis of structure-based modeling of the conformational dynamics underlying helicase functions. However, direct all-atom molecular dynamics simulations (4) are in practice limited to nanoseconds while the biologically relevant time scales are much longer (milliseconds to seconds). Coarsegrained modeling (5) based on simplified structural representations and energy functions offers a practical recipe for modeling the ‘long-time’ conformational dynamics of helicase structures.

M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_17, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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In this chapter, we will demonstrate the application of a suite of computational techniques based on a coarse-grained elastic network model to a monomeric helicase—the NS3 helicase of hepatitis C virus (6). These techniques allow us to predict and visualize collective domain motions encoded in the protein structures (7), probe allosteric couplings between key functional sites (such as ATP-binding site and DNA/RNA-binding site of a helicase) (7, 8), and simulate ATP-binding-induced global conformational changes (7, 9). We will focus on the technical details of the methodology and refer the readers to our original paper (7) for more biological discussions. These general techniques are applicable not only to NS3 helicase but also to other multi-domain protein structures. They have been made available through a web-server (enm.lobos.nih.gov).

2. Methods The modeling procedures consist of the following four components: 2.1. Normal Mode Analysis of Elastic Network Model

Given the C atomic coordinates of a protein crystal structure from Protein Data Bank (for example, an NS3 helicase structure with the PDB code 1A1V, see Note 1), we build an elastic network model (10–12) by connecting all pairs of C atoms that are within a cutoff distance (RC ¼ 10 A˚) using harmonic springs with a uniform force constant C (13). C can be determined by fitting the crystallographic B factors, although the fitting is not necessary for the techniques presented here. The total elastic energy is 1 X EENM ¼ C ðdij dij0 Þ2 ; [1] 2 05 dij

Rc

where dij is the distance between the C atoms i and j, and dij0 is the distance between C atoms i and j as given in the crystal structure. We expand the above potential energy function to second order: X 1 1 EENM X T H X ¼ C X T Hij X ; [2] 2 2 05 dij

Rc

where X ¼ X X0 , X is a 3N-dimensional vector representing the Cartesian coordinates of the N C atoms, X0 gives P the equilibrium C coordinates in the crystal structure, H ¼ C d 0 5Rc Hij is the ij Hessian matrixh (second derivatives of potential energy), where i Hij ¼ ð1=2Þr2 ðdij dij0 Þ2 .

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For the Hessian matrix H, we perform the standard normal mode analysis (NMA) to solve 3N normal modes. Each normal mode m has an eigenvalue lm and an eigenvector Vmwhich satisfyHVm ¼ lm Vm . The lowest 6 zero modes correspond to three translations and three rotations and are therefore removed from the mode spectrum (mode numbering starts from #1 for the lowest non-zero mode). The eigenvector of each mode can be compared with an observed conformational change by calculating the overlap between them, which is the cross-correlation coefficient between these two 3N-dimensional vectors. The eigenvectors of the lowest few normal modes have been found to compare well (with high overlap values) with the protein conformational changes observed crystallographically (see Note 1) (12, 14). 2.2. Dynamical Domain Partition for a Normal Mode

To visualize the collective conformational changes described by a low-frequency mode, we partition the multi-domain protein structure into dynamical domains which move (rotate and translate) as rigid bodies. We use the following procedure: 1. We deform the crystal structure along the direction of the eigenvector of a give mode by RMSD ¼ 1 A˚ to generate the end conformation. 2. We partition the protein into five-residue-long peptides. For each peptide i, we rotate and translate its initial conformation (given by the crystal structure) to fit its end conformation with minimal RMSD, and then we record the corresponding rotation matrix Ui and translation vector Ti. 3. We cluster all (Ui, Ti) using the following iterative K-means algorithm (ten iterations): a. Initial centroids are assigned to the peptides containing residues at the center of each structural domain of a multi-domain protein; b. Each peptide i is assigned to a centroid I whose (UI, TI), if applied to the initial conformation of peptide i, gives the minimal RMSD for fitting the end conformation of peptide i; those peptides assigned to the same centroid form a cluster; c. The centroid I of each cluster is re-selected from its member peptides such that its (UI, TI) minimizes the RMSD of fitting the whole cluster to the end conformation. 4. Finally we sort all clusters in decreasing size, and each cluster with at least 20 residues (or four peptides) is named a dynamical domain, and the (UI, TI) of its centroid describes the rigid-body motion of this domain. See Note 2 for an example.

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2.3. Fluctuation-Based Correlation Analysis of Hot-Spot Residues

The correlation analysis aims to identify key residues strongly coupled to the fluctuations of a subset of residues at a functional site (such as the ATP-binding site or the DNA/RNA-binding site of a helicase). Following a previous work (8), we compute the overall meansquare fluctuation (MSF) for the given subset (S) of residues as follows: D E X jPS Vm; j2 2 1 S ; [3] FS ¼ jPS~ rS j / TrðPS ðHSS ÞPS Þ ¼ lm 1mM where ~ rS is the structural displacement of the residues in subset S away from the coordinates given in the crystal structure; Ps is the projection operator that projects out the six translational and rotational components from ~ rS (because we are only interested in 1 is the S the internal fluctuations of the subset residues); HSS submatrix of the inverse of the ENM Hessian matrix 1 HSS

T X Vm;S Vm;S ¼ ; lm 1mM

lm and Vm;S are the eigenvalue and the S component of eigenvector of mode m. A cutoff at mode M ¼ 3 N/10 is used to compute 1 HSS (N is the total number of residues). Next we introduce a residue-position-specific perturbation to explore the resulting change of FS. As described in a previous paper (8),weintroducethefollowingtranslationallyandrotationallyinvariant perturbation at residue position i (it perturbs the force constant of the ˚ ): springs connecting the C atom i to its neighbors within RC ¼ 10 A X ðdik d 0 Þ2 ik Ei ¼ C ; [4] 2 05 k:dik

Rc

from which the corresponding Hessian matrix Hi can be calculated as follows: X Hi ¼ C Hik ; [5] 0 5R k:dik c

h i where Hik ¼ ð1=2Þr2 ðdik dik0 Þ2 :

Then the change of FS in response to Hi is FSji / Tr½PS ðH 1 ÞSS PS ¼ Tr½PS ðH 1 H i H 1 ÞSS PS :

[6]

Finally we can define the following correlation function for position i: FSji CFi ¼ : [7] C FS CFi measures the relevance (or importance) of residue i to the MSF of residues in subset S.

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We can identify the ‘hot-spot residues’ that are strongly coupled to the fluctuations of residues in subset S—all residues are sorted in the order of declining CFi and the top 10% of them are predicted as hot-spot residues. The hot-spot residues are dynamically important, because their local interactions with neighboring residues control the stability and dynamics of the subset S residues. Namely, by perturbing the elastic interactions involving a hot-spot residue, the MSF of subset S will be changed significantly. Therefore, mutations of a hot-spot residue will significantly alter the dynamical fluctuations of residues in subset S and thereby affect its biological activity (such as ligand binding). See Note 3 for an example. 2.4. Deformation Analysis of ATPBinding-Induced Conformational Change

The deformation analysis aims to predict the ATP-binding-induced global conformational change given the input of a local structural change at ATP-binding site. It can be applied to the modeling of other ligand-binding-induced conformational changes. To simulate the ATP-binding-induced local structural change at the ATP-binding site, we consider two local conformations of ATP-binding site (7)—the open conformation that captures the empty state, and the closed conformation that captures the ATPbound state. To structurally deform the ATP-binding site from the open to closed conformation, we superimpose the closed conformation to the open conformation (using C atoms only) to minimize the RMSD between them. Then we extract the structural displacement vector~ rS for the subset of ATP-binding residues. Finally we compute the induced structural displacement ~ rE of the rest of the ENM (named environment or E component) in response to the given local deformation ~ rS (9): 1 ~ HES~ rE ¼ HEE rS ;

[8]

where HEE and HES are two sub-matrices of the ENM Hessian matrix " # HSS HSE H ¼ HES HEE Together, ~ rS and ~ rE predict the ATP-binding-induced global conformational change. See Note 4 for an example.

3. Notes Here we will illustrate the usage of the above methods using NS3 helicase as an example.

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1. For an NS3 crystal structure (PDB code: 1A1V), we have constructed elastic network model and then performed normal mode analysis (7). It is found that the lowest two modes (#1 and #2, see Fig. 17.1) capture most of the observed conformational changes from 1A1V to several other NS3 structures (7), which validates the use of low-frequency modes to describe domain motions in NS3 helicase.

Fig. 17.1. Dynamical domain partition of the conformational changes described by mode #1 (a), mode #2 (b), and the ATP-binding-induced structural change (c). In all cases, there are two rotations of domain 1 (white), domain 2 (silver) with respect to domain 3 (black). The rotational axis is shown as an arrow—the color of the arrow stem is the same as the fixed domain (domain 3), while the color of the arrow head is the same as the moving domain (domain 1 or domain 2). The bound DNA is shown as spheres in (c).

2. The dynamical domain partition has been applied to the lowest two modes (#1 and #2) solved for an NS3 helicase structure (7) (see Fig. 17.1). For NS3 helicase, three initial centroids are assigned to the peptides containing residues at the center of domain 1 (residue 227), domain 2 (residue 366), and domain 3 (residue 564). The following domain motions are found: Mode #1 describes a closure rotation (8) of domain 2 toward domain 3, and a small twisting motion (0.7) of domain 1 relative to domain 3 (see Fig. 17.1a).

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Mode #2 describes two twisting motions of domain 2 (9) and domain 1 (2) relative to domain 3 (see Fig. 17.1b). 3. We have used the correlation analysis to identify the hot-spot residues which are strongly coupled to the ATP-binding site and DNA/RNA-binding site in NS3 helicase (7) (see Table 17.1.):

Table 17.1 Hot-spot residues for two subsets of residues—ATP-binding site and DNA/RNAbinding site in NS3 helicase Subset S

Residues of subset S

Hot-spot residues (8 clusters)

ATP-binding site

207–212 (motif I) 229–231 (motif Ia) 290–293 (motif II) 322,323 (motif III) 467 (motif VI)

1. 190,203,204,206–214 (motif I) 2. 291,293,294 (motif II) 3. 322–330 (motif III) 4. 412 (DNA/RNA binding) 5. 432 (DNA/RNA binding) 6. 450,456–463 (motif VI) 7. 479–482,484,485 8. 521

DNA/RNAbinding site

230,232,254,255,269,271,272,275,298 (domain 1) 369–371,392,393,411,413 (domain 2) 432,434,448,450 (interface: domains 2 and 3) 501,502 (interface: domains 1 and 3)

1. 293,294 (motif II) 2. 323–327,329,330 (motif III) 3. 369,370 (DNA/RNA binding) 4. 411,412,414 (DNA/RNA binding) 5. 430,432–436 (DNA/RNA binding) 6. 446,448–452,456–459,461 (motif VI) 7. 477, 479–482, 484, 485, 489, 490, 493 8. 521

For the ATP-binding site, we find that its hot-spot residues span an extensive network that not only covers the ATP-binding site, but also extends to the distant DNA/RNA-binding site (see Table 17.1.). Among them are several highly conserved residues of SF2 motifs implicated for helicase functions (for example, E291 and H293 of motif II, T322-P325 of motif III, Q460-G463 of motif VI). Some conserved DNA/RNA-binding residues (D412, V432) are found to be dynamically coupled with the ATP-binding site, supporting their important role in modulating ATPase activity by DNA/RNA binding. For the DNA/RNA-binding site, we find its hot-spot residues also span a network that not only overlaps with the DNA/RNAbinding site, but also reaches the ATP-binding site (see Table 17.1.).

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Among them are several highly conserved residues from SF2 motifs implicated for helicase functions (for example, H293 of motif II, A323–P325 of motif III, Q460 and R461 of motif VI). Some of those hot-spot residues are directly involved in ATP binding (for example, H293 of motif II, A323–P325 of motif III, Q460 and R461 of motif VI), which supports their critical role in modulating DNA/RNA binding in an ATP-dependent manner. Therefore, the above two sets of hot-spot residues, although defined for two spatially separated subsets of residues, share many common residues with each other, and mediate dynamical couplings between the ATP-binding site and the DNA/RNA-binding site. This dynamics-based mechanism facilitates the modulation of DNA/RNA binding by ATP binding and vice versa. 4. For the ATP-binding site of NS3 helicase, the open conformation is obtained from an NS3 crystal structure (PDB code: 1A1V), and the closed conformation is obtained from a bovine F1 ATPase crystal structure (PDB code: 1BMF, chains B & F) (see Table 17.2). The use of ATP-bound conformation of F1 ATPase to mimic the ATP-bound conformation of NS3 helicase is justified because both NS3 helicase and F1 ATPase belong to the large family of RecA-like ATPases, which possess highly conserved structural motifs in the ATP-binding site (15, 16).

Table 17.2 Alignment of residues between the HCV NS3 helicase and the bovine F1 ATPase at the ATP-binding site PDB code

Range of residue numbers

1A1V

A207–212

A229–231

A290–293

A322,323

A467

1BMF

F159–164

F186–188

F256–259

F308,309

B373

We compute the overlaps between the ATP-binding-induced conformational change and individual normal modes solved for an NS3 helicase structure (7). It is found that the ATP-bindinginduced conformational change is dominated by mode #1 (overlap ¼ 0.89) (7), suggesting that ATP binding causes domain 2 to close toward domain 3 and domain 1 to undergo a small twisting motion relative to domain 3 (see Fig. 17.1a). The ATP-binding-induced conformational change is also analyzed by dynamical domain partition. We find that it consists of two rotations between three dynamical domains (see Fig. 17.1c). Domain 2 closes toward domain 3 by a rotation of 26, which is accompanied by a smaller twisting motion of domain 1 by 11 relative to domain 3. As a result of the above domain motions, the

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helicase’s domain 2 grips tighter on DNA/RNA and simultaneously pushes it toward domain 1 and 3; meanwhile, by swinging away from DNA/RNA, domain 1 weakens its DNA/RNA-binding affinity, which allows DNA/RNA to slide through the cleft between domains 1 and 3. These coordinated domain motions are consistent with the key propositions of the inchworm model for NS3 helicase (17). References 1. Gorbalenya A. E., Koonin E. V., Donchenko A. P., and Blinov V. M. (1989) Two related superfamilies of putative helicases involved in replication, recombination, repair and expression of DNA and RNA genomes. Nucleic Acids Res. 17, 4713–4730. 2. Lee C. G. and Hurwitz J. (1992) A new RNA helicase isolated from HeLa cells that catalytically translocates in the 30 to 50 direction. J. Biol. Chem. 267, 4398–4407. 3. Ray B. K., Lawson T. G., Kramer J. C., Cladaras M. H., Grifo J. A., Abramson R. D., Merrick W. C., and Thach R. E. (1985) ATP-dependent unwinding of messenger RNA structure by eukaryotic initiation factors. J. Biol. Chem. 260, 7651–7658. 4. Karplus M. and McCammon J. A. (2002) Molecular dynamics simulations of biomolecules. Nat. Struct. Biol. 9, 646–652. 5. Tozzini V. (2005) Coarse-grained models for proteins. Curr. Opin. Struct. Biol. 15, 144–150. 6. Gwack Y., Kim D. W., Han J. H., and Choe J. (1997) DNA helicase activity of the hepatitis C virus nonstructural protein 3. Eur. J. Biochem. 250, 47–54. 7. Zheng W., Liao J. C., Brooks B. R., Doniach S. (2007) Toward the mechanism of dynamical couplings and translocation in hepatitis C virus NS3 helicase using elastic network model. Proteins 67, 886–896. 8. Zheng W. and Brooks B. R. (2005) Identification of dynamical correlations within the myosin motor domain by the normal mode analysis of an elastic network model. J. Mol. Biol. 346, 745–759. 9. Zheng W. and Brooks B. R. (2005) Probing the local dynamics of nucleotide-binding

10.

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pocket coupled to the global dynamics: myosin versus kinesin. Biophys. J. 89, 167–178. Hinsen K. (1998) Analysis of domain motions by approximate normal mode calculations. Proteins 33, 417–429. Atilgan A. R., Durell S. R., Jernigan R. L., Demirel M. C., Keskin O., and Bahar I. (2001) Anisotropy of fluctuation dynamics of proteins with an elastic network model. Biophys. J. 80, 505–515. Tama F. and Sanejouand Y. H. (2001) Conformational change of proteins arising from normal mode calculations. Protein Eng. 14, 1–6. Tirion M. M. (1996) Large amplitude elastic motions in proteins from a single-parameter atomic analysis. Phys. Rev. Lett. 77, 1905–1908. Krebs W. G., Alexandrov V., Wilson C. A., Echols N., Yu H., and Gerstein M. (2002) Normal mode analysis of macromolecular motions in a database framework: developing mode cosncentration as a useful classifying statistic. Proteins 48, 682–695. Bird L. E., Subramanya H. S., and Wigley D. B. (1998) Helicases: a unifying structural theme? Curr. Opin. Struct. Biol. 8, 14–18. Ye J., Osborne A. R., Groll M., and Rapoport T. A. (2004) RecA-like motor ATPases—lessons from structures. Biochim. Biophys. Acta 1659, 1–18. Kim J. L., Morgenstern K. A., Griffith J. P., Dwyer M. D., Thomson J. A., Murcko M. A., Lin C., and Caron P. R. (1998) Hepatitis C virus NS3 RNA helicase domain with a bound oligonucleotide: the crystal structure provides insights into the mode of unwinding. Structure 6, 89–100.

Chapter 18 Duplex Unwinding with DEAD-Box Proteins Eckhard Jankowsky and Andrea Putnam Abstract DEAD-box proteins, which comprise the largest helicase family, are involved in virtually all aspects of RNA metabolism. DEAD-box proteins catalyze diverse ATP-driven functions including the unwinding of RNA secondary structures. In contrast to many well-studied DNA and viral RNA helicases, DEAD-box proteins do not rely on translocation on one of the nucleic acid strands for duplex unwinding, but directly load onto helical regions and then locally pry the strands apart in an ATP-dependent fashion. In this chapter, we outline substrate design and unwinding protocols for DEAD-box proteins and focus on the quantitative evaluation of their unwinding activity. Key words: RNA, RNA helicase, DExH/D, DEAD-box, RNP, PAGE, unwinding, DDX, eIF4A, Ded1p, Mss116p.

1. Introduction Virtually all aspects of RNA metabolism involve DEAD-box proteins, the largest family of the helicase superfamily 2 (SF2). There are at least 25 DEAD-box proteins in Saccharomces cerevisiae, 37 in humans, and 5 in Escherichia coli (1). All currently available structures of DEAD-box proteins reveal the ‘‘helicase core’’ of SF2 proteins: two conserved RecA-like domains, arranged in tandem (2). DEAD-box proteins are defined by 13 characteristic sequence motifs, including all of the typical SF2 ‘‘helicase’’ motifs (Fig. 18.1). The name ‘‘DEAD-box’’ is derived from a short sequence in motif II, reading D-E-A-D, in single letter code (Fig. 18.1b). However, a given DEAD-box protein is not exclusively identified by the sequence of motif II, but by the

M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_18, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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Fig. 18.1. Characteristic DEAD-box sequence motifs. (a) Location of the characteristic sequence motifs in the two RecA like helicase domains that form the ‘‘helicase core’’ of DEAD-box proteins. Dark frames (e.g., Q, I, II) indicate motifs mainly involved in ATP binding, white frames (e.g., Ia, Ib, IV) indicate motifs mainly involved in RNA binding, and grey frames (e.g., III, Va, Vb) indicate motifs mainly involved in the coordination between ATP binding/hydrolysis and RNA binding/unwinding (2). The location of the characteristic sequence motifs corresponds to their location in other SF2 proteins. (b) Sequence logos for the characteristic DEAD-box motifs. Logos were created from a sequence alignment comprising the 25 DEAD-box proteins from S. cerevisiae, the 37 human DEAD-box proteins and the 5 DEAD-box proteins from E. coli.

sequence identity of all characteristic motifs (1). Several genuine DEAD-box proteins feature a motif II not reading D-E-A-D (e.g., Sub2p, ref. 3). DEAD-box proteins generally display RNA stimulated ATPase activity and most DEAD-box proteins unwind RNA duplexes in vitro, provided suitable substrates are used (2). Accordingly, DEAD-box proteins are often referred to as RNA helicases (4). Recent data suggest that certain DEAD-box proteins also unwind RNA duplexes in the cell (5, 6). Yet, other DEADbox proteins function as immobile clamps for subsequent formation of stable protein complexes on RNA (e.g., eIF4A-III in the exon junction complex, references 7–10), or in the remodeling of ribonucleoprotein complexes (11). In vitro, the remodeling of some ribonucleoprotein complexes can occur without duplex unwinding (12), and it is an open question whether unwinding of RNA secondary structure plays a role in all physiological functions of DEAD-box proteins (13).

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Nonetheless, ATP-dependent unwinding of RNA duplexes by DEAD-box proteins in vitro provides a direct readout for the ability of a given enzyme to remodel RNA in a reaction driven by ATP. In this chapter, we describe PAGE-based duplex unwinding assays for DEAD-box proteins. Several protocols and notes are largely identical to those published by us in an earlier volume of this series, in a chapter devoted to methods for measuring unwinding and protein displacement reactions for RNA helicases in general (14). In the chapter below, we focus exclusively on DEAD-box proteins, and outline the specific considerations for substrate design and unwinding reactions prompted by the distinct unwinding mechanism of these enzymes. In addition, we concentrate on the quantitative evaluation of the unwinding activity. Quantitative unwinding data are critical for systematic structure function studies, for evaluating effects of co-factors on DEAD-box proteins, and for illuminating the molecular mechanism of these enzymes. 1.1. The Distinct Unwinding Mechanism by DEAD-Box Proteins

Most DEAD-box proteins exert their function in a highly localized fashion, as parts of multi-component RNA–protein complexes such as those catalyzing pre-mRNA splicing, ribosome biogenesis, or translation initiation (2). The RNAs involved in these processes frequently contain thousands of nucleotides and adopt highly complex three-dimensional structures. Yet, cellular RNAs rarely, if ever, feature duplexes containing more than two continuous helical turns, and most helices in cellular RNAs are shorter than one helical turn. It has long been noted that DEAD-box proteins and other cellular RNA helicases have most likely evolved to unwind such ‘‘short’’ duplexes, and that this unwinding has to occur strictly locally, in order to preserve the integrity of the complexes in which the enzymes function (1). Recent evidence indicates that DEAD-box proteins are especially well adapted to the local remodeling of short RNA duplexes. In contrast to many well-studied DNA and viral RNA helicases, DEAD-box proteins do not rely on translocation on one of the nucleic acid strands for duplex unwinding, but directly load onto helical regions and then locally pry the strands apart in an ATPdependent fashion (15–17). Because the distinct unwinding mode of DEAD-box proteins has to be taken into account when designing appropriate substrates and unwinding experiments, it is instructive to briefly outline our current understanding of the mechanism by which DEAD-box proteins separate RNA duplexes (Fig. 18.2). DEAD-box proteins are loaded directly on the duplex region, aided by single-stranded or structured nucleic acid regions (Fig. 18.2, step 1). These regions have to be proximal, but do not need to be covalently connected to the duplex. As a consequence, DEAD-box proteins unwind tailed model substrates without apparent polarity, but nonetheless require unpaired regions for efficient unwinding (15, 16).

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Fig. 18.2. Current model for the kinetic mechanism of duplex unwinding by DEAD-box proteins (18). See text for details.

Duplex loading involves multiple protomers (17). The exact mechanism of the loading process has not yet been elucidated, although it is clear that the loading can occur at any place in the duplex, either at an end or in the middle, as well as on either strand (17). Duplex loading requires or is accompanied by ATP binding (17). Upon loading, the DEAD-box protein locally opens the duplex strands (ref. 17, Fig. 18.2, step 1). This strand opening also depends on ATP, and recent data indicate that ATP hydrolysis is not required for this strand opening, suggesting that ATP binding suffices (18, 19). The local helix opening reduces the number of basepairs in the duplex, and the remaining basepairs dissociate without further action from the enzyme (Fig. 18.2, step 2). Unwinding rate constants therefore decrease with duplex length and stability, because more or more stable basepairs dissociate slower (1, 2, 17, 20, 21). As no translocation of the DEAD-box protein is involved, unwinding of duplexes with more than two helical turns becomes exceedingly slow (22). ATP hydrolysis, although dispensable for duplex unwinding, is nonetheless critical for release of the DEAD-box protein from the RNA (Fig. 18.2 step 6), and thus for enzyme recycling (18). Not every ATP-driven local helix opening necessarily leads to complete strand separation. Even though helix opening does not require ATP hydrolysis, it is nevertheless possible that ATP hydrolysis occurs after the helix has been opened by ATP binding, but before the strands have separated (Fig. 18.2, step 4). While ATP hydrolysis promotes enzyme dissociation, strand separation may take place before the enzyme actually dissociates (Fig. 18.2, step 5). Alternatively, the enzyme may dissociate before complete

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helix separation (Fig. 18.2, step 7). In this case, the strands quickly re-anneal (refs. 19, 23, Fig. 18.2, step 8). Such ‘‘nonproductive’’ ATP hydrolysis events are more prevalent for longer and more stable duplexes, because the actual strand separation is slower and unwinding events thus occur less frequently. Accordingly, unwinding of longer or more stable helices involves larger numbers of hydrolyzed ATPs per duplex separated (19, 23).

2. Materials 2.1. Substrate Design

Despite the distinct unwinding mode of DEAD-box proteins, the simplest substrates for measuring duplex unwinding in vitro resemble those used for examining canonical DNA and RNA helicases. These substrates contain a defined duplex and an unpaired tail (Fig. 18.3). The unpaired region does not have to be physically attached to the duplex, but proximity to the duplex is required (16). The most straightforward way to ensure this arrangement is an extension of one of the duplex strands (Fig. 18.3).

Fig. 18.3. Duplex substrates. The asterisk indicates the most practical position of the radiolabel. The dotted line marks the unpaired region, which does not have to be physically attached to the duplex but must be in proximity to the duplex (16).

Like most other helicases, DEAD-box proteins generally unwind duplexes in vitro in a sequence non-specific manner and, thus, substrate sequences can be freely chosen. (2). Of note are DbpA-related bacterial DEAD-box proteins, which require a specific RNA hairpin for recruitment to the duplex that, however, can be of any sequence (24). As outlined above, DEAD-box proteins are well adapted to unwind short duplexes, but score poorly when confronted with duplexes exceeding one-and-a-half to two helical turns (20, 22, 25). Therefore, it is critical to use short duplexes for measuring unwinding by DEAD-box proteins. A practical lower limit for RNA duplexes is roughly 9–12 bp for reactions at room temperature and roughly 14–16 bp for reactions at 37C, but unwinding reactions have been measured with duplexes as short as 6 bp (15).

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In addition to duplex length, the stability of a given duplex sequence affects observed unwinding velocities; the more stable the duplex, the slower it is unwound (20, 25). Respective effects of duplex lengths vs. stability on unwinding rate constants of DEAD-box proteins have not yet been examined systematically, but available data suggest that neither overall stability nor the number of basepairs in a duplex exclusively determine observed unwinding rate constants (own unpublished results). Although DEAD-box proteins are able to unwind duplexes without single-stranded extensions, unwinding efficiency is generally enhanced by single-stranded regions (16, 17, Note 2). The single-stranded region facilitates loading of the enzyme on the duplex, as outlined above (Fig. 18.2). Accordingly, less enzyme is required to saturate tailed substrates, compared to blunt-end duplexes (17). Notwithstanding, the unwinding activities of some DEAD-box proteins, including eIF4A, are only weakly stimulated by single-stranded regions (20, 26). The orientation of the single-stranded region with respect to the duplex does not have strong effects on the unwinding efficiency, i.e., DEAD-box proteins will unwind duplexes with unpaired regions either 30 or 50 to the duplex region (Fig. 18.3). However, the single-stranded region has to exceed a critical length for maximal stimulation. For the DEAD-box protein Ded1p, maximal stimulation is seen with unpaired regions of 20 nt or more (17). For most DEAD-box proteins this critical length has not been determined, and it is possible that other DEAD-box proteins require fewer unpaired nucleotides for maximal stimulation. The sequence of the unpaired region does not appear to affect unwinding efficiencies, as shown for the DEAD-box proteins Ded1p and Mss116p (17). Since duplex loading can occur anywhere in the duplex, DEAD-box proteins are able to unwind mixed RNA–DNA duplexes, regardless of which strand is RNA (16, 17). When using RNA–DNA duplexes, it is important to note that DNA– RNA helices are less stable than RNA–RNA duplexes of the same sequence. For very short duplexes, stability can be problematic for DNA–RNA but not for RNA–RNA duplexes. It is most practical to utilize chemically synthesized oligonucleotides for mechanistic studies, because these RNAs have exactly the desired length. Although in vitro transcribed RNAs can be employed as well, the potentially heterogenous ends should be trimmed exactly to the appropriate length using either DNAzymes (27) or an alternative method such as RNaseH digestion or ribozyme cleavage.

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1. RNA oligonucleotides (deprotected and purified). 2. T4 polynucleotide kinase (10,000 U/mL) (New England Biolabs). 3. 10 T4 PNK buffer: 700 mM Tris–HCl (pH 7.6), 100 mM MgCl2, 50 mM dithiothreitol (DTT). 4. g32P gamma-labeled ATP (5 mCi/mL, 7000 Ci/mmol ¼ 23.8 mM). 5. 50 mM ATP. 6. 10 duplex annealing buffer: 100 mM MOPS, pH 6.5, 10 mM EDTA, 0.5 M KCl). 7. 10 TBE: 89 mM Tris base, 89 mM boric acid, 2 mM EDTA. 8. 1 TBE, 0.5 TBE (dilutions of the above). 9. Denaturing polyacrylamide gel: 20% acrylymide:bis 19:1, 7 M Urea, 1 TBE. 10. Non-denaturing polyacrylamide Gel: 15% acrylymide:bis 19:1, 0.5 TBE. 11. 5 Denaturing loading buffer: 80% formamide, 0.1% bromophenol blue (BPB), 0.1% xylene cyanol (XC). 12. 5 Non-denaturing gel loading buffer: 50% glycerol, 0.1% BPB, 0.1% XC. 13. Gel elution buffer: 300 mM NaOac, 1 mM EDTA, 0.5% SDS. 14. 100% EtOH. 15. 50% glycerol.

2.3. Unwinding Reactions

1. 10 Helicase reaction buffer (HRB): 400 mM Tris-HCl, pH 8.0, 5 mM MgCl2, 0.1% IGEPAL (tert-octylphenoxy-poly[oxyethylene]ethanol), 20 mM DTT (may vary for different helicases). 2. 5–25 nM radiolabeled RNA duplex. 3. Unlabeled RNA duplex, where applicable. 4. Unlabeled RNA trapping strand, where applicable. 5. Purified DEAD-box protein. 6. 20 mM equimolar ATP/MgCl2, or ATP analog/MgCl2. 7. 2 Helicase reaction SDS stop buffer (HRSB): 50 mM EDTA, 1% SDS, 0.1% BPB, 0.1% XC, 20% glycerol. 8. Non-denaturing polyacrylamide gel: 15% acrylymide:bis 19:1, 0.5 TBE. 9. Whatman chromatography paper (Fisher). 10. Gel dryer (equipped with membrane or oil pump). 11. PhosphorImager screen/PhosphorImager.

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3. Methods 3.1. RNA Labeling and Duplex Preparation (see Note 1)

1. Mix 1 mL 100 mM top strand RNA (Fig. 18.3), 1 mL 10 T4 PNK buffer, 2 mL 50 mM ATP, 2.1 mL g32P-ATP (23.8 mM), and 2.4 mL water. This yields a fraction of 0.33 radioactive labels/total labels. This fraction can be varied depending on the reference date for the g32P-ATP to give a stronger or weaker signal. 2. Add 1.5 mL T4 PNK. The final volume is 10 mL. 3. Incubate at 37C for 60 min (labeling efficiency is usually higher than 80%, see Note 3). 4. Inactivate the kinase by adding 2 mL denaturing gel loading buffer and heating to 95C for 2 min. 5. Pre-run 20% denaturing gel (0.8 mm thick) for roughly 30 min to reach separation temperature (50 C). 6. Load sample on gel and run at 30 V/cm for at least 2 h, depending on the length of the RNA. 7. Remove glass plates and expose gel to film (Kodak) to localize the labeled RNA. Cut out labeled strand, crush gel slice into a 1.5-mL tube using a syringe and add 600 mL elution buffer. The buffer has to cover the gel. Elute over night at 4C under gentle shaking. 8. Centrifuge to sediment residual acrylamide debris (16 K g, 5 min) 9. Remove elution buffer, split into two 1.5 mL tubes and add 3 volume of 100% EtOH and 2 mL 50% glycerol. 10. Place on dry ice for at least 1 h, and centrifuge (16 K g) for 30 min at 4C. 11. Remove supernatant and dry pellet in SpeedVac for at least 15 min. 12. Recombine pellets by mixing one pellet in 10 mL water, resuspend, and transfer to other tube. Rinse tube with 6 mL water and transfer again. Completion of transfer can be verified with a Geiger counter. 13. To the tube with combined, dissolved pellets, add 2 mL 100 mM unlabeled complementary RNA and 2 mL duplex annealing buffer. 14. Heat to 95C and gradually cool to room temperature over at least 30 min. 15. After room temperature is reached, add 4 mL 5 non-denaturing gel loading buffer and load on 15% non-denaturing PAGE (0.8 mm thick). Ensure that gel has been pre-run for at

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least 30 min and load sample on running gel. Run gel at room temperature (20 V/cm) for at least 1 h (longer for duplexes exceeding 30 bp). It is advisable to run labeled singlestranded RNA as a size marker on a parallel lane. 16. Remove glass plates, expose gel to X-ray film, localize and cut out labeled duplex. Crush gel slice into a 1.5-mL tube using a syringe and add 600 mL elution buffer. The buffer has to cover the gel. Elute over night at 4C under gentle shaking. 17. Repeat steps 8–11, and dissolve pellet in buffer (50 mM MOPS, pH 6.0, 50 mM KCl). 18. Measure concentration of purified duplex by aliquoting 1 mL into a scintillation vial and count in a scintillation counter. 3.2. Pre-steady State Unwinding Reactions

Unwinding reactions for helicases are commonly measured under pre-steady state conditions, i.e., with excess enzyme over the substrate. This is because unwound strands re-anneal spontaneously. This re-annealing process is a bimolecular reaction with both strands present at identical concentrations, and therefore, the effective re-annealing rate increases roughly with the square of the initial duplex concentration. Spontaneous annealing rate constants under usual reaction conditions are approximately kon 107 M1 min1 (22, 28), which translates into half lives of approximately t1/2 20 min at 5 nM of the respective strands, or t1/2 0.2 min at 50 nM strands. If annealing rates become comparable to unwinding rates, the reaction amplitude decreases, diminishing the unwinding signal. The re-annealing problem is often addressed by inclusion of excess scavenger RNA or DNA that is identical to the labeled strand. This scavenger binds to the unlabeled duplex strand, keeping the displaced labeled strand free. While this reaction regime efficiently enhances low reaction amplitudes, it complicates the quantitative interpretation of rate constants because the actual process measured is a strand exchange, which can occur spontaneously and can be catalyzed by DEAD-box proteins without ATP (22, 28). Moreover, the scavenger RNA can interact with free enzyme and alter the effective enzyme concentration. To maintain a straightforward, easily interpretable reaction regime, it is thus desirable to prevent significant spontaneous re-annealing during the unwinding reaction by choosing low substrate concentrations. There is no rational lower limit to the substrate concentration, but it should be noted that even at high-enzyme concentrations and near diffusion-limited enzymesubstrate binding, the substrate-binding step can become exceedingly slow at duplex concentrations below 0.1 nM. In addition, adsorption of RNA to the tube walls may become a problem at sub-nanomolar substrate concentrations, although this problem can be circumvented to some extent by inclusion of low

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concentrations of detergent in the reactions (see Sections 2 and 2.2). In our hands, duplex concentrations from 0.1 to 2 nM have proven practical choices. While spontaneous re-annealing of unwound duplex can be largely precluded, it has recently become clear that several DEADbox proteins (as well as other helicases) actively facilitate strand annealing (21, 22, 29, 30). The DEAD-box proteins Ded1p and Mss116p are among the strongest known strand annealers (21, 22). Although DEAD-box proteins do not display strand annealing activity per se (22), it is important to specifically probe for strand annealing activity when investigating an unknown DEAD-box protein. Protocols for strand annealing reactions are outlined below (Section 3.2.2). Prominent strand annealing activity is usually noted if unwinding reactions show distinct reaction amplitudes, even at sub-nanomolar substrate concentrations. If detected, strand annealing activity has to be taken into account when deriving quantitative information from unwinding time courses (Section 3.2.3). 3.2.1. Multiple Cycle, Presteady State Unwinding Reactions

Irrespective of whether or not a given DEAD-box protein displays strand annealing activity, unwinding reactions with enzyme excess (pre-steady state) are performed as shown in Fig. 18.4. Although DEAD-box proteins only perform one cycle of strand separation before dissociating from the substrate, as long as the enzyme can re-bind the substrate during the course of the reaction, multiple cycles of binding and unwinding can occur. This reaction regime is therefore referred to as multiple cycles. For pre-steady state, multiple cycle reactions, enzyme and substrate are pre-incubated without ATP, and the unwinding reaction is started by addition of ATP (Fig. 18.4a). Alternatively, enzyme can be incubated with ATP and the reaction can be started with substrate (Fig. 18.4b). Irrespective of the reaction start,

Fig. 18.4. Multiple cycle, pre-steady state reactions. (a) Scheme for reaction started with ATP. (b) Scheme for reactions started with substrate addition. See text for details.

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aliquots are removed at appropriate times and the reaction in these aliquots is stopped with SDS. Stopped aliquots can be stored on ice until the entire time course is measured. Aliquots are then applied to non-denaturing PAGE and the fractions of duplex and unwound substrate are determined for each aliquot (see Note 1). 1. Mix 3 mL of 10 HRB, 3 mL of 5 nM labeled RNA duplex, 3 mL 10 helicase (diluted with protein storage buffer) to desired protein concentration, and 21 mL of water to a final value of 30 mL (Note 4). 2. Incubate at reaction temperature for at least 5 min. This preincubation time may need to be optimized. 3. Aliquot 3 mL into 3 mL 2 helicase reaction stop buffer (HRSB) for zero time point. 4. Add 3 mL of 20 mM ATP/MgCl2 and mix rapidly by pipetting up and down to initiate the unwinding reaction. 5. Aliquot 3 mL reaction into 3 mL HRSB at desired time points and place on ice. 6. Aliquot 3 mL reaction into 3 mL HRSB and heat to 95C for 2 min (use as single-stranded size marker). Place on ice. 7. Load aliquots on a 15% non-denaturing PAGE (0.8 mm thick). 8. Run for 60 min at 10 V/cm (depending on duplex length). For short duplexes (<13 bp) run at 4C to prevent duplex dissociation during electrophoresis. 9. Remove glass plates and place gel on Whatman Chromatography paper. Cover gel with plastic wrap and dry on gel dryer. Quantify radioactivity in duplex (ID) and single strand or product (SSt) bands using a PhosphorImager. The fraction single strand [Frac SSt] formed at each time point is calculated according to: ½Frac SSt ¼ ISSt =ðISSt þ ID Þ 3.2.2. Strand Annealing Reactions

Strand annealing, in essence, is the reverse of the unwinding process. Annealing reactions and corresponding quantification are preformed accordingly. 1. Mix 3 mL of 10 HRB, 3 mL 10 ATP/MgCl2, 3 mL 10 Helicase (diluted with protein storage buffer) to desired protein concentration, and 21 mL of water to a final value of 27 mL. 2. Denature RNA duplex at 95C for 2 min. 3. Initiate the reaction by adding 3 mL of 5 nM denatured labeled RNA and mix rapidly by pipetting up and down to initiate the reaction. 4. Follow steps 5–9 of the pre-steady state unwinding protocol (Section 3.2.1).

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Most DEAD-box proteins tested to date catalyze strand annealing without ATP (Fig. 18.5). However, the bacterial DEAD-box protein CsdA requires ATP for this reaction (29). To specifically test for strand annealing activity in the presence of ATP, an unwinding and an annealing reaction at identical conditions are performed and the reaction amplitudes are compared (Fig. 18.6). If the DEAD-box protein under investigation catalyzes strand annealing, the reaction amplitude is identical for both unwinding and strand annealing reaction (Fig. 18.6c), because a steady state between unwinding and strand annealing is established (ref. 22, Fig. 18.6c, see Note 5).

Fig. 18.5. RNA unwinding and strand annealing activties of the DEAD-box protein Ded1p. (a) Representative PAGE of an unwinding time course of a substrate with 16 bp and a 25-nt unpaired region 30 to the duplex. Reactions were conducted at 19C with 0.5 nM substrate, 600 nM Ded1p, 1 mM ATP, as outlined in the protocol and Fig. 18.4a (5 min pre-incubation of Ded1p and substrate). Mobilities of duplex and single-stranded RNAs are indicated by the cartoons on the left, the asterisk marks the radiolabel. The block arrow emphasizes that the reaction was started with duplex RNA. The zero time point represents the reaction before ATP addition. Aliquots were removed between 5 s and 5 min. (b) Representative PAGE of a strand annealing time course with (left panel) and without Ded1p (right panel). No ATP was present during the reactions, otherwise reaction conditions were as in panel (a). The block arrow indicates that the reaction was started with single-stranded RNA. The zero time point represents the reaction before DED1 addition. Aliquots were removed from 0.5 to 30 min. Lane c on the right panel indicates the control for the duplex. (c) Steady state between ATP-driven unwinding and ATP-independent strand annealing, both catalyzed by a number of DEAD-box proteins (2, 22).

3.2.3. Calculating Rate Constants from Unwinding Time Courses Under Multiple Cycle, Pre-steady State Conditions

To obtain quantitative kinetic information from multiple cycles, pre-steady state unwinding reactions, the fraction of unwound duplex is plotted vs. reaction time. Observed rate constants kobs are determined by fitting these time courses to the integrated rate law for a homogenous first order reaction: Frac SSt ¼ A ð1 eðkobstÞ Þ A is the reaction amplitude. If no strand annealing is observed, kobs equals the observed unwinding rate constant k’unw, a composite rate constant describing a multitude of basic reactions steps, even at enzyme saturation (cf. Fig. 18.2). Nonetheless, in most cases k’unw is sufficient for a quantitative comparison of different unwinding reactions. If multiphasic time courses are observed, the

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Fig. 18.6. ATP-dependent steady state between RNA unwinding and strand annealing catalyzed by the DEAD-box protein Ded1p. (a) Representative PAGE for a strand annealing reaction with Ded1p and 0.1 mM ATP. The RNA substrate and reaction conditions, except for the ATP concentration, were identical to those used in Fig. 18.5. The block arrow indicates that the reaction was started with single-stranded RNA. Aliquots were removed from the reaction at the times indicated in panel (c), (below). (b) Representative PAGE for unwinding reaction with Ded1p and 0.1 mM ATP. The RNA substrate and reaction conditions, except for the ATP concentration, were identical to those used in Fig. 18.5. The arrow indicates that the reaction was started with duplex RNA. Aliquots were removed at the times indicated in panel (c), (below). (c) Time courses of strand annealing (*) and unwinding (l) reactions in the presence of DED1 and 0.1 mM ATP. Data points represent the average of at least three independent reactions; error bars indicate one standard deviation. The line through the data points represents the best fit to the integrated form of a homogenous first order rate law. Identical reaction amplitudes for unwinding and annealing reactions under identical reaction conditions indicate a steady-state between unwinding and strand annealing that can be approached from either the duplex or the single strands (22).

kinetic analysis becomes significantly more complex and requires separate measurements of association and dissociation of enzyme from the RNA, which will not be discussed here. If strand annealing activity is not negligible, the steady state between unwinding and annealing reactions has be considered, according to d½SSt=dt ¼ kunw ½Dpx kann ½SSt; kann is the observed annealing rate constant, [Dpx] is the duplex concentration. Since [Dpx] + [SSt] ¼ const., integration of the differential equation yields, after re-arrangement for the fraction of single strand: Frac SSt ¼ kunw ðkunw þ kann Þ1 ð1 eðkunwþkannÞt This equation can be used to directly extract observed unwinding and annealing rate constants from unwinding time courses. A more comprehensive description of the kinetic treatment for reactions where both unwinding and strand annealing occur can be found in refs. (22, 28). 3.2.4. Single Cycle, Presteady State Unwinding Reactions

Unwinding reactions by DEAD-box proteins reported to date were mostly conducted under the above described multiple cycle, pre-steady state conditions. As mentioned, unwinding rate constants obtained under these conditions describe a composite of elementary reaction steps (Fig. 18.2). To obtain kinetic

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information for more elementary processes, especially rate constants for the actual strand separation, it is critical to ensure that enzyme dissociating from the substrate during the course of the reaction can not re-bind. This reaction regime is termed single cycle because one measures only the reaction that corresponds to a single cycle of enzyme binding. Single cycle reactions have been used for the analysis of canonical, translocating DNA and RNA helicases, to measure helicase processivity, translocation rates, and translocation step sizes (27, 31). For DEAD-box proteins, which do not translocate, these parameters do not apply. However, at enzyme and ATP saturation, the measured rate constant under single cycle conditions corresponds directly to the rate constant for strand separation (Fig. 18.2), a fundamental kinetic parameter for any DEAD-box protein. Reaction amplitudes report the ratio between unwinding rate constant and dissociation rate constants, and this ratio, as a function of protein and ATP concentrations, is critical for quantification of functional ATP binding steps. In single cycle, pre-steady state unwinding conditions, rebinding of the DEAD-box protein to the substrate is prevented by inclusion of a large molar excess of scavenger RNA over the enzyme (Fig. 18.7). The scavenger RNA serves to trap all free enzymes in the reaction, and should therefore be of sufficient length to bind the DEAD-box protein under study. Singlestranded RNAs with more than 25 nt have proven practical. Furthermore, the scavenger RNA should not interact with the substrate RNA, to prevent undesired artifacts in the measured rate constant. Except for the inclusion of scavenger RNA which is added at the reaction start together with the ATP (Fig. 18.7), single cycle, pre-steady state unwinding reactions are conducted as multiple cycle, pre-steady state unwinding reactions (for protocol see Section 3.2.1).

Fig. 18.7. Reaction scheme for single cycle, pre-steady state reactions.

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Steady state reactions, the mainstay of classical enzyme kinetics, are only rarely used to characterize helicases. In contrast to many other enzymes, which mostly encounter excess substrate in the cell, both DNA and RNA helicases in their physiological environment are often present at higher concentrations than their substrates (2), and thus operate under presteady state conditions in many cases. Nevertheless, characterizing helicases under steady state conditions is instructive for elucidating kinetic parameters of steps such as enzyme recycling. Steady state reactions require substrate in excess over the enzyme. As outlined above, with increasing substrate concentrations the corresponding re-annealing of the unwound strands diminishes the fraction of unwound product in the steady state. Even though unwinding occurs continuously, the fraction of visible product may become too small to for meaningful data interpretation. To circumvent this problem, it is necessary to resort to a strand exchange regime (Fig. 18.8a). The unwinding reactions are conducted with duplex substrate at concentrations clearly exceeding enzyme concentrations, and an excess of unlabeled top strand over the substrate concentration (Fig. 18.8a,b). At the concentrations of substrate and unlabeled top strand necessary for these reactions (high nanomolar to micromolar range), the unwound strands immediately re-form duplexes. Due to the excess of unlabeled top strand, the labeled strand remains preferentially free, until the reaction reaches a steady state determined by the ratio of labeled and unlabeled top strand (Fig. 18.8c,d). In technical terms, steady-state unwinding reactions are performed in a fashion similar to pre-steady state reactions, with the main difference being substrate concentration and presence of unlabeled top strand (Fig. 18.8b). 1. Mix 3 mL of 10 HRB, 3 mL of 5 nM labeled RNA duplex, 3 mL of 20 mM unlabeled RNA duplex, 3 mL of 100 mM RNA short strand, 3 mL 10 Helicase (diluted with protein storage buffer) to desired protein concentration, and 15 mL of water to a final value of 30 mL. 2. Incubate at reaction temperature for at least 5 min. 3. Aliquot 3 mL into 3 mL 2 helicase reaction stop buffer (HRSB) for time point indicating reaction start. 4. Add 3 mL of a mixture containing 2 mM ATP/MgCl2 and mix rapidly to initiate the unwinding reaction. 5. Aliquot 3 mL reaction into 3 mL HRSB at desired time points and place on ice.

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Fig. 18.8. Steady-state unwinding reactions. (a) Strand exchange regime to follow steady-state unwinding reactions. Radiolabeled duplex substrate together with and excess of unlabeled top-strand are subjected to the reaction. Constant (steady state) unwinding of duplex and re-annealing of labeled as well as unlabeled top strand result in a reaction amplitude, which depends on the molar ratio between labeled and unlabled top strand in the reaction. For example, initial concentrations of 1 mM duplex (with labeled top strand) and 5 mM unlabeled top strand will result in a final reaction amplitude of A1 ¼ 0.83. (b) Reaction scheme for steady-state reaction. (c) Representative PAGE for a steady-state unwinding reaction of the DEAD-box protein Ded1p using a substrate with 10 bp and an unpaired region of 25 nt 30 to the duplex. Reactions were conducted as shown in panel (b) and described in the text; at 19C with 50 nM Ded1p, 2 mM labeled duplex, 10 mM unlabeled top strand, and 2 mM ATP. Aliqouts were removed as shown in panel (d), below. Cartoons indicate the mobility of duplex and free top strand, the asterisk marks the radiolabel. (d) Time course for the steady-state unwinding reaction shown in panel (c). The solid line represents a fit to a single exponential equation. See text for details regarding the calculation of kinetic parameters from the plot.

6. Take one aliquot and bring to 95C for 2 min. Allow the reaction to cool to the reaction temperature slowly. Aliquot 3 mL reaction into 3 mL HRSB (use as end point marker to determine correct steady state of the strand exchange reaction, Fig. 18.8c). Place on ice. 7. Follow steps 7–9 of pre-steady state unwinding protocol (Section 3.2.1) Steady-state unwinding velocities (v) are calculated from the time courses measuring the single exponential approach to the steady state according to (23): Frac SSt ¼ A ð1 e v ¼ kobs ½Dpx

ðkobsÞt

Þ

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kobs is the observed rate constant measured from the time course (Fig. 18.8d), A is the reaction amplitude, [Dpx] is the duplex concentration. Steady-state parameters (e.g., vmax, Km) are obtained by plotting unwinding velocities vs. enzyme or ATP concentrations, as in classical steady-state studies of enzyme kinetics. It is critical to examine whether the large excess of unlabeled top strand interferes with unwinding by the DEAD-box protein. This is best done by measuring the effect of the concentrations of unlabeled top strand (that will be used in the steadystate reactions) on pre-steady state reactions (Section 3.2.1). If the unlabeled top strand interferes with the DEAD-box protein under study, these effects need to be taken into account, and if necessary, the effective concentration of the enzyme must be corrected accordingly. We have found that top strands with 10 nt or less at concentrations of 5–10 mM have no notable effect on the DEADbox protein Ded1p, whereas top strands with 13 nt or more decrease the effective concentration of the enzyme under study (own unpublished results).

4. Notes 1. This protocol is largely identical to a protocol published by us in an earlier volume of this series, in a chapter devoted to protocols describing unwinding and protein displacement reactions for RNA helicases in general (14). 2. The unwinding activity of the DEAD-box protein CYT-19 is also enhanced by structured RNAs proximal to the duplex (15), and it is possible that strand separation by other DEADbox proteins is also stimulated by structured RNAs located proximal to the duplex. 3. If the concentration of duplex RNA is determined by measuring the radioactivity of labeled RNA, it is essential to maintain a reproducibly high efficiency of labeling with 32P. Variations of more than 30–50% in labeling efficiency cause significant fluctuations in the actual duplex concentration in the reaction and might critically affect the observed kinetics and/or the reaction amplitude. It is therefore important to verify labeling efficiency for each oligonucleotide used. 4. Reaction conditions such as temperature, salt concentrations, pH, and concentration and ratio of ATP to Mg2þ affect unwinding activity. These conditions will need to be optimized for each enzyme. The reaction temperature is critical, not only because of the effect of temperature on enzyme

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function, but also because of the effect of temperature on RNA annealing rates. The rate of spontaneous re-annealing increases by a factor of approximately 5–10 from 20C to 37C. The salt concentration in the reaction may significantly affect unwinding rates by DEAD-box proteins. For example, increasing monovalent salt concentrations from 20 to 150 mM decreases unwinding rates by approximately tenfold for the DEAD-box protein Ded1p (own, unpublished results). We have not observed strong differences between distinct monovalent cations and anions for unwinding activities tested, but significant effects of chloride on the activity of the DEAD-box protein eIF4A have been observed under certain reaction conditions (32). If an unknown DEAD-box protein is being tested, it might be advisable to utilize acetate or glutamate rather than chloride. Most DEAD-box proteins that have been investigated in vitro have affinities for ATP with Kd(ATP) in the range of 0.1-2 mM. ATP concentrations around 1-2 mM are thus a good initial point when optimizing reaction conditions. The ratio of ATP to Mg2þ also affects reaction rates and affinities of the enzyme for the RNA. To be hydrolyzed, ATP has to be bound to Mg2þ. If [ATP] > [Mg2þ], the uncomplexed ATP may still bind to the enzyme, but essentially acts as an inhibitor. Therefore, excess ATP can complicate quantitative interpretation of the data and excess of Mg2þ over ATP is clearly preferable for mechanistic studies. However, DEAD-box proteins can be sensitive to excess Mg2þ. Optimizing the ratio of ATP to Mg2þ for the respective set of experiments is therefore of considerable importance. 5. DEAD-box-protein-catalyzed strand annealing activity increases with the length of the respective strands. For a more comprehensive discussion of the subject see ref. (22). References 1. Linder P. (2006) Dead-box proteins: a family affair—active and passive players in RNP-remodeling. Nucleic Acids Res. 34, 4168–4180. 2. Jankowsky E. and Fairman M. (2007) RNA helicases—one fold for many functions. Curr. Opin. Struct. Biol. 17, 316–324. 3. Zhang M. and Green M. R. (2001) Identification and characterization of yUAP/ Sub2p, a yeast homolog of the essential human pre-mRNA splicing factor hUAP56. Genes Dev. 15, 30–35. 4. Linder P., Lasko P. F., Ashburner M., Leroy P., Nielsen P. J., Nishi K., Schnier J., and

Slonimski P. P. (1989) Birth of the D-E-AD box. Nature 337, 121–122. 5. Liang X. H. and Fournier M. J. (2006) The helicase Has1p is required for snoRNA release from pre-rRNA. Mol. Cell Biol. 26, 7437–7450. 6. Kos M. and Tollervey D. (2005) The putative RNA helicase Dbp4p is required for release of the U14 snoRNA from preribosomes in Saccharomces cerevisiae. Mol. Cell 20, 53–64. 7. Shibuya T., Tange T. O., Sonenberg N., and Moore M. J. (2004) eIF4AIII binds spliced mRNA in the exon junction complex and is

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13. 14.

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essential for nonsense-mediated decay. Nat. Struct. Mol. Biol. 11, 346–351. Ballut L., Marchadier B., Baguet A., Tomasetto C., Seraphin B., and Le Hir H. (2005) The exon junction core complex is locked onto RNA by inhibition of eIF4AIII ATPase activity. Nat. Struct. Mol. Biol. 12, 861–869. Andersen C. B., Ballut L., Johansen J. S., Chamieh H., Nielsen K. H., Oliveira C. L., Pedersen J. S., Seraphin B., Le Hir H., and Andersen G. R. (2006) Structure of the exon junction core complex with a trapped DEAD-box ATPase bound to RNA. Science 313, 1968–1972. Bono F., Ebert J., Lorentzen E., and Conti E. (2006) The crystal structure of the exon junction complex reveals how it maintains a stable grip on mRNA. Cell 126, 713–725. Bowers H. A., Maroney P. A., Fairman M. E., Kastner B., Luhrmann R., Nilsen T. W., and Jankowsky E. (2006) Discriminatory RNP remodeling by the DEAD-box protein DED1. RNA 12, 903–912. Fairman M., Maroney P. A., Wang W., Bowers H., Gollnick P., Nilsen T. W., and Jankowsky E. (2004) Protein displacement by DExH/D RNA helicases without duplex unwinding. Science 304, 730–734. Linder P. (2004) The life of RNA with proteins. Science 304: 694–695. Jankowsky E. and Fairman M. (2008) in RNA-Protein Interaction Protocols, Ed. Lin R. J. Humana Press, Totowa, NJ, Vol. 488, pp. 343–355. Tijerina P., Bhaskaran H., and Russell R. (2006) Nonspecific binding to structured RNA and preferential unwinding of an exposed helix by the CYT-19 protein, a DEAD-box RNA chaperone. Proc. Natl. Acad. Sci. U.S.A. 103, 16698–16703. Yang Q. and Jankowsky E. (2006) The DEAD-box protein Ded1 unwinds RNA duplexes by a mode distinct from translocating helicases. Nat. Struct. Mol. Biol. 13, 981–986. Yang Q., Del Campo M., Lambowitz A. M., and Jankowsky E. (2007) DEAD-box proteins unwind duplexes by local strand separation. Mol. Cell 28, 253–263. 18.Liu F., Putnam A., and Jankowsky E. (2008) ATP hydrolysis is required for DEAD-box protein recycling but not for duplex unwinding. Proc. Natl. Acad. Sci. U.S.A. 105, 20209–20214.

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19. Chen Y., and Russell R. (2008) The DEADbox protein CYT-19 uses a single ATP to completely separate a short RNA duplex. Proc. Natl. Acad. Sci. U.S.A. 105, 20203–20209. 20. Rogers G. W., Richter N. J., and Merrick W. C. (1999) Biochemical and kinetic characterization of the RNA helicase activity of eukaryotic initiation factor 4A. J. Biol. Chem. 274, 12236–12244. 21. Del Campo M., Tijerina P., Bhaskaran H., Mohr S., Yang Q., Jankowsky E., Russell R., and Lambowitz A. M. (2007) Do DEADbox proteins promote group II intron splicing without unwinding RNA? Mol. Cell 28, 159–166. 22. Yang Q. and Jankowsky E. (2005) ATP- and ADP-dependent modulation of RNA unwinding and strand annealing activities by the DEAD-box protein DED1. Biochemistry 44, 13591–13601. 23. Bizebard T., Ferlenghi I., Iost I., and Dreyfus M. (2004) Studies on three Escherichia coli DEAD-box helicases point to an unwinding mechanism different from that of model DNA helicases. Biochemistry 43, 7857–7866. 24. Diges C. M. and Uhlenbeck O. C. (2001) Escherichia coli DbpA is an RNA helicase that requires hairpin 92 of 23S rRNA. EMBO J. 20, 5503–5512. 25. Rogers G. W. J., Lima W. F., and Merrick W. C. (2001) Further characterization of the helicase activity of eIF4A. Substrate specificity. J. Biol. Chem. 276, 12598—12608. 26. Rogers G. W. J., Richter N. J., and Lima WF, M. W. (2001) Modulation of the helicase activity of eIF4A by eIF4B, eIF4H, and eIF4F. J. Biol. Chem. 276, 30914–30922. 27. Jankowsky E., Gross C. H., Shuman S., and Pyle A. M. (2000) The DExH protein NPH-II is a processive and directional motor for unwinding RNA. Nature 403, 447–451. 28. Yang Q., Fairman M. E., and Jankowsky E. (2007) DEAD-box-protein-assisted RNA structure conversion towards and against thermodynamic equilibrium values. J. Mol. Biol. 368, 1087–1100. 29. Chamot D., Colvin K. R., Kujat-Choy S. L., and Owttrim G. W. (2005) RNA structural rearrangement via unwinding and annealing by the cyanobacterial RNA helicase CrhR. J. Biol. Chem. 280, 2036–2044.

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30. Rossler O. G., Straka A., and Stahl H. (2001) Rearrangement of structured RNA via branch migration structures catalysed by the highly related DEAD-box proteins p68 and p72. Nucleic Acids Res. 29, 2088–2096. 31. 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. 32. Lorsch J. R. and Herschlag D. (1998) The DEAD box protein eIF4A. 1. A minimal kinetic and thermodynamic framework reveals coupled binding of RNA and nucleotide. Biochemistry 37, 2180–2193.

Chapter 19 Analysis of the RNA Helicase p68 (Ddx5) as a Transcriptional Regulator Samantha M. Nicol and Frances V. Fuller-Pace Abstract The DEAD box RNA helicase p68 (Ddx5) has been demonstrated to act as a transcriptional co-activator for a number of highly regulated transcription factors (e.g. estrogen receptor alpha and the tumour suppressor p53) and to be recruited to promoters of genes that are responsive to activation of these transcription factors, suggesting that it may play a role in transcription initiation. We have investigated the function of p68 as a co-activator of the tumour suppressor p53, with a particular emphasis on the importance of p68 in the induction of p53 transcriptional activity by DNA damage. These studies have involved RNAi-mediated suppression of p68 in cells expressing wild-type p53 and determining its effect on the expression of cellular p53 target genes in response to DNA damage. Additionally a significant amount of our research has focused on the study of the role of p68 in transcriptional initiation; this has included an investigation of the recruitment of p68 to the promoters of p53-responsive genes and of the importance of p68 in influencing recruitment of p53. Here we present detailed methods for RNAi knockdown of p68 expression, determination of its effect on expression of p53-responsive genes by quantitative RT-PCR and Western blotting, and chromatin immunoprecipitation techniques for determining recruitment of p68 and p53 to p53-responsive promoters. Key words: DEAD box RNA helicase, p68 (Ddx5), p53 tumour suppressor, transcription regulation, siRNA, chromatin immunoprecipitation, p21 promoter, quantitative PCR/RT-PCR.

1. Introduction The DExD/H box family of RNA helicases includes a large number of proteins that play important roles in cellular processes that involve modulation of RNA structures, such as RNA processing, RNA export, ribosome assembly and translation. However, it is now clear that several members of this family are multifunctional proteins that also play important roles in transcription regulation. One such M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_19, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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protein is the prototypical DEAD box protein p68 (Ddx5), which is known to be involved in several cellular processes that require manipulation of RNA structures, including mRNA/ rRNA processing and transcript clearance/export [reviewed in (1)]. However, work from several laboratories has demonstrated that p68 also acts as an important co-activator for a range of transcription factors that are themselves highly regulated, including estrogen receptor alpha (2), androgen receptor (3), the myogenic regulatory factor MyoD (4) and the tumour suppressor p53 (5). Moreover, p68 has been shown to be recruited to the promoters of responsive genes under conditions in which these transcription factors are activated (3–6), consistent with a role in transcription initiation. Additionally, in some contexts, p68 can act as a promoter-dependent transcriptional repressor (7), suggesting that its role in transcriptional regulation may be influenced by other factors that it interacts with at promoters. In our study of p53 co-activation by p68 (5), we demonstrated that p68 potently synergises with p53 to activate transcription from p53-responsive promoters. Strikingly, we showed that RNAi-mediated suppression of p68 expression in cells that express wt p53 inhibited induction of several cellular p53 target genes in response to DNA damage, suggesting that p68 is important for DNA damage-induced p53 transcriptional activation. We found that p68 is critical for the induction of the cell cycle arrest gene p21 and that it is recruited to the p21 promoter in response to treatment with the DNA damaging agent etoposide. Interestingly p68 recruitment was found to be p53dependent and to increase concomitantly with p53 recruitment in response to etoposide treatment. These observations, coupled with earlier findings that p68 interacts with RNA polymerase II and the co-activators CBP/p300 (2, 8), suggest that p68 may be involved in DNA damage-induced recruitment of components of the transcriptional machinery to the p21 promoter (9) or, perhaps, in the formation or stabilisation of the transcription initiation complex. This idea is underscored by the report that p68 appears to be important for the recruitment of Brg-1, TBP and RNA Pol II to MyoD-responsive promoters (4). To undertake these studies we have knocked down p68 expression using p68-specific siRNAs and determined its effect on the induction of p53-responsive genes after treatment with the DNA damaging agent, etoposide, by quantitative RT-PCR (qRT-PCR) and Western blotting. We have also examined recruitment of p68 and p53 to the p21 promoter by chromatin immunoprecipitation and semi-quantitative as well as quantitative PCR (qPCR). Detailed protocols for these procedures are described.

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2. Materials 2.1. siRNA Oligonucleotides

2.2. Quantitative RT-PCR (qRT-PCR) Primers for Determination of mRNA Expression Levels

Custom siRNAs: obtained as ‘ready-to-use’ annealed duplexes (Dharmacon/Thermo Scientific) as follows (see Note 1). p68 siRNA: 50 -AAC UCU AAU GUG GAG UGC GAC-30 control siRNA: 50 -CAG TCG CGT TTG CGA CTG G-30 1. qRT-PCR primers (Applied Biosystems, ABI) supplied ready to use, at a standard concentration of 20 . Aliquot prior to storage at –20C to prevent repeated freeze/thawing. p68—Hs00189323_m1 p21—Hs00355782_m1 Beta-Actin—Hs99999903_m1 2. qRT-PCR primers (MWG-Biotech) made up to a stock concentration of 100 pmol/ml in RNase-free water. Aliquot prior to storage at –20C to prevent repeated freeze/thawing. p53 forward—CAG CCA AGT CTG TGA ACT TGC A p53 reverse—GTG TGG AAT CAA CCC ACA GCT p53 probe—50 FAM—TCC CCT GCC CTC AAC AAG ATG TTT TGC C—TAMRA 30

2.3. Primers for Chromatin Immunoprecipitation (ChIP)

Primers for the p21 promoter and GAPDH control (MWG-Biotech) made up to a stock concentration of 100 pmol/ml, in RNasefree water. Aliquot and store at –80C. The primers used were adapted from those described in (9–11).

2.3.1. qPCR

p21-B forward:50 TGG AGA TCA GGT TGC CCT TTT 30 p21-B reverse:50 ACA AAG TTG TTG ATT GTC ACA TGC T 30 p21-B probe:50 FAM-TAG TCT CTC CAA TTC CCT CCT TCC CGG-TAMRA 30 p21-C forward:50 GTG GCT CTG ATT GGC TTT CTG 30 p21-C reverse:50 CTG AAA ACA GGC AGC CCA AG 30 p21-C probe:50 FAM-TGG CAT AGA AGA GGC TGG TGG CTA TTT TG-TAMRA 30 GAPDH forward:50 GTA TTC CCC CAG GTT TAC AT 30 GAPDH reverse:50 TTC TGT CTT CCA CTC ACT CC 30 GAPDH probe:50 FAM- CCG TCA AGG CTG AGA ACGGG TAMRA 30

2.3.2. Semi-quantitative PCR

p21-G forward:50 CAC CAC TGA GCC TTC CTC AC 30 p21-G reverse:50 CTG ACT CCC AGC ACA CAC TC 30 p21-M forward:50 CCC GGA AGC ATG TGA CAA TC 30 p21-M reverse:50 CAG CAC TGT TAG AAT GAG CC 30

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2.4. Antibodies 2.4.1. Western Blot Antibodies

1. p68—PAb204 (Upstate/Millipore)—0.2 mg/ml. 2. p53—DO-1 (available from various commercial sources, including Oncogene Science)—0.7 mg/ml. 3. p21—anti-p21WAF1 (Santa-Cruz C-19)—0.2 mg/ml. 4. Actin (Sigma-Aldrich A2066)—1 in 400 dilution of stock.

2.4.2. Chromatin Immunoprecipitation Antibodies

1. p68—PAb204 (Upstate/Millipore)—3 mg in 500 ml.

2.5. Cell Culture (see Note 2)

1. U2OS (human osteosarcoma) and MCF-7 (human breast adenocarcinoma).

2. p53—DO-1 (available from various commercial sources, including Oncogene Science)—3 mg in 500 ml.

2. Etoposide (Sigma-Aldrich) prepare as a 100 mM stock in dimethyl sulphoxide (DMSO) (Sigma-Aldrich) and use at concentrations ranging from 50 to 100 mM, depending on the experiment. 2.6. p68 siRNA

1. siRNA oligonucleotides as above (Dharmacon) 2. RNase-free water and 5 siRNA buffer (Dharmacon) 3. Opti-MEM1 medium (Invitrogen) 4. LipofectamineTM 2000 transfection reagent (Invitrogen)— for U2OS cells 5. LipofectamineTM RNAiMAX transfection reagent (Invitrogen)—for MCF-7 cells

2.7. Preparation of RNA and Protein from Cells

1. Qiagen RNeasy1 kit 2. Qiagen QIAshredderTM spin columns 3. 70% ethanol 4. b-mercaptoethanol 5. Acetone

2.8. cDNA Synthesis

1. Random hexamer—3 mg/ml (Invitrogen) 2. 100 mM dNTP set (GE Healthcare) 3. M-MLV reverse transcriptase—200 U/ml (Invitrogen) 4. RQ1 DNase—1 U/ml (Promega)

2.9. qRT-PCR for mRNA Measurements/ qPCR for ChIP

1. RNase-free H2O (Fisher Scientific) 2. 2 Universal PCR Master Mix (ABI) 3. Primers and probes as in Section 2.3 above 4. cDNA or DNA from ChIP 5. 96-well plates and optical caps (ABI)

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1. GoTaq1 Flexi DNA Polymerase (Promega) 2. 5 GoTaq1 buffer (Promega) 3. 25 mM MgCl2 4. 100 mM dNTPs (GE Healthcare)—used at 25 mM 5. DNA from ChIP

2.11. Chromatin Immunoprecipitation (ChIP)

1. Phosphate buffered saline (PBS): Prepare using five phosphate buffered saline tablets (Sigma-Aldrich) per litre H2O (see Note 9). 2. 1.5% Paraformaldehyde: 0.75 g of paraformaldehyde in 50 ml PBS. Heat to 65C with addition of 5 M NaOH until dissolved and then cool to 37C. 3. Cell collection buffer: Ice-cold PBS, protease inhibitor (P.I.) cocktail (Roche). 4. SDS lysis buffer: 50 mM Tris–HCl, pH 8.0, 10 mM EDTA, pH 8.0, 1% SDS, EDTA-free P.I. Cocktail. 5. Dilution buffer: 20 mM Tris–HCl, pH 8.0, 150 mM NaCl, 2 mM EDTA, pH 8.0, 1% TritonX-100, EDTA-free P.I. Cocktail. 6. Low Salt Wash: 20 mM Tris–HCl, pH 8.0, 150 mM NaCl, 2 mM EDTA, pH 8.0, 0.1% SDS, 1% Triton X-100, EDTAfree P.I. Cocktail. 7. High Salt Wash: 20 mM Tris–HCl, pH 8.0, 500 mM NaCl, 2 mM EDTA, pH 8.0, 0.1% SDS, 1% Triton X-100, EDTAfree P.I. cocktail (see Note 8). 8. LiCl wash: 10 mM Tris–HCl, pH 8.0, 0.25 M LiCl, 1 mM EDTA, pH 8.0, 1% Igepal, 1% DOC (see Note 9). 9. TE buffer: 10 mM Tris–HCl, pH 8.0, 1 mM EDTA, pH 8.0. 10. Elution buffer: 1% SDS and 0.1 M NaHCO3.

2.12. Other Reagents

1. Protein A sepharose: 50% slurry in 1 PBS (Sigma-Aldrich) 2. RNase A (Sigma-Aldrich) 3. Proteinase K: 10 mg/ml in sterile H2O (Sigma-Aldrich) 4. Salmon sperm DNA: 2 mg/ml stock in sterile H2O (SigmaAldrich) 5. QIAquick1 Gel extraction kit

2.13. Equipment

1. MSE Soniprep 150 sonicator. 2. Standard equipment for electrophoresis of polyacrylamide gels (as for SDS PAGE) and UV trans-illuminator for semiquantitative PCR. 3. Mx3005PTM TaqMan PCR machine with MxPro300 software. Analyse data by the Ct method in Microsoft Excel.

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3. Methods 3.1. Knockdown of p68 Expression by siRNAReverse Transfection

The methods below are based on ‘reverse transfection’ protocols described for the transfection reagents LipofectamineTM 2000 and LipofectamineTM RNAiMAX by Invitrogen, which are used for transfecting U2OS and MCF-7 cells respectively. Reverse transfection involves first placing the transfection reagent/siRNA mix onto the cell culture dish and then overlaying the cells, rather than overlaying the transfection reagent/siRNA mix onto a cell monolayer. For both cell lines we have found that reverse transfection gives the best results for p68 siRNA knockdown (see Note 3). The efficiency of p68 knockdown and its effect on p53 transcriptional activity, as measured by the induction of the p53 target gene p21, is examined by qRT-PCR and Western blotting (see Fig. 19.1).

A: qRT-PCR

B: Western blot

p68

1.4

1.4

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0.8 0.6 0.4 0.2

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p53

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p68 p68+

siRNA –/+ etoposide

NS NS+ p68 p68+ siRNA –/+ etoposide

Fig. 19.1. Effects of p68 siRNA knockdown on induction of expression of the p53 target gene p21 in response to treatment with the DNA damaging agent, etoposide. (a) qRT-PCR shows that p68 mRNA expression (left panel) is indeed knocked down by the siRNA, while there is little effect on p53 mRNA expression (middle panel). However, induction of p21 mRNA expression by etoposide is markedly inhibited (right panel). All values for mRNA are given relative to actin. NS: non-specific, control siRNA. /þ refers to the presence of etoposide in the medium (100 mM for 4 h). Please note the scale for p21 is different from those for p68 and p53. (b) Western blot showing effect of p68 siRNA knockdown on p68, p53 and p21 protein expression and the response to etoposide treatment (100 mM for 4 h). Actin is used as a loading control. 3.1.1. LipofectamineTM 2000 Reverse Transfection—U2OS Cells

1. Warm an appropriate amount (in a 50-ml falcon tube) of Opti-MEM1 medium to room temperature for 30 min prior to transfection. Protect the Opti-MEM1 from light by wrapping the tube in aluminium foil. 2. Pipette appropriate amounts of Opti-MEM1 and LipofectamineTM 2000 in a 15-ml falcon tube or 1.50-ml microfuge tube (depending on the volume required see table below), mix gently and incubate at room temperature for 5 min.

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3. Pipette an appropriate amount of Opti-MEM1 into the dish and add the required amount of siRNA oligonucleotide duplex (see table below); mix by tilting the plate. 4. Following the 5 min incubation, add the Opti-MEM1/LipofectamineTM 2000 complex to the dish, mix gently and incubate in the Microflow cabinet for 15 min. 5. During this incubation, trypsinise the cells, resuspend in antibiotic-free medium (D-MEM without penicillin/streptomycin) and determine the number of cells as described in Section 2.5. 6. Following the 15-min incubation, add the required amount of cells to the dish, mix gently to ensure even distribution of the cells and place back in the incubator for the required length of time, depending on the experiment. We find that 48 h is generally sufficient to achieve 60–90% knockdown of p68 expression. 7. Prior to harvesting cells treat with the DNA damaging agent, etoposide, as required. Cell Total culture Opti-MEM1 LipofectamineTM siRNA oligo (ml number dishes (ml)* 2000 (ml) of 20 mM stock) of cells 10 cm

1500

30

30

0.75–1.5 106

6 cm

500

15

15

7.5 105

6-well

100

5

5

2.5 105

*Note this amount of Opti-MEM1 is used both for preparing the transfection reagent and for adding to the siRNA; see Notes 2 and 3 above.

3.1.2. LipofectamineTM RNAiMAX Reverse Transfection—MCF-7 Cells

1. Warm an appropriate amount (in a 50-ml falcon tube) of Opti-MEM1 medium to room temperature for 30 min prior to transfection. Protect the Opti-MEM1 from light by wrapping the tube in aluminium foil. 2. Pipette an appropriate amount of Opti-MEM1 into the dish and add the required amount of siRNA oligonucleotide duplex (see table below); mix by tilting the plate. 3. Add the required amount of LipofectamineTM RNAiMAX, mix again by tilting the plate and incubate at room temperature for 20 min. 4. During this incubation, trypsinise the cells, resuspend in antibiotic-free medium (D-MEM without penicillin/streptomycin) and determine the number of cells as described in Section 2.5.

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5. Following the 20-min incubation add the required amount of cells to the dish, mix gently to ensure even distribution of the cells and place back into the incubator for the required length of time, depending on the experiment. We find that 48 h is generally sufficient to achieve 60–90% knockdown of p68 expression. 6. Prior to harvesting cells treat with the DNA-damaging agent, etoposide, as required.

3.2. RNA and Protein Extraction from Cells

3.2.1. RNA Extraction

Cell culture dishes

OptiMEM1 (ml)

LipofectamineTM RNAiMAX (ml)

siRNA oligo (ml of 20 mM stock)

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RNA extraction from untransfected or siRNA-transfected cells (which have been untreated or treated with the DNA damaging agent etoposide, depending on the requirements of the experiment) is performed using QIAshredderTM columns and RNeasy1 Mini Kits following the manufacturer’s instructions. 1. Lyse cells directly on the tissue culture dish as directed in the instructions and homogenise by placing the lysate onto a QIAshredderTM column. Centrifuge the column at 14,000 g for 2 min at room temperature. 2. Add an equal volume of 70% ethanol to the eluate from the QIAshredderTM column, mix gently by pipetting and apply to the RNeasy1 Mini column and centrifuge for 15 s at > 8000 g as instructed (in successive aliquots of up to 700 ml if the volume exceeds this). Retain 300 l of the flow-through for protein extraction and discard the remainder of the flow-through. 3. Wash the RNeasy1 Mini column serially according to the manufacturer’s instructions (see Note 4). After the washes elute the RNA in the RNase-free water provided in the kit. Following the final elution measure the RNA concentration at 260/280 nm using a spectrophotometer and store at –80C.

3.2.2. Protein Extraction

1. To the 300 ml flow-through from step 2, above, add 1200 ml of acetone and mix gently. Do not vortex (see Note 5). 2. Centrifuge at 14,000 g for 10 min at room temperature. 3. Aspirate off the acetone and add 100 ml of ice-cold ethanol. Mix gently, do not vortex.

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4. Centrifuge at 14,000 g for 10 min at room temperature. 5. Aspirate off the ethanol and air-dry the pellet for 10 min. 6. Add 150 ml of standard SDS sample buffer and boil for 10 min, vortexing occasionally. This protein can then be used for analysis by standard SDS-PAGE and Western blotting (see Fig. 19.1b). 3.3. Reverse Transcription/cDNA Synthesis

This is a two-step protocol involving DNA digestion followed by reverse transcription. This procedure can be carried out either using water baths at 37, 65 and 70C, or using a PCR machine programme for the required incubations. 1. For each reaction, place 1 mg of RNA in a microfuge tube and add 2 ml of 5 first strand buffer and 1 ml of RQ1 DNase. Make up the final volume to 10 ml with RNase-free water. Incubate at 37C for 30 min. 2. Following this incubation add 1 ml of RQ1 DNase stop solution and incubate the sample at 65C for 10 min and then chill on ice. 3. Add 2 ml of RNase-free water and 1 ml of random hexamer (300 ng/ml), incubate at 70C for 10 min followed by a brief chill on ice. Finally, add 6 ml of RT-mix (containing 2 ml of 5 first strand buffer, 2 ml of 10 mM dNTPs, 1 ml of 100 mM DTT and 1 ml M-MLV reverse transcriptase 200 U/ml) and incubate for 10 min at 25C followed by 1 h at 37C and, subsequently, 15 min at 70C. Store the resulting cDNA at –20C.

3.4. qRT-PCR (TaqMan)

For the analysis of mRNA levels in the untransfected/siRNAtransfected and/or untreated/treated samples, we perform quantitative real-time PCR using the Stratagene Mx3005PTM TaqMan PCR machine with MxPro300 software and cDNA (prepared as described in Section 3.3) as follows. 1. Set up qPCR reactions in 96-well plates (Applied Biosystems). All reactions include Universal PCR Master Mix, primer sets (including forward, reverse and probe) and the appropriate cDNA template. For every primer set a non-template control (NTC) is included to monitor possible primer contamination. (a). For ABI primers each reaction requires 6 ml of 2 Universal PCR Master Mix (Applied Biosystems), 0.6 ml of the appropriate ABI primer mix (including the forward and reverse primers and the probe) and 3.4 ml RNase-free water. (b). For MWG-primers each reaction requires 6 ml of 2 Universal PCR Master Mix, 1.2 ml of a primer mix (20 ml stock prepared using 17.5 ml of RNase-free water, 1 ml of forward and 1 ml of reverse primer and 0.5 ml of probe) and 2.8 ml of RNase-free water.

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However, to ensure consistency across all wells, prepare a stock master/primer mix for the required number of reactions, including NTC reactions, and dispense in 10-ml aliquots per well. 2. To each well add 10 ml of the master/primer mix (as above) and 2 ml of the appropriate cDNA, diluted 1 in 36 in RNase-free water. Seal the plates with Optical Caps (Applied Biosystems) and centrifuge briefly at 1000 g prior to loading into the TaqMan PCR machine. For our experiments the samples are amplified on the following Fast 2 Step PCR programme. (However, this might need to be adjusted for different primers/mRNAs.) 1 cycle at 95C for 10 min 40 cycles at 95C for 15 s 60C for 1 min Analyse TaqMan results by the Ct method in Microsoft Excel with an appropriate control sample as a reference sample and in all cases RNA levels are calculated relative to a ‘housekeeping’ mRNA, e.g. actin. For our studies on the effect of p68 siRNA knockdown on the induction of expression of p53 responsive genes after treatment with etoposide, we use untransfected cells, which have not been treated with etoposide, as the reference sample with a value of 1 relative to actin (see Fig. 19.1a and Note 6). 3.5. Chromatin Immunoprecipitation (ChIP)

The protocol below, adapted from previously described protocols (5, 6), gives the amounts used per 10 cm plate. We usually prepare six plates in each case and pool the harvested cells (step 2). For a standard ChIP, 2 106 cells are seeded in each plate 16–24 h prior to cross-linking with formaldehyde. When required, etoposide is added at 100 mM for 2 h prior to cross-linking. 1. To cross-link remove medium from cells, wash twice in warm 1 PBS and add 10 ml of 1.5% formaldehyde. Leave at 37C for 10 min. 2. Wash cells twice with ice-cold 1 PBS. To harvest cells add 2 ml of ice-cold collection buffer, leave for 2–3 min and then scrape off into a 15-ml falcon tube. 3. Pellet cells by centrifugation at 110 g for 10 min at 4C. 4. Remove supernatant and lyse pellet in 900–1000 ml of lysis buffer per 6 plates (depending on pellet size), transfer to a 1.5-ml centrifuge tube and leave suspension on ice for 10 min. 5. Sonicate (at a setting of 10 m) while keeping the tube on ice, seven times for 20 s, with 1 min between each sonication, to prevent overheating and DNA denaturation. This should give DNA fragments ranging from 200 to 1000 bp (see Note 7). Centrifuge at 10,000 g for 10 min at 4C to remove debris.

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6. Remove the supernatant (chromatin), measure the volume and place into a clean 15-ml falcon tube. At this stage take a 10-ml aliquot and check sonication efficiency by electrophoresis through a 1% agarose gel, ethidium bromide staining and visualisation using a UV trans-illuminator. 7. Dilute the chromatin fivefold in dilution buffer. At this stage the chromatin lysate can be aliquoted into 500- to 600-ml aliquots in microfuge tubes, snap frozen on dry ice and stored at –80C. 8. To set up the chromatin immunoprecipitation, pre-clear a 500- to 600-ml aliquot of the chromatin lysate with 30 ml of protein A 50% slurry with salmon sperm DNA (2 mg), on a rotating wheel at 4C for 2 h. This step removes any proteins binding non-specifically to the protein A. 9. Following pre-clearing, centrifuge the lysate at 400 g for 3 min at 4C. Place supernatant into fresh microfuge tube, removing 100 ml for use as an input control sample. Add an appropriate volume of the required antibody (to give 3 mg of antibody) and place on rotating wheel at 4C for overnight incubation. 10. The next day add 50 ml of protein A 50% slurry with salmon sperm DNA (2 mg) and place back on rotating wheel at 4C for 2 h. 11. Centrifuge at 400 g for 3 min at 4C. 12. Serially wash beads in 500 ml of low salt wash, high salt wash (see Note 8), LiCl wash and then twice in 1 TE buffer, on the rotating wheel at 4C for 5 min in each case centrifuging 400 g for 3 min at 4C between washes. 13. Finally centrifuge at 400 g for 3 min at 4C. 14. Elute complexes from the beads, by placing on rotating wheel at room temperature for 15 min with 50 ml of elution buffer. 15. Centrifuge at 1500 g for 3 min and repeat elution step (step 14). Combine the eluates from the two elution steps, i.e. final volume 100 ml. 16. Reverse cross-linking of the immunoprecipitated and the input samples by adding 4 ml of 5 M NaCl and 1 ml of RNAse A and placing in 65C water bath overnight. 17. Recover DNA by adding 4 ml of 1 M Tris, pH 6.5, 2 ml of 0.5 M EDTA, pH8.0, and 2 ml of 10 mg/ml proteinase K. Place in a 45C water bath for 1 h. 18. Purify DNA using QIAquick1 Gel Extraction Kit according to manufacturer’s instructions. 19. Store samples at –20C. These can now be used for qPCR or semi-quantitative PCR with appropriate primers.

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1. qPCR from ChIP DNA is performed largely as that for cDNA described in Section 3.4, with the appropriate primers and PCR programmes as indicated in that section. However, when setting up the plate add only 9.5 ml of the stock master/primer mix (instead of 10 ml) with 2.5 ml of the appropriate non-diluted ChIP DNA sample.

3.6. qPCR from ChIP

2. Analyse TaqMan results by the Ct method in Microsoft Excel with the relative input sample as the reference sample. The fold change from the input can be converted into percentage of input by multiplying fold change by 100. GAPDH and ‘no antibody’ controls are used to rule out non-specific immunoprecipitation (see Fig. 19.2a). A: q-PCR

B: semi-quantitative PCR

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p21 promoter PCR product Primer dimers

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Etoposide ChIP

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p68 (PAb204)

Fig. 19.2. Chromatin immunoprecipitation showing recruitment of p53 and p68 to the p21 promoter in response to etoposide treatment (100 mM for 2 h). (a) qPCR providing a quantitative measure of recruitment as a percentage of input. (b) Semi-quantitative measurement of the recruitment of p53 and p68; this also shows an increase in response to etoposide treatment, although precise comparison is, of course, not possible. In both (a) and (b), no PCR products were obtained in the ‘no antibody’ control (data not shown for (a) but values were below detection). The antibodies used for the IP are indicated.

3.7. Semi-quantitative PCR from ChIP

1. Dilute primers to a final concentration of 5 pmol/ml. Set up a 50-ml reaction with GoTaq1 Flexi DNA Polymerase (Promega) as follows: 10 ml of 5 buffer, 3 ml of 25 mM MgCl2, 0.4 ml of 25 mM dNTPs (GE Healthcare) 8 ml of the ChIP DNA sample, 5 ml of forward and reverse primers, 18.4 ml of RNase-free H2O and 0.2 ml of Taq. Note: for input samples use only 2.5 ml of input DNA. 2. Amplify the DNA using the following PCR programme: 1 cycle at 95C for 4 min 35 cycles at 95C for 30 s

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60C for 1 min 72C for 45 s 1 cycle at 72C for 5 min 3. Analyse PCR products by electrophoresis through a 15% Tris–glycine polyacrylamide gel as for standard sodium dodecylsulphate (SDS) polyacrylamide gel electrophoresis (PAGE) but omitting the SDS. 4. After electrophoresis, stain DNA by ethidium bromide (as for standard agarose gels) and visualise using a UV trans-illuminator (see Fig. 19.2b).

4. Notes 1. siRNA oligonucleotides are provided by Dharmacon/ Thermo Scientific as lyophilised samples. These should be made up in 1 siRNA buffer (prepared using the 5 siRNA buffer and RNase-free water, both purchased from Dharmacon/Thermo Scientific) as a 200 mM stock, aliquoted out in 100 ml amounts and stored at –80C. Prior to use, 20 mM working solutions are prepared from these stocks and aliquoted out in appropriate amounts, ranging from 20 to 50 ml depending on the scale of experiments performed, and again stored at –80C. This procedure avoids repeated freeze/thawing. 2. Maintain U2OS and MCF-7 cells in Dulbecco’s Modified Eagle’s Medium (D-MEM) supplemented with 10% foetal bovine serum, 2 mM L-glutamine, 50 units/ml penicillin and 50 mg/ml streptomycin (all from Invitrogen) and grow at 37C and 5% CO2. Omit the antibiotics for siRNA transfections. Detach for passaging cells by rinsing in phosphate buffered saline (PBS-see below) and trypsinise using trypsinEDTA (Invitrogen). Determine cell count by counting using a haemocytometer. Cells should be passaged every 2–3 days with a 1:3 dilution for MCF-7 and 1:5 dilution for U2OS. Do not allow cells to become too confluent and when passaging do not seed cells too sparsely. This is particularly important for MCF-7 cells. Discard cells after 15 passages for MCF-7 and 20 passages for U2OS. Early passage stocks are stored in liquid nitrogen and can be used for experiments 2– 3 passages after recovery from liquid nitrogen. 3. For siRNA knockdown the medium does not need to be changed if the cells are to be harvested 48 h after transfection. However, if a longer period is required to achieve

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sufficient knockdown the medium should be changed 24 h after transfection. It is useful to check whether changing the medium affects knockdown/recovery of cells depending on cell type and the target that is being knocked down by siRNA. 4. For the first wash (RW1 in the Qiagen RNeasy1 kit) we find that better/cleaner yields are obtained if the RW1 buffer is left on the column for 5 min prior to centrifugation. At the elution stage the RNase-free water is left for 1 min on the column prior to centrifugation. 5. Once acetone has been added to the 300-ml flow-through from the RNeasy1 Mini column the sample can be stored at –20C prior to continuing the protein extraction protocol. This can prove useful if several samples are being processed at the same time for RNA/protein extraction. 6. All values for mRNA levels are calculated relative to a ‘housekeeping’ mRNA, e.g. actin, GAPDH or TBP, with an appropriate reference, or control, sample taken as having a relative value of 1. For example, in our studies, untransfected cells (with normal levels of p68), which had not been treated with etoposide, were given a value of 1 relative to the actin mRNA level in that sample (see Fig. 19.1). 7. The sonication is one of the most important steps in this protocol. It is critical that the DNA fragment size obtained by sonication is reproducible (at 200–1000 bp) to avoid false-positive or negative results. It is best to sonicate a volume of between 600 and 800 ml and care must be taken to avoid over-sonication since this will result in denaturing the proteins. It is also important to keep the sample cold and to wait 1 min between the 20 s sonication bursts. It is also advisable to check sonication efficiency by electrophoresis of a sample. Once the procedure has been standardised for a particular system it is best to use the same sonication equipment as different sonicators can give very different results. 8. Normally the high salt wash after immunoprecipitation contains 0.1% SDS. However, for some antibodies (e.g. PAb204 for p68), this should be omitted (5, 6). Conditions should be optimised for different antibodies. 9. PBS can be prepared as a batch and stored at room temperature; LiCl wash buffer also can be prepared as a batch but aliquoted and stored at –20C. All other buffers must be prepared fresh just prior to use.

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Acknowledgements This work is supported by the Association for International Cancer Research (06-613). References 1. Fuller-Pace F. V. (2006) DExD/H box RNA helicases: multifunctional proteins with important roles in transcriptional regulation. Nucleic Acids Res. 34, 4206–4215. 2. Endoh H., Maruyama K., Masuhiro Y., Kobayashi Y., Goto M., Tai H., Yanagisawa J., Metzger D., Hashimoto S. and Kato S. (1999) Purification and identification of p68 RNA helicase acting as a transcriptional coactivator specific for the activation function 1 of human estrogen receptor alpha. Mol. Cell Biol. 19, 5363–5372. 3. Clark E. L., Coulson A., Dalgliesh C., Rajan P., Nicol S. M, Fleming S., Heer R., Gaughan L., Leung H. Y., Elliott D. J. et al. (2008) The RNA helicase p68 is a novel androgen receptor coactivator involved in splicing and is overexpressed in prostate cancer. Cancer Res. 68, 7938–7946. 4. Caretti G., Schiltz R. L., Dilworth F. J., Di Padova M., Zhao P., Ogryzko V., FullerPace F. V., Hoffman E. P., Tapscott S. J. and Sartorelli V. (2006) The RNA helicases p68/p72 and the noncoding RNA SRA are coregulators of MyoD and skeletal muscle differentiation. Dev. Cell 11, 547–560. 5. Bates G. J., Nicol S. M, Wilson B. J., Jacobs A. M., Bourdon J. C., Wardrop J., Gregory D. J., Lane D. P, Perkins N. D and Fuller-Pace F. V. (2005) The DEAD box protein p68: a novel transcriptional coactivator of the p53 tumour suppressor. EMBO J. 24, 543–553.

6. Metivier R., Penot G., Hubner M. R., Reid G., Brand H., Kos M. and Gannon F. (2003) Estrogen receptor-alpha directs ordered, cyclical, and combinatorial recruitment of cofactors on a natural target promoter. Cell 115, 751–763. 7. Wilson B. J., Bates G. J., Nicol S. M., Gregory D. J., Perkins N. D. and Fuller-Pace F. V. (2004) The p68 and p72 DEAD box RNA helicases interact with HDAC1 and repress transcription in a promoter-specific manner. BMC Mol. Biol. 5, 11. 8. Rossow K. L. and Janknecht R. (2003) Synergism between p68 RNA helicase and the transcriptional coactivators CBP and p300. Oncogene 22, 151–156. 9. Espinosa J. M., Verdun R. E. and Emerson B. M. (2003) p53 functions through stress- and promoter-specific recruitment of transcription initiation components before and after DNA damage. Mol. Cell 12, 1015–1027. 10. Kaeser M. D. and Iggo R. D. (2002) Chromatin immunoprecipitation analysis fails to support the latency model for regulation of p53 DNA binding activity in vivo. Proc. Natl. Acad. Sci. U.S.A. 99, 95–100. 11. Donner A. J., Szostek S., Hoover J. M. and Espinosa J. M. (2007) CDK8 is a stimulusspecific positive coregulator of p53 target genes. Mol. Cell 27, 121–133.

Chapter 20 A Method to Study the Role of DDX3 RNA Helicase in HIV-1 Chia-Yen Chen, Venkat R.K. Yedavalli, and Kuan-Teh Jeang Abstract Viral replication requires the use of host cell proteins and enzymes. Many viruses utilize viral helicases at various stages of their life cycle; these viruses have evolved to encode directly helicase or helicase-like proteins. In contrast, the genomes of retroviruses are devoid of viral helicases. Human immunodeficiency virus (HIV-1) has adopted the ability to use one or more cellular RNA helicases for its replicative life cycle. In this chapter, we briefly summarize the approach for assaying the RNA unwinding activity of RNA helicases measuring the effect of helicase inhibitors on HIV-1 replication. Key words: RNA helicases, human immunodeficiency virus type 1 (HIV-1), DEAD-Box domain, DDX3.

1. Introduction Helicases, including DNA and RNA helicases, are enzymes that separate stretches of duplexed DNA and/or RNA into singlestranded components in an energy-dependent manner. Based on motifs and sequences, RNA helicases are grouped into three superfamilies (SF1–SF3) and two smaller families (F4 and F5) (1). Most of the RNA helicases are in the SF2 superfamily and are conserved among different species. They can be found in organisms ranging from bacteria to humans to viruses. Viral replication requires the use of host cell proteins and enzymes. Many viruses utilize viral helicases at various stages of their life cycle; these viruses have evolved to directly encode helicase or helicase-like proteins. However, viruses that synthesize their genome within the cell nucleus tend to exploit cellular

M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_20, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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helicases and thus do not encode for any RNA helicases. Human immunodeficiency virus type 1 (HIV-1) is one example of this latter class of viruses. HIV-1 is a human retrovirus that packages two copies of positive strand full-length viral RNA per virion. HIV-1 infects human cells using a major receptor (CD4) with one of several co-receptors (CCR5, CXCR4, DC-SIGN), and it infects mostly T-helper (TH) cells, macrophages, and some microglial and dendritic cells (2). The viral genome is > 9 kilobases and encodes nine proteins. HIV-1 structural proteins include Gag (group-specific antigen), Pol (polymerase), and Env (envelope). In addition, HIV1 encodes two regulatory proteins, the transcriptional transactivator (Tat) and the regulator of post-transcriptional gene expression (Rev). The virus also has four genes that encode accessory proteins: Nef, Vif, Vpr, and Vpu. Replication of HIV-1 RNA starts inside an infected cell with the reverse transcription of its RNA into a cDNA form followed by its integration into the host chromosomal DNA to form a provirus. HIV-1 RNA is then transcribed from the proviral DNA by the cellular RNA polymerase II, processed like cellular mRNAs including 5’- and 3’-end modifications as well as splicing, and is then exported into the cytoplasm for translation. There are three processes pertaining to HIV-1 RNA that do not normally occur for cellular RNAs: nuclear export of introncontaining HIV-1 RNAs, packaging of viral RNAs into the spacelimited interior of virions, and completion of the reverse transcription of HIV-1 genomic RNA in the cytoplasm. In these regards, it is important to appreciate that HIV-1 encodes the nucleocapsid (NC) protein which bears RNA chaperone activity and has been shown to regulate HIV-1 RNA packaging and viral reverse transcription (3). A single HIV-1 transcript in its unspliced and spliced forms directs the synthesis of all viral proteins. Although export of intron-containing cellular transcripts from the nucleus into the cytoplasm is restricted in mammalian cells, HIV-1 must use unspliced RNA for the packaging of its genome into new virions and for the translation of Gag protein. Similarly, HIV-1 has to use a partially spliced transcript for translation of its Env protein. Thus, the virus must overcome the cell’s normal constraints of retaining unspliced/partially spliced RNAs in the nucleus preventing the nuclear-cytoplasmic export of such entities. To overcome these constraints, HIV-1 has evolved the Rev protein. Rev utilizes CRM1 as a cellular cofactor for Rev-dependent export of unspliced and partially spliced HIV-1 RNA. There is evidence that for export of HIV-1 RNAs, Rev/CRM1 activity also needs an ATP-dependent co-factor, RNA helicase DDX3 (4). DDX3 is a nucleocytoplasmic shuttling protein that binds CRM1 and localizes to nuclear membrane pores. Experimentally, abolition of the cell’s DDX3 activity suppressed Rev-RRE (Rev-responsive element)

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function for unspliced and partially spliced HIV-1 RNAs (4), supporting that DDX3 is a human RNA helicase that functions in the CRM1 RNA export pathway.

2. Materials 2.1. Purification of DDX3 Protein

1. LB broth. 2. Chitin beads (New England Biolabs (NEB)). 3. Column buffer: 20 mM Tris–HCI, 1000 mM NaCI, 0.5% Triton X-100, 0.1 mM EDTA, and 20 mM PMSF. 4. Cleavage buffer: 20 mM Tris–HCl (pH 7.5), 500 mM NaCl, 1 mM EDTA, and 50 mM DTT.

2.2. RNA Unwinding Assay

1. ATPase/helicase buffer: 20 mM Tris–HCl, pH 8.0, 70 mM KCl, 2 mM MgCl2, 2 mM dithiothreitol, 15 units of ATPase/ helicase buffer. 2. Maxiscript T7 in vitro transcription kit (Ambion). 3. MaxiScript T3 in vitro transcription kit (Ambion). 4. Sample loading buffer: 10 mM EDTA, 40% glycerol, bromphenol blue, xylene cyanol. 5. EasyTides uridine 50 -triphosphate,[a-32P] (Perkin Elmer). 6. 5 RNA annealing Buffer: 30 mM HEPES (pH 7.4), 100 mM potassium acetate, 2 mM magnesium acetate. 7. 10 Tris boric acid EDTA (TBE) buffer. 8. Accugel 29:1 (National Diagnostics), TEMED, 10% ammonium persulfate.

2.3. SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE)

1. Separating buffer (4): 1.5 M Tris–HCl, pH 8.7, 0.4% SDS (Protogel Resolving Buffer, National Diagnostics). Store at room temperature. 2. Stacking buffer (4): 0.5 M Tris–HCl, pH 6.8, 0.4% SDS (Protogel Resolving Buffer, National Diagnostics). Store at room temperature. 3. 30% acrylamide/bis solution (37.5:1 w/v) (ProtoGel (30%), National Diagnostics). 4. N,N,N,N’-tetramethyl-ethylenediamine (TEMED, Bio-Rad, Hercules, CA). 5. 10% Ammonium persulfate: prepare 10% solution in water. 6. Running buffer (10): 125 mM Tris, 960 mM glycine, 0.5% (w/v) SDS (Tris–glycine–SDS PAGE buffer (10), National Diagnostics). Store at room temperature.

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7. Prestained protein marker, broad range (7–175 kDa) (NEB). 8. Hoeffer S600 electrophoresis unit (GE Biosciences). 9. Sample buffer (SDS reducing buffer) (store at room temperature). a. Deionized water 3.8 ml, 0.5 M Tris–HCl, pH 6.8, 1.0 ml, glycerol 0.8 ml. b. 10% (w/v) SDS 1.6 ml, 2-mercaptoethanol 0.4 ml, 1% (w/v) bromophenol blue. c. 0.4 ml, add water to 8.0 ml. 2.4. Western Blotting

1. Tris–glycine transfer buffer: 25 mM Tris (do not adjust pH), 190 mM glycine, 20% (v/v) methanol. 2. Casein hydrolysate (USB). 3. CSPD (Applied Biosystems). 4. Nitro-Block (Applied Biosystems). 5. PVDF membrane (Millipore). 6. Semi-dry blotting apparatus. 7. 10 assay buffer: 200 mM Tris–HCl (pH 9.8), 10 mM MgCl2. 8. Blocking buffer: 1X PBS containing 0.2% casein hydrolysate, 0.1% Tween 20 detergent.

2.5. HIV-1 RT Assay

1. RT assay buffer: 60 mM Tris–HCl, pH 7.8, 75 mM KCl, 5 mM MgCl2, 0.1% Nonidet-P40, 1.04 mM EDTA, 5 mg/ ml poly rA, 0.16 mg/ml Oligi dT(18), 4 mM DTT. 2. Alpha 32P dTTP (Perkin Elmer). 3. DE81 paper (Whatman). 4. Phosphorimaging plate/Phosphorimager (Fuji Medical, FLA7000). 5. 20 SSC buffer (Invitrogen). 6. Plasmid pNL4-3 (proviral HIV-1 molecular clone).

3. Methods 3.1. Purification of DDX3 Protein

1. DDX3 cloned into pTYB11 vector, fused at its N-terminus with chitin binding domain was used to transform Escherichia coli BL21-DE3 cells. Cells were grown at 25C overnight.

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2. A single colony was selected and grown at 25C in LB medium containing 100 mg/ml ampicillin.When the OD 600 of the culture reaches 0.8, protein expression was induced at 15C with IPTG at a final concentration of 0.5 mM. 3. Cell extracts were prepared by lysing the cells by sonication in column buffer. Extracts were clarified by centrifugation and supernatant was used for protein purification. 4. Chitin column (20 ml for 1 culture) was equilibrated with 10 volumes of column buffer and slowly loaded with the clarified lysate. 5. The columns were then washed with at least 20 bed volumes of column buffer to thoroughly remove the unbound proteins. 6. The column were then washed two times with 3 bed volumes of cleavage buffer [20 mM HEPES or Tris–HCl (pH 7.5), 500 mM NaCl, 1 mM EDTA] and then once with cleavage buffer containing 50 mM DTT. The column flow was stopped without draining the cleavage buffer completely. The columns were then left at 8C for one day to allow for cleavage. 7. Cleaved protein was eluted by adding 0.5 bed volumes of cleavage buffer and continuing the column flow with cleavage buffer. 8. The eluted protein was dialyzed and concentrated before use. Purification was confirmed by SDS-PAGE electrophoresis and coomassie blue staining of gel. 3.2. RNA Unwinding Assay

1. The pBluescript plasmid was used to generate partially doublestranded RNA for the RNA unwinding assay (Fig. 20.1). The pBluescript vector was digested to completion with Kpn I and transcribed with T3 polymerase to generate a 120-base long transcript. The vector was also digested with EcoRI and transcribed with T7 polymerase in the presence of (32p) UTP. 2. The two complementary transcripts were purified and suspended in 1 annealing buffer. Samples were heated for 1 min to 95C and gently cooled to RT in the heating block. 3. The double-stranded RNA was then incubated with purified DDX3 in ATPase/helicase buffer. 4. After incubation for 30 min at 37C, the reactions were stopped by the addition of a solution containing 10 mM EDTA, 40% glycerol, bromphenol blue, and xylene cyanol. 5. The reaction was resolved in a 10% polyacrylamide gel (30 ml of Accugel 29:1, 10 ml 10 TBE, and 60 ml of H2O, 1 for 2 h). The gel was dried and exposed to a phosphorimaging plate.

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Fig. 20.1. A schematic illustration of a helicase assay. (a) Preparation of partial RNA duplex for RNA unwinding assay. (b) RNA unwinding assay.

3.3. SDS-PAGE Electrophoresis

1. Prepare a 0.75-mm-thick, 10% gel by mixing 3.75 ml of 4 separating buffer with 5.0 ml acrylamide/bis solution, 6.25 ml water, 150 mL ammonium persulfate solution, and 10 mL TEMED (Fig. 20.2). Pour the gel, leaving space of 3–4 cm for a stacking gel, and overlay with water. Wait until the gel polymerizes (approx 20 min). 2. Pour off water and prepare the stacking gel by mixing 0.65 ml of 30% acrylamide/0.8% bisacrylamide, 1.25 ml of 40 Tris-Cl/ SDS, pH 6.8, and 3.05 ml H2O. Add 100 ml of 10% ammonium persulfate and 5 ml TEMED. Swirl gently to mix. Insert the comb and pour the stacking gel avoiding air bubbles. Once the stacking gel polymerizes remove the comb gently and rinse the well with deionized water. 3. Prepare the running buffer by diluting 100 ml of the 10 Tris–glycine SDS running buffer with 900 ml of deionized water. 4. Add diluted Tris–glycine SDS running buffer to the upper and lower chambers of the gel unit and load the 25 mL of each sample in a well. Include one well for prestained protein molecular weight markers.

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Fig. 20.2. An illustration of an approach for assaying the effect of Helicase inhibitors on HIV-1 replication.

5. Complete the assembly of the gel unit and connect to a power supply. The gel can be run overnight at 45 V. 3.4. Western Blotting

1. The samples that have been separated by SDS-PAGE are transferred to PVDF membrane electrophoretically (Fig. 20.2). The gel unit is disconnected from the power supply and disassembled. The stacking gel is removed and discarded. 2. Five 3 M sheets wetted with transfer buffer are laid over the electrode (anode). The PVDF membrane is activated in methanol and after rinsing in transfer buffer laid on top of the 3 M paper. The separating gel is then laid on top of the PVDF membrane. Five more sheets of 3MM paper are wetted in the transfer buffer and carefully laid on top of the gel, ensuring that no bubbles are trapped in the resulting sandwich. The lid (electrode/cathode) is put on the gel sandwhich and the transfer is performed at a constant current of 250 mA for 2 h. 3. Following transfer, incubate the blot in blocking buffer (at least 30 ml) for 60–120 min. 4. Dilute primary antibody in blocking buffer (30 ml). Incubate with blot for 30–60 min. 5. Wash the membrane at least twice for 5 min in blocking buffer. Use at least 30 ml for all washes.

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6. Dilute secondary antibody-AP (alkaline phosphatase) conjugate 1:5000 in blocking buffer (30 ml). Incubate with blot for 30–60 min. 7. Wash three times for 5 min each as in step 5, then rinse two times for 2 min with 1 assay buffer. 8. Drain blots by touching a corner on a paper towel, then place on plastic wrap on a flat surface (do not let blots dry). 9. Pipette a thin layer of CSPD solution containing 5% NitroBlock onto the blot and incubate for 5 min. 10. Drain excess substrate solution and wrap the blot in plastic sheets. Smooth out bubbles or wrinkles. 11. Blots may be imaged by placing them in contact with standard X-ray film. 3.5. Reverse Transcriptase Assay (RT Assay) to Measure Inhibition of HIV

Reverse transcriptase assays provide an inexpensive approach to quantify the amount of virus present in the sample (Fig. 20.2). RT assay is an indirect measure of virus particles present in sample; it measures the amount of viral protein reverse transcriptase, which is incorporated into the virions (5). 1. Culture supernatants from pNL4-3 transfected HeLa cells or PBMC infected with HIV-1 were used in the assay. In case of HIV-1-infected peripheral blood mononuclear cells, PBMC, cell culture supernatants were collected every third day. HeLa cells were transfected with 2 mg of pNL4-3 (HIV-1 molecular clone) using Lipofectamine—Lipofectamine plus reagent. Forty-eight hours post-transfection, the culture supernatant was collected and assayed for RT activity. 2. 10 ml of culture supernatant was mixed with 50 ml of RT assay buffer containing 32P dTTP (2 ml/ml of RT assay buffer). 3. The reaction mix was incubated for 2 h at 37C. Subsequently 10 ml of the reaction mix was spotted on DE81 paper and allowed to dry. 4. The paper was washed three times with 2 SSC buffer, dried, and used to expose a phosphorimaging plate. 5. The RT activity was quantified by phosphorimager FLA-7000 (Fuji Medical) and also using scintillation counter (Beckman).

4. Notes In order to use radioactive isotopes in RNA unwinding assay and RT assays, users should follow their institutional guidelines for safe use of radioactive compounds in research. The users must be certified and properly trained in the use of radioactivity.

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Acknowledgments Research in KTJ’s laboratory is supported by intramural funds from NIAID, NIH, USA, and by the IATAP program from the office of the Director, NIH. References 1. Gorbalenya A. E. and Koonin E. V. (1989) Viral proteins containing the purine NTPbinding sequence pattern. Nucleic Acids Res. 17, 8413–8440. 2. Peterlin B. M. and Trono D. (2003) Hide, shield and strike back: how HIV-infected cells avoid immune eradication. Nat. Rev. Immunol. 3, 97–107. 3. Levin J. G., Guo J., Rouzina I., and MusierForsyth K. (2005) Nucleic acid chaperone activity of HIV-1 nucleocapsid protein: critical role in reverse transcription and

molecular mechanism. Prog. Nucleic Acid Res. Mol. Biol. 80, 217–286. 4. Yedavalli V. S., Neuveut C., Chi Y. H., Kleiman L., and Jeang K. T. (2004) Requirement of DDX3 DEAD box RNA helicase for HIV-1 Rev-RRE export function. Cell 119, 381–392. 5. Willey R. L., Smith D. H., Lasky L. A. et al. (1988). In vitro mutagenesis identifies a region within the envelope gene of the human immunodeficiency virus that is critical for infectivity. J. Virol. 62,139–147.

Chapter 21 Molecular Characterization of Nuclear DNA Helicase II (RNA Helicase A) Suisheng Zhang and Frank Grosse Abstract Nuclear DNA helicase II (NDH II) was first isolated from calf thymus using a DNA-unwinding assay. Subsequently it has been shown to be a homologue of human RNA helicase A (RHA) and the maleless protein (MLE) from Drosophila. Accordingly, the protein possesses both DNA and RNA unwinding activities. Also, it can use all four NTPs or dNTPs to fuel the reaction. At its N-terminus it possesses two double-strand RNA binding domains (dsRBD I and II), while the C-terminus comprises an imperfect glycine (G)- and arginine (R)-rich repeat, a so-called RGG-box that preferably binds to ssDNA or ssRNA. Many proteins interact with NDH II both at its N- and C-terminus and thereby mediate transcriptional regulation, RNA processing, and transport, the DNA damage response and genome surveillance. The latter includes the histone variant g-H2AX and the Werner syndrome helicase (WRN). Here we describe experimental approaches to obtain mechanistic information about this important nuclear helicase. Key words: DDX9, MLE, NDH II, RHA, unwindase, helicase assays, helicase function, domain mapping, DNA helicase, RNA helicase, DEXH box, maleless (MLE), double-stranded RNA binding domain (dsRBD), RGG-box.

1. Introduction Nuclear DNA helicase II (NDH II), also known as RNA helicase A, DDX9, and MLE protein, belongs to the superfamily II of DEXH box helicases (1). In addition to the seven canonical helicase motifs (Ia, Ib to VI) of this family NDH II further contains three nucleic acid binding motifs, i.e., two N-terminal doublestrand RNA binding domains (dsRBDs) and a C-terminal RGGbox (2). While a contribution of these additional nucleic acid binding sites to the unwinding activity of NDH II remains unclear, there is some evidence for their involvement in targeting specific M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_21, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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nucleic acid structures and mediating protein–protein contacts (1, 3). Increasing evidence is emerging for clinical importance of this helicase, for example, in autoimmune diseases (4), in tumorigenesis (5), and in checkpoint control processes (6). NDH II seems also to be involved in the DNA damage response as suggested by its physical and functional interaction with the histone variant gH2AX (7) and the Werner syndrome helicase (8). WRN helicase is deficient in patients displaying a human progeria that leads to premature ageing and a predisposition to tumorigenesis (9). Despite the fact that NDH II was initially identified as a DNA helicase, some workers had difficulties in measuring its DNA but interestingly not its RNA unwinding activity. Although it is quite clear now that NDH II and its homologues can unwind both nucleic acid substrates equally well (10, 11), the obvious technical problems in studying DNA-unwinding induced us to provide detailed experimental protocols for a successful examination of the enzymatic properties of this essential enzyme. The procedures presented here can be adapted to RNA unwinding studies as well. In this case, however, further precautions, such as treating buffers with diethylpyrocarbonate and/or using RNase inhibitors such as RNasin, should be considered. Hopefully the detailed protocols given here will help to rapidly expand our knowledge on human NDH II and its orthologues from other higher eukaryotes such as worms, plants, and insects.

2. Materials 2.1. Chemicals and Solutions 2.2. Native and Recombinant Proteins

All chemicals were of at least analytical grade, the water was bidistilled, and all aqueous solutions were sterilized adequately before use. 1. Native NDH II can be purified from calf thymus by conventional chromatography (12). The final purification step may either be performed with ATP-agarose (12) or preferably with poly(rI)(rC)-agarose (11), the latter of which is currently not commercially available. A useful procedure for the preparation of poly(rI)(rC)-agarose is given by (13). Bovine NDH II obtained from conventional column purification consisted of two polypeptides of 130 and 100 kDa, representing degradation products with losses of the N- and C-terminal nucleic acid binding domains. Despite this, both degraded forms were active in nucleic acid unwinding and nucleic acid-stimulated ATP hydrolysis (12). Human NDH II can be easily expressed in insect cells as an N-terminal (His)6-tagged recombinant protein using the Bacmid technology (Life

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Technologies) (2). Recombinant NDH II was purified by Ni2+-NTA-agarose and then by poly(rI)(rC)-agarose. These two steps are sufficient to yield more than 90% homogenous enzyme free of nucleases, which inevitably affect the helicase assay. To avoid nuclease contaminations, NDH II should be collected from the final elution step in individual fractions and those showing the highest purity (with regard to nuclease contaminations) should be used. Protein concentration was determined by densitometry after SDS-PAGE, using bovine serum albumin as a standard. The protein remains active for more than one year when stored at –20C in the presence of 50% glycerol. 2. The nucleic acid binding of domains of NDH II have been mapped by Northwestern blotting. To achieve this goal, fragments of NDH II were expressed as GST-fusion proteins as described before (2, 8). Alanine substitutions of phenylalanine 211 (F211A) or lysines 235 and 236 (K235A, K236A) of dsRBD II were introduced by PCR-based site-directed mutagenesis (14). The GST-fusion proteins were expressed in Escherichia coli followed by suspension of the bacteria from 2 ml culture medium in 500 ml of SDS-PAGE loading buffer (15), a brief sonication and heating at 95C for 5 min. The recombinant proteins can be stored at –20C for more than 1 year. 2.3. Oligonucleotides, Bacteriophage M13 Single-Stranded DNA (ssDNA) and Synthetic Homopolymeric Nucleic Acids

1. An oligodeoxyribonucleotide (50 -ACTCTAGAGGATC CCCGGGTACGTTATTGCATGAAAGCCCGGCTG,45mer) with 22 nucleotides complementary to M13mp 18’s position 6243–6264, and a non-complementary tail of 23 nucleotides at its 30 end was designed. The oligonucleotides should be of high quality and purified by electrophoresis, such as those from Purimex (Grebenstein, Germany). Small aliquots were stored at a concentration of 20 mg/ml in order to avoid repeated freeze and thaw cycles (see Note 1). 2. M13mp18 ssDNA was prepared as described (16), but can be also obtained commercially (GE-Healthcare). The DNA was dissolved in TE buffer (10 mM Tris–HCl, pH 8.0, and 1 mM EDTA) and its concentration was measured by optical absorbance at 260 nm where 1 optical density per cm (1 OD) equals 40 mg/ml M13-DNA. To avoid multiple freeze and thaw cycles, the DNA was divided into small aliquots at about 3 mg/ml (1 pmol/ml) for storage at –20C. 3. Poly(rI) or poly(rI)(rC) (GE-Healthcare).

2.4. Radioactive Labeling of the Nuclei Acids

1. 50 -end labeling of oligo- and polynucleotides was achieved by T4 polynucleotide kinase (New England BioLabs). 2. g-32P-ATP (5000 Ci/mmol) (GE-Healthcare).

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3. 50 -end labeling buffer: 80 mM Tris–HCl, pH 7.4, 10 mM MgCl2, and 5 mM DTT. 4. To separate the free nucleotide from the labeled nucleic acid the spun column method employing Sephadex G-50 was used (14). Autoclaved and silanized glass wool was used as plug for 1-ml tuberculin syringes filled with column material. Alternatively, commercially available spun columns can be used (GE-Healthcare). 2.5. Preparation of the Helicase Substrate

1. DNA annealing buffer (2 ): 12 mM Tris–HCl, pH 7.5, 14 mM MgCl2, 100 mM NaCl, and 2 mM DTT. 2. Bio-gel A-5 M (Bio-Rad). 3. Bio-gel A-5 M chromatography buffer: 10 mM Tris–HCl, pH 7.5, 100 mM NaCl, and 0.5 mM EDTA.

2.6. Helicase Assays

1. 40% polyacrylamide gel solution with a ratio of 39–1 (acrylamide to bisacrylamide) prepared in water and stored at 4C. 2. 5 TBE buffer: 25 mM Tris–borate, pH 8.3, 2.5 mM EDTA. 3. Native polyacrylamide gels (15%) were prepared by mixing 3.75 ml of the 40% gel stock solution, 2 ml of 5 TBE, and water to a volume of 10 ml. 60 ml of 10% ammonium peroxidisulfate (APDS) and 5.5 ml N,N,N’,N’-tetramethylethylene diamine (TEMED) and 8 10 cm gels with a thickness of 1 mm were cast. Polymerization was allowed to complete at room temperature overnight, or, if necessary, at 37C for at least 4 h. 4. 5 helicase reaction buffer: 100 mM Tris–HCl, pH 7.5, 50% glycerol, 17.5 mM MgCl2, 500 mg/ml BSA, and 25 mM DTT; store at –20C. 5. A 100 mM ATP solution was prepared and neutralized using 1 M Tris–base and pH indicator sticks; store at –20C. 6. Helicase reaction stop buffer: 1% SDS, 200 mM EDTA, 50% glycerol, 0.1% bromophenol blue, and 0.1% xylene cyanole FF; store at –20C.

2.7. Filter Binding Assay

1. 96-well vacuum blotter (Millipore, Dassel, Germany) 2. Nitrocellulose membrane (Protran B85, 0.45 mm) with the size of the 96-well vacuum blotter (Millipore). 3. Filter binding buffer: 20 mM HEPES, pH 8.0, 50 mM NaCl, 1 mM DTT, and 1 mM EDTA. 4. The nitrocellulose membrane was pre-treated by incubation with 0.3 N NaOH for 10 min, washed two times for 5 min each with water, and then equilibrated with the above binding buffer for at least 16 h (see Note 2).

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5. Whatman 3MM paper was cut out to yield the same size as the nitrocellulose membrane. The 3MM paper was equilibrated with binding buffer just before use. 2.8. Agarose Gel Shift Assay

1. Agarose (Life Technologies, GIBCO). 2. 1.5% agarose gel in 1 TBE buffer: 5 mM Tris–borate, pH 8.3, 0.5 mM EDTA. 3. Agarose gel shift loading buffer: 0.25% bromophenol blue, 0.25% xylene cyanol FF, and 30% glycerol in water (14).

2.9. Northwestern Blot

1. 10% SDS polyacrylamide gel (15). 2. Hybond-C nitrocellulose membrane (GE-Healthcare). 3. Proteins were electro-transferred from the SDS polyacrylamide gel to the nitrocellulose membrane using a semi-dry blot apparatus (e.g., from Sigma). 4. Transfer buffer: 25 mM Tris, 190 mM glycine, 0.1% SDS, and 20% methanol. 5. Hybond-C nitrocellulose membrane, cut to a similar size of the gel, was wetted with water, and then equilibrated with transfer buffer for 15 min before use. 6. Six sheets of Whatman 3MM paper, cut to the same size as the gel and equilibrated with transfer buffer. 7. Ponceau solution: 0.1% (w/v) Ponceau S in 10% (w/v) acetic acid. 8. TBS buffer: 25 mM Tris–HCl, pH 7.8, 140 mM NaCl, and 3 mM KCl. 9. 8 M urea in TBS solution. 10. Binding buffer: 10 mM HEPES, pH 8.0, 25 mM NaCl, 10 mM MgCl2, 0.1 mM EDTA, 1 mM DTT. 11. Blocking buffer: dissolve 5% (w/v) milk powder in binding buffer (see Note 6).

3. Methods 3.1. Preparation of DNA Helicase Substrate

1. The 45-mer deoxyoligonucleotide was 50 -labeled by T4 polynucleotide kinase in a 20-ml reaction mixture containing 20 pmol oligonucleotide, 60 mCi g-32P-ATP (18 pmol), and 20 units of T4 polynucleotide kinase in 50 -labeling buffer for 30 min at 37C. 2. At the end of the labeling reaction, the polynucleotide kinase was inactivated by heating at 65C for 15 min.

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3. Free g-32P-ATP was removed from the labeled oligonucleotide by the spun column method (14). 4. The labeled oligonucleotide in the eluant of the spun column (100 ml) was mixed with 20 pmol M13mp18 ssDNA in 20 ml. 5. 120 ml 2 annealing buffer was mixed with the nucleic acids and water to a final volume of 240 ml. 6. Annealing proceeded by heating to 95C for 10 min, followed by incubation at 60C for 30 min, at 37C for 30 min, and finally at room temperature for 30 min. 7. The annealing mixture was loaded onto a Bio-gel A-5 M column (1 ml bed volume, Bio-Rad) pre-equilibrated with Bio-gel A-5 buffer and then eluted with the same buffer. 8. Fractions of 100 ml were collected. Elutions of the annealed DNA and the unannealed oligonucleotide were monitored by liquid scintillation counting. These measurements can be done with closed 0.25 ml eppendorf tubes placed into the scintillation bottles. 9. The eluted radioactive fractions in the void volume (the first peak) contained annealed DNA for the helicase assay. These fractions were not pooled in order to select the substrate with the highest specific radioactivity. 3.2. Helicase Assay

1. A 15% native polyacrylamide gel was prepared. 2. A 10-ml helicase reaction mixture was prepared to contain 20 mM Tris–HCl, pH 7.5, 10% glycerol, 3.5 mM MgCl2, 100 mg/ml BSA, 5 mM DTT, 6 mM (nucleotides) helicase substrate, 3 mM ATP, and up to 280 nM NDH II (see Note 4). 3. The reaction mixture was incubated at 37C for 30 min. 4. The reaction was terminated by adding 3 ml stop solution to the reaction mixture, and immediate chilling on ice. 5. The reaction mixture was loaded to the native polyacrylamide gel. 6. Electrophoresis was performed at 1 mA/cm constant current for about 45 min until the bromophenol blue marker reached the bottom of the gel. 7. The gel on one glass plate was wrapped into Saran wrap and then exposed at –80C to X-ray film or PhosphoImager screen. 8. Unwinding of dsDNA was determined according to the separation of the released oligonucleotide that migrated considerably faster than the annealed dsDNA, which remained at the upper position of the gel. The position of the free oligonucleotide can be referred to as a ‘‘heat control’’ obtained by

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Fig. 21.1. Measurement of the DNA-unwinding activity of NDH II. (a) NDH II unwinds dsDNA in the presence of any one of the four common rNTPs or dNTPs. This is in contrast to the NDH II-copurifying helicase NDH I that prefers ATP or dATP. (b) Time-dependent unwinding activity of NDH II. Unwinding can be quantified as time- or enzyme concentrationdependent percentage of displaced oligonucleotide from the dsDNA substrate. The displacement can be determined by scintillation counting of the excised radioactive bands or by evaluation of the PhosphoImager signal (12).

denaturing the substrate at 95C for 5 min and subsequent chilling on ice with 3 ml stop solution. This releases the labeled oligonucleotide quantitatively from the ssDNA template. On the other hand, a control without addition of any protein provided the marker for the annealed substrate (Fig. 21.1). 3.3. Filter Binding Assay

1. Synthetic homopolymeric nucleic acids such as poly(rI) or poly(rI)(rC) were 50 -labeled by T4 polynucleotide kinase and g-32P-ATP following a similar molar ratio of nucleic acid to g-32P-ATP as described above for preparing the helicase substrate. A labeled M13 ssDNA was obtained by hybridization with a 50 -labeled oligonucleotide as described above for the helicase substrate. 2. 2 mM (nucleotides) of the nucleic acid probes were incubated with increasing amounts of NDH II (from 0 to 280 nM) in 10 ml binding buffer containing 20 mM HEPES, pH 8.0, 50 mM NaCl, 1 mM DTT, and 1 mM EDTA for 20 min at room temperature. 3. Meanwhile the 96-well vacuum blotter was mounted by placing the pre-treated nitrocellulose membrane over Whatman 3MM paper that was pre-equilibrated with binding buffer. A pre-filtration of the binding buffer through the membrane was examined to ensure that there was no blockage or leakage between the nitrocellulose membrane and the blotter.

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4. The samples at the end of incubation were loaded to the slots of the blotter and then sucked through the nitrocellulose membrane using a vacuum pump. 5. The nitrocellulose membrane was washed under vacuum for three times each with 500 ml of binding buffer. 6. The vacuum blotter was dismantled and the nitrocellulose membrane was carefully removed to avoid any breakage or contamination. 7. The nitrocellulose membrane was dried and slots cut out according to the loading positions. 8. Nucleic acids retained on the membrane were measured by scintillation counting. 9. Nucleic acid binding was quantified according to nucleic acid retention as a function of the addition of NDH II. This provided an estimate of the nucleic acid binding constant as well as the binding length of NDH II (Fig. 21.2). a)

b)

Fig. 21.2. Nitrocellulose binding assay of NDH II. (a) Binding of NDH II to poly(rI)(rC). (b) Determination of the association constant of NDH II according to the binding data (11).

3.4. Gel Shift by Agarose Gel Electrophoresis

1. 1.5% agarose in TBE was melted in a microwave oven and cooled down to about 60C (a hot but hand endurable temperature). The agarose was poured into a 2-mm-thick space between two glass plates (20 20 cm), which can be slightly opened to facilitate filling. The comb was inserted at the top of the gel to a depth just enough for holding the sample volume (see Note 3). 2. A labeled M13 ssDNA probe as applied for the filter binding assay was used for the agarose gel shift assay.

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3. 6 mM (nucleotides) M13 ssDNA were incubated for 20 min at room temperature with increasing amount (0–280 nM) of NDH II in 10 ml binding buffer as used for the filter binding assay. To measure the binding specificity, increasing amounts of nucleic acid competitors (1.5–15 mM nucleotides) were added. 4. At the end of incubation time samples were mixed with 3 ml of loading buffer and loaded to the agarose gel prepared in 1 TBE buffer. 5. Electrophoresis was performed in 1 TBE buffer at a low and constant voltage (10–30 V) for a period of 16–18 h (see Note 5). 6. Electrophoresis was stopped when the bromophenol blue dye reached the bottom of the agarose gel. 7. One glass plate was carefully removed to avoid damage of the agarose, which together with the other glass plate was packed into Saran wrap and then exposed to an X-ray film overnight at –80C (Fig. 21.3).

Fig. 21.3. Determination of single strand nucleic acids binding of NDH II by agarose gel electrophoresis. Binding of NDH II to ssDNA was competed by ssRNA but not by dsRNA (17).

3.5. Northwestern Blotting

1. 5–10 ml of bacterial lysates containing GST-fusion proteins were separated through a 10% SDS polyacrylamide gel. 2. After electrophoresis the proteins were transferred to a HybondC nitrocellulose membrane using a semi-dry blotter, which was assembled by placing in an order from the cathode (–) to the anode (+) three sheets of Whatman 3MM paper, the protein gel, the Hybond-C nitrocellulose membrane, and three sheets of Whatman 3MM paper. 3. Electrotransfer proceeded for 1 h at a current of 0.8 mA per cm2. 4. After electrotransfer, the nitrocellulose membrane was removed from the semi-dry blotter and stained with Ponceau S solution in a culture dish for a few minutes. The stained protein on the membrane was photographed.

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5. The nitrocellulose membrane was washed with a few changes of TBS to remove the dye. 6. The membrane was placed into 8 M urea in TBS for 10 min, followed by ten steps of dilutions with TBS, i.e., by replacing after every 10 min one-third of the volume of the previous urea-containing TBS with TBS buffer without urea. During this procedure the protein undergoes partial renaturation. 7. After urea removal the membrane was blocked with 5% milk powder in binding buffer for 1 h at room temperature to saturate unspecific binding sites. 8. The membrane was incubated for at least 30 min with binding buffer in a minimum volume (10 ml) containing a nucleic acid probe, e.g., 50 -labeled poly(rI) or poly(rI)(rC). 9. After binding, the radioactive solution was removed and the membrane was washed three times for about 5 min each with binding buffer (see Note 7). 10. After washing the membrane was packed into Saran wrap and exposed to X-ray film for visualizing the nucleic acid binding signals (Fig. 21.4).

Fig. 21.4. Characterization of the nucleic acid binding domains of NDH II by Northwestern blots. (a) dsRNA binding of the N-terminal dsRBD I (aa 1–130), dsRBD II (aa 131–318) and dsRBD I+II (aa 1–318) of NDH II. Apparently, dsRBD II displayed a higher affinity to dsRNA than dsRBD I while dsRBD I + II bound co-operatively, i.e., binding to dsRBD I + II was stronger than that of either of the individual domain (2). (b) dsRNA binding of the three dsRBD II mutants F211A, K235A, and K236A (unpublished data). The K235A mutant completely abolished dsRNA binding. Note that the band at 66 kDa represents unspecific binding.

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4. Notes 1. Repeated freezing and thawing cycles of the oligonucleotide may lead to a serious decrease in the efficiency of 50 -labeling and thus the sensitivity of the assay. Once this occurs, the oligonucleotide should be discarded and replaced by a new batch. 2. The described processing of the nitrocellulose membrane is necessary for an improvement of its flow rate. Without this pre-treatment a high background level and/or a variation of the retained nucleic acid can arise that might give wrong positive results. 3. A too cold agarose solution would be difficult to cast between the glass plates for vertical electrophoresis. Moreover, the well depth should be carefully controlled because the gel may be disrupted when a too deeply inserted comb is removed. 4. The storage buffer of NDH II contains 50% glycerol. Glycerol may inhibit the helicase activity when exceeding 10% in the assay mixture. 5. No attempt should be made to accelerate the speed of the gel shift by increasing the voltage for electrophoresis because an increased electric field might disrupt the protein–nucleic acid complex by denaturing the protein at temperatures above 37C. 6. Blocking buffer should always be prepared immediately before use because prolonged storage of this solution might lead to bacterial growth and, as a result contamination with bacterial nucleases that destroy the nucleic acid probe. 7. Too extensive washing may lead to a complete elimination of the radioactive signal whereas insufficient washing causes a higher background. This problem may be avoided by closely monitoring the radioactivity during the washing procedure using a Geiger counter so that the time to stop washing, usually judged by experience, can be adjusted.

Acknowledgments The work was supported by Deutsche Forschungsgemeinschaft Grant Gr 895/5-2. We are indebted to H. Pospiech for his critical comments.

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References 1. Fuller-Pace F. V. (2006) DExD/H box RNA helicases: multifunctional proteins with important roles in transcriptional regulation. Nucleic Acids Res. 34, 4206–4215. 2. Zhang S. and Grosse F. (1997) Domain structure of human nuclear DNA helicase II (RNA helicase A). J. Biol. Chem. 272, 11487–11494. 3. Zhang S. and Grosse F. (2004) Multiple functions of nuclear DNA helicase II (RNA helicase A) in nucleic acid metabolism (review). Acta Biochim. Biophys. Sin. (Shanghai) 36, 177–183. 4. Takeda Y., Caudell P., Grady G., Wang G., Suwa A., Sharp G. C., Dynan W. S., and Hardin J. A. (1999) Human RNA helicase A is a lupus autoantigen that is cleaved during apoptosis. J. Immunol. 163, 6269–6274. 5. Toretsky J. A., Erkizan V., Levenson A., Abaan O. D., Parvin J. D., Cripe T. P., Rice A. M., Lee S. B., and Uren A. (2006) Oncoprotein EWS-FLI1 activity is enhanced by RNA helicase A. Cancer Res. 66, 5574–5581. 6. Schlegel B. P., Starita L. M., and Parvin J. D. (2003) Overexpression of a protein fragment of RNA helicase A causes inhibition of endogenous BRCA1 function and defects in ploidy and cytokinesis in mammary epithelial cells. Oncogene 22, 983–991. 7. Mischo H. E., Hemmerich P., Grosse F., and Zhang S. (2005) Actinomycin D induces histone gH2AX foci and complex formation of gH2AX with Ku70 and nuclear DNA helicase II. J. Biol. Chem. 280, 9586–9594. 8. Friedemann J., Grosse F., and Zhang S. (2005) Nuclear DNA helicase II (RNA helicase A) interacts and stimulates the

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exonuclease of Werner syndrome helicase (WRN). J. Biol. Chem. 280, 31303–31313. Hanada K. and Hickson I. D. (2007) Molecular genetics of RecQ helicase disorders. Cell Mol. Life Sci. 64, 2306–2322. Lee C.-G., Chang K. A, Kuroda M. I., and Hurwitz J. (1997) The NTPase/helicase activities of Drosophila maleless, an essential factor in dosage compensation. EMBO J. 16, 2671–2681. Zhang S. and Grosse F. (1994) Nuclear DNA helicase II unwinds both DNA and RNA. Biochemistry 33, 3906–3912. ZhangS.andGrosseF.(1991)Purificationand characterization of two DNA helicases from calfthymus.J.Biol.Chem.266,20483–20490. Fenner B. J., Goh W., and Kwang J. (2006) Sequestration and protection of doublestranded RNA by the betanodavirus b2 protein. J. Virol. 80, 6822–6833. Sambrook J., Fritsch E. F., and Maniatis T. (1989), in Molecular Cloning: A Laboratory Manual, Cold Spring Harbor Laboratories, Cold Spring Harbor, NY. Harlow E., and Lane D. (1999), in Using Antibodies: A Laboratory Manual, Cold Spring Harbor Laboratories, Cold Spring Harbor, NY. Reckmann B., Grosse F., and Krauss G. (1983) The elongation of mismatched primers by DNA polymerase from calf thymus. Nucleic Acids Res. 11, 7251–7260. Zhang S., Herrmann C. and Grosse F. (1999) Pre-mRNA and mRNA binding of human nuclear DNA helicase II (RNA helicase A). J. Cell Sci. 112, 1055–1064.

Chapter 22 Regulation of Inter- and Intramolecular Interaction of RNA, DNA, and Proteins by MLE Hyangyee Oh, Andrew M. Parrott, Yongkyu Park, and Chee-Gun Lee Abstract Drosophila maleless (MLE) is a member of helicase superfamily 2 and functions as a dosage compensation factor essential for the development of male flies. This function provides a good opportunity to investigate diverse biochemical activities associated with MLE in the context of a defined in vivo pathway, i.e., the transcriptional activation of X-linked genes. We have shown for the first time that MLE catalyzes the unwinding of both DNA and RNA and that MLE helicase activity is essential for its in vivo function. Also, we have provided evidence that MLE stimulates the transcriptional activity of roX2 on the X chromosome. We have also found that MLE interacts with dsDNA, topoisomerase II, and nucleosome. This observation supports a current model of dosage compensation: transcriptional activation of X-linked genes is causally associated with conformational change in the male X chromosome, subsequent to the targeted association of the dosage compensation complex (DCC). Key words: Maleless (MLE), roX2, dosage compensation, topoisomerase II.

1. Introduction In Drosophila, dosage compensation is achieved by transcriptional activation of X-linked genes (1). To date, genetic analysis has identified eight transacting factors that are necessary for the onset or maintenance of dosage compensation. Based on observations that loss of their functional allele leads to a male-specific lethal phenotype, these factors are named the MSL (male-specific lethal) proteins. They include six protein factors such as MSL1 (2), MSL2 (3, 4), MSL3 (5), MLE (maleless) (6), MOF (male-absent on the first) (7), and JIL-1 (8) and two non-coding RNAs, roX1 and roX2 (RNA on X) (9, 10). M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_22, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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It is generally believed that dosage compensation occurs in several discernible steps, starting with the targeted association of roX RNAs and MSL complex with the X chromosome. Upon the expression of MSL2 protein, MSL proteins assemble a complex on approximately 35 sites of the X chromosome, called ‘‘chromatin entry sites’’ (11–14). Once the chromatin entry sites are fully occupied with the MSL complex in the presence of MLE, the flanking chromatin region becomes competent in binding the MSL complex (15, 16). This ‘‘spreading’’ or ‘‘nucleation’’ process appears to require the histone acetyltransferase (HAT) activity of MOF (17). We have shown that mleGET, a mutant MLE defective in ATPase activity, leads to a poor association of the MSL complex with the male X chromosome and lethality in male embryos (18). In addition, MLE is also involved in the transcriptional activation of roX2 and its subsequent association with MSL complexes (17, 19). Detailed mechanistic understanding of dosage compensation has greatly advanced due to the employment of diverse biochemical assays. For example, dsRNA/dsDNA unwinding assays, ATPase, nucleotide UV-crosslinking, immunoprecipitation coupled with RT-PCR, and multiplex PCR and reporter gene assays have yielded important clues to the molecular basis of how ATP-dependent and ATP-independent activities of MLE contribute to dosage compensation.

2. Materials 2.1. Expression and Purification of Recombinant MLE

1. Sf9 cells (Invitrogen). 2. Grace’s Medium (Invitrogen) supplemented with 10% fetal bovine serum (FBS, Sigma), 1% Pluronic F-68 solution (Sigma-Aldrich), and 1% penicillin-streptomycin (SigmaAldrich). 3. Chromatography resins: Pro-Bond1 (Invitrogen) and hydroxyapatite (Bio-Rad). 4. Hypotonic buffer: 50 mM Tris–HCl, pH 8.0, 10 mM KCl, 1.5 mM MgCl2, 20 mM Na2HPO4, pH 7.4, and 0.5 mM PMSF (phenyl methylsulfonate fluoride, Sigma-Aldrich). 5. Nuclei re-suspending (NS) buffer: 50 mM Tris–HCl, pH 8.0, 1.5 mM MgCl2, 20 mM Na2HPO4, pH 7.4, and 0.5 mM PMSF. 6. Column wash (CW) buffer: 20 mM sodium phosphate, pH 6.0, 0.5 M NaCl, and 0.5 mM PMSF. 7. Buffer P20: 20 mM sodium phosphate, pH 7.4, 0.5 mM PMSF, 0.05% Nonidet P-40 (Bufferad), and 10% glycerol.

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8. Buffer P250: 0.25 M sodium phosphate, pH 7.4, 0.5 mM PMSF, 0.05% Nonidet P-40, and 10% glycerol. 9. Buffer A250: 20 mM HEPES–NaOH, pH 7.4, 0.1 mM EDTA, 0.5 mM PMSF, 2 mM dithiothreitol (DTT), 0.25 M NaCl, and 12.5% glycerol. 10. Protease inhibitors: All reagents are from Sigma-Aldrich. (a) 0.5 M EGTA (1,000 ); store at room temperature. (b) A mixture of leupeptin (10 mg/ml) and aprotinin (10 mg/ml) in de-ionized H2O (500 ); store at –20C. (c) Pepstatin (1 mg/ml) in ethanol (100 ); store at –20C. 11. Dialysis tubing: Spectrapor1 membrane (Spectrum) (MWCO 6-8000). (a) Membranes are cut into pieces (15 cm) and autoclave in a beaker (2 l) containing 2% sodium bicarbonate and 1 mM EDTA for 10 min at 10 lb/in2. (b) Wash extensively with de-ionized H2O and repeat the autoclave. (c) After cooling, store the tubing at 4C. 12. Acrylamide:bis (30:1) stock solution: dissolve 30 g acrylamide (Invitrogen) and 1 g bis-N,N’-methylene-bis-acrylamide (Bio-Rad) in 100 ml of de-ionized H2O. Filter the solution through a 0.45-mm filter (Nalgene) and store it at 4C. 13. SDS-polyacrylamide gel: gels (7–12.5%) are prepared using a 30% acrylamide:bis (30:1) stock solution and 4 separation buffer (0.5 M Tris–HCl, pH 8.8, 0.4% SDS), whereas a stacking gel (4% acrylamide) is prepared using 4 stacking buffer (0.5 M Tris–HCl, pH 6.8, 0.4% SDS). 14. SDS sample buffer (5 ): 0.1 M Tris–HCl (pH 6.8), 0.5% SDS, 0.05% bromophenol blue (BPB), and 50% glycerol. 15. SDS-PAGE running buffer (10 ): dissolve 30.2 g Tris base, 144 g glycine, and 10 g SDS per liter. (It is not necessary to adjust the pH of the stock solution.) 2.2. Preparation of ssRNA and dsRNA

1. Plasmid DNAs: pSP65 (Promega), pGEM1 (Promega), pGEM3 (Promega), and pGEM3CS– (20). 2. Enzymes: SP6 RNA polymerase (Promega), T7 RNA polymerase (Promega), RNasin (Promega), RNase P1 (Boehringer Mannheim), U2AF (21), topoisomerase II (22) (Vaxron), and restriction endonucleases (New England BioLabs, NEB) such as PvuII, HaeII, RsaI, XbaI, and AccI. 3. Light box, permanent marker, and disposable micropestles (Eppendorf). 4. Circular glass fiber filter paper (25 mm diameter): GF/C (Enzo).

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5. Trichloroacetic acid (Fisher): prepare 10 and 1% solutions (w/v) in de-ionized H2O and store at 4C. 6. Carrier DNA solution: dissolve 25 mg salmon sperm DNA (Sigma-Aldrich) in 50 ml of 5 mM sodium pyrophosphate (pH 7.0), sonicate 6–8 times for 15 s using a microtip probe on ice until it gets pipettable, and then store at 4C. 7. Formamide dye mix (2 ): to 10 ml formamide, dissolve 10 mg xylene cyanol FF (XC) and 10 mg BPB. 8. Native dye mix (5 ): 0.1 M Tris–HCl (pH 7.4), 0.05% XC, 0.05% BPB, 0.1% Nonidet P-40, and 50% glycerol. 9. Denaturing polyacrylamide gel (about 18 20 cm): using 30% acrylamide:bis stock solution and 5 TBE, prepare a 10% denaturing gel containing 50% urea and 0.5 TBE. 10. Native polyacrylamide gel (about 18 20 cm): using 30% acrylamide:bis stock solution and 5 TBE, prepare an 8% native gel containing 0.5 TBE. 11. Electrophoresis mobility shift assay (EMSA): (a) Prepare 5–10% acrylamide/5% glycerol composite gel about 1.5 h before use, using a 30% acrylamide stock solution (see Section 2.1, step 12), 50% glycerol (v/v), and 5 TBE buffer. (b) Pre-run the gel for at least 30 min at constant 20 mA. 12. Load the reaction mixture, continue the electrophoresis until BPB migrates about two-thirds of the gel. 13. DNA/RNA hybridization buffer: 50 mM Tris–HCl (pH 8.0), 0.5 M NaCl, 0.1 mM EDTA, and 0.5% SDS. 14. Extraction buffer (2 ): 1 M ammonium acetate, 10 mM EDTA, and 0.2% SDS. 15. RNA/DNA dilution buffer: 20 mM HEPES–NaOH (pH 7.4), 50 mM KCl, 0.5 mM EDTA, 0.01% Nonidet P-40, and RNasin (Promega) (250 units/ml). 16. TBE (5 ): per liter, add 54 g Tris base, 27.5 g boric acid, and 20 ml 0.5 M EDTA (pH 8.0). 17. Reaction termination buffer (5 ): 0.1 M Tris–HCl (pH 7.4), 50 mM EDTA, 0.1% BPB, 0.1% XC, and 0.1% NP-40, 0.5% SDS, and 50% glycerol. 2.3. Preparation of dsDNA

1. Plasmid DNA: pBluescriptIIKS– (Stratagene). 2. Enzymes: T4 polynucleotide kinase and Klenow fragment (NEB); Taq DNA polymerase (NEB). 3. -32P-dCTP and g-32P-ATP: Specific radioactivity, 3,000 Ci/ mmol.

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4. 1 TBS: 50 mM Tris-–HCl, pH 8.0, 150 mM NaCl, and 0.1 mM EDTA. 5. Sephadex G-50 (Amersham Biosciences) in 1 TBS. 6. Duplex DNAs for the helicase assay are prepared using the following synthetic DNAs dissolved in de-ionized H2O at 100 nM: (a) 98-mer; 50 -GAATA CAAGC TTGGG CTGCA GGTCG ACTCT AGAGG ATCCC CGGGC GAGCT CGAAT TCGGG TCTCC CTATA GTGAG TCGTA TTAAT TTCGA TAAGC CAG-30 (b) 38-mer; 50 -GAATA CACGG AATTC GAGCT CGCCC GGGGA TCCTC TAG-30 (c) 5/30-mer; 50 -AGAGT CGACC TGCAG CCCAA GCTTG TATTC-30 (d) 5COM/30-mer; 50 -GAATA CAAGC TTGGG CTGCA GGTCG ACTCT-30 (e) 3/30-mer; 50 -CTGGC CGACT CATCA-30

TTATC

GAAAT

TAATA

7. For the preparation of dsDNA substrate for EMSA, prepare the following DNA primers: (a) T3 and T7 primers (Stratagene). (b) ProxA; 50 -TTGGA ATCCC GCTAT TTTCG GATTC ATGCA GTTCC CATTA TATTT TATTC GGTA-30 . (c) ProxB; 50 -TACCG AATAA AATAT AATGG GAACT GCATG AATCC GAAAA TAGCG GGA-30 . 2.4. ATPase Assay

1. g-32P-ATP (MP 3,000 Ci/mmol.

Biochemicals):

specific

radioactivity,

2. RNAs: Yeast tRNA (Roche) and RNA homopolymers such as poly(A), poly(U), poly(G), and poly(C) (GE Healthcare) are dissolved in DEPC-treated H2O at 10 mg/ml and stored in aliquots at –20C. 3. DNA homopolymers: Poly(dA), poly(dT), poly(dC), and poly(dG) (GE Healthcare) are dissolved in de-ionized H2O (10 mg/ml) and stored in aliquots at –20C. 4. Thin layer chromatography (TLC) plate: cellulose PEI-F plate (J.T. Baker). 5. TLC buffer: 1 M formic acid and 0.5 M LiCl. 6. TLC developing tank with lid (27 7 25 cm) (Kontes). 2.5. Nucleotide UVCrosslinking and Western Blotting

1. -32P-GTP (MP 3,000 Ci/mmol.

Biochemicals):

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radioactivity,

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2. 96-well plate (round bottom): Flexible and untreated PVC (Costar). 3. Western transfer buffer (1 ): per liter, dissolve 5.8 g Tris base, 29 g glycine, 1 g SDS, and 200 ml methanol. 4. PVDF filter: Immobilon-P (Millipore). Cut the filter into appropriate sizes and submerge it in methanol for 10 min followed by Western transfer buffer for additional 10 min. 5. Blocking solution: 5% non-fat dry milk (Carnation) in 1 PBST. 6. Secondary antibody: anti-mouse and anti-rabbit IgGs conjugated with horseradish peroxidase (Amersham Biosciences). 7. Enhanced chemiluminescent (ECL) reagents (Amersham Biosciences) and Bio-Max ML film (Kodak). 8. DE-81 and Whatman-3 M paper (Whatman). 2.6. Immunoprecipitation and RT-PCR Analysis

1. 1 Extraction buffer: prepare a fresh buffer consisting of 20 mM HEPES–NaOH (pH 7.6), 70 mM KCl, 2 mM DTT, 0.1% NP-40, 1 mM PMSF, and 1 unit/ml RNasin (Promega), and protease inhibitor (1 ) (see Section 2.1, step 10). 2. Polyclonal antibodies: rabbit pre-immune sera and polyclonal antibodies specific for MLE or MSL1 (13). 3. Protein A-Sepharose (50% slurry) (Sigma-Aldrich). 4. Lysis buffer for total RNA isolation: Ultraspec reagent (Biotecx). 5. RNase-free DNase RQI (1 unit/ml) and 10 DNase I buffer (Promega). 6. SuperScript II RT and 10 Advantage II PCR buffer (Clontech). 7. Taq DNA polymerase (NEB). 8. 100 mM dNTP set (Amersham Biosciences). 9. PCR primers roX2-5/rox2-3; 50 -TTGGC ATTTT GCTCT TGTTT TTCTC-30 /50 -CGTTA CTCTT GCTTC ATTTT GCTTC G-30 .

2.7. Multiplex PCR and Reporter Gene Assays

1. Expression vectors: pMT/V5-b-galactosidase (Invitrogen), pMT-MLE (19), and pMT-MSL1 (19). 2. A reporter construct: Prox2-luciferase (19). 3. S2 cells (Invitrogen): cells are grown in Drosophila-SFM (Invitrogen).

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4. Transfection reagents: Prepare 2 M CaCl2 and 2 HBS (50 mM HEPES–NaOH, pH 7.05, 1.5 mM Na2HPO4.2H2O, 280 mM NaCl, 10 mM KCl, and 12 mM dextrose) (adjust pH with 1 N HCl or 1 N NaOH if necessary) and store them in aliquots at –20C. 5. Copper sulfate solution (100 ): Prepare 100 mM solution and store it at room temperature. 6. 1 luciferase lysis buffer: 50 mM potassium phosphate, pH 7.4, 0.2% Triton X-100, 2 mM DTT, and 5 mM b-glycerophosphate. 7. Spin column (Qiagen). 8. PCR primer sets: Prepare the following primer sets (100 mM) in TE buffer: (a) MSL1-5/MSL1-3; 50 -GAAGA TCTAT GAGCG CCA30 /50 -CTGCT TTAAT TCCTC ATTCT GCG-30 . (b) DSRPK1-3/dSRPK11-5; 50 -GATGA ATGCA ACGTC CACGT AAAG-30 /50 -CATCC TTTTG CGACC ACTCG TAC-30 . (c) MLE-5/MLE-3; 50 -CAAAA CCTCG GTGAA TTGCA GCAA-30 /50 -CTGAT CCTCT ATTGC TTTCA-30 . (d) RoX2-5/rox2-3 (see Section 2.6, step 9).

3. Methods Diverse in vivo and in vitro assays make possible investigations into the biochemical basis by which MLE brings about a twofold increase in the transcription activity of numerous genes on the male X chromosome. For example, by employing recombinant MLE, it can be determined whether the in vivo interaction between MLE and roX2 RNA is dependent on the ATPase or helicase activity of MLE, and whether the conformation of bound roX2 RNA is subject to MLE helicase activity. Conversely, how the association of roX2 influences the ATPase and helicase activities of MLE or its interaction with specific DNA (19) or with non-specific DNA (23) can be explored. Once the minimal roX2 RNA binding region is identified, in vivo reporter gene assays will enable us to address the above issues from the aspect of transcription regulation of X-linked genes, an in vivo context that mimicks dosage compensation.

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3.1. Expression and Purification of Recombinant MLE Using Baculoviruses

1. Sf9 cells, in exponential growth phase, are centrifuged at 800 g for 15 min and are re-suspended in fresh Grace’s medium to a density of 2 106 cells/ml. About 1 109 Sf9 cells in 0.5-l medium are needed for a reproducible and reliable yield of recombinant MLE. 2. Infect Sf9 cells with baculoviruses at 5.0 MOI (multiplicity of infection) in a 2-l culture flask, and incubate them in a refrigerated incubator at 27C, while stirring at 100 rpm. 3. Two days later, cells are centrifuged at 800 g for 15 min and washed once with ice-cold 1 PBS containing 1 protease inhibitors. After centrifugation, decant the supernatant and measure the packed cell volume (PCV), which is typically approximately 5–6 ml. All subsequent purification steps should be processed at 4C. 4. Re-suspend Sf9 cells in 2.5 PCV of ice-cold hypotonic buffer supplemented with 1 protease inhibitors. 5. After incubation for 15 min on ice, cells are disrupted by homogenization: 10 strokes using a type B pestle. 6. Homogenate is cleared by centrifugation at 2,000 g for 10 min. Pellet, containing nuclei, is re-suspended in 1.5 PCV of NS buffer with 1 protease inhibitors. 7. Quickly add 1/10 volume of 5 M NaCl to nuclear and cytoplasmic fractions, and incubate them on a nutator for 20 min. 8. Clear the mixtures by centrifugation at 15,000 g for 30 min. 9. Pool the clear supernatants in a tube containing Pro-Bond1 resin (4 ml) pre-equilibrated with NS buffer containing 0.5 M NaCl. 10. After overnight incubation on a nutator, pack the resin into a column, and wash the column resin with 60 ml of CW buffer. 11. Bound proteins, including MLE, are eluted with a linear gradient (50 ml) of 0–0.5 M imidazole in CW buffer in a total of 40 fractions (1.2 ml/fraction). 12. Aliquots (5 ml) of fractions are subject to a 7.5% SDS-PAGE (Fig. 22.1), and fractions enriched in MLE are pooled and dialyzed overnight against 2 l of buffer P20 containing 20% glycerol and 1 mM DTT. 13. The following day, pooled fractions are loaded onto a hydroxyapatite column (3 ml) equilibrated with buffer P20 containing 0.2 M NaCl. 14. Bound proteins are eluted with a linear gradient (60 ml) of 20–250 mM Na phosphate, pH 7.4, using buffer P20 and P250 containing 0.25 M NaCl, in a total of 60 fractions.

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Fig. 22.1. Purification of hexahistidine-tagged MLE using a Ni+-conjugated Pro-Bond column. Clear cell lyate, prepared from Sf9 cells infected with baculoviruses, was loaded onto the column, and bound proteins were eluted with a linear gradient of imidazole (0–0.5 M) as described in Section 3.1, step 11. Aliquots (5 ml) of the following fractions were resolved on a 7.5% SDS-PAGE, and proteins were visualized by Coomassie blue staining. Hexahistidine-tagged MLE (145 kDa) is indicated by arrowhead. L, clear cell lysate; FT, flow-through fraction; W1, supernatant fraction obtained from the first batchwise wash; W2, supernatant fraction obtained from the second batch-wise wash; 3–33, fractions eluted with the indicated imidazole gradient.

15. Aliquots (10 ml) of fractions are resolved on a 7.5% SDSPAGE, and fractions enriched in MLE are pooled, dialyzed against 2 l of buffer A250, and stored in aliquots at –70C. 3.2. Preparation of ssRNA and dsRNA

1. A standard helicase substrate, a partial dsRNA, is prepared using two in vitro transcription reactions. The first reaction (100 ml), which is for 32P-labeled 38-mer RNA, is set up with 10 ml of 10 reaction buffer (provided by Manufacturer), 5 mM DTT, 20 units RNasin, 2.5 mg pSP65 cut with XbaI, 0.5 mM ATP, 0.5 mM CTP, 0.5 mM UTP, 50 mM GTP, and 40.7 ml -32P-GTP. The second reaction, for a longer 98-mer RNA, is assembled essentially the same way except it is provided with 2.5 mg pGEM1 cut with PvuII, 0.5 mM GTP, and 0.5 ml -32P-GTP (see Fig. 22.2 and Note 1). 2. After incubation for 100 min at 37C, reactions are terminated by the addition of 100 ml of 1 RNA/DNA extraction buffer, which is followed by conventional phenol/chloroform and chloroform extraction. 3. To the aqueous phase recovered in step 2, add glycogen (20 mg) and 2.5 volume of ice-cold ethanol, mix well, and place in powdered dry-ice. 4. After 5 min incubation (see Note 2), centrifuge the mixture at 12,000 g for 15 min at 4C and discard the supernatant using a pipette.

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Fig. 22.2. Preparation of dsRNA and dsDNA substrates. (A) DNA templates and RNA polymerases necessary for the preparation of dsRNA substrates are summarized relative to upper or lower RNA strand constituting each dsRNA. For example, 52-nt long upper strand of 50 -tailed dsRNA is synthesized in a transcription reaction provided with pGEM3SacI and SP6 RNA polymerase, whereas a 46-nt long lower strand is prepared using pGEM3-AccI and T7 RNA polymerase. RNA polymerases used are indicated in parenthesis. (B) An autoradiogram shows the position of indicated dsRNA and dsDNA substrate relative to xylene cyanol (XC) and bromophenol blue (BPB) on an 8% native polyacrylamide gel (30:1). In lanes 1–2, the sequence and structure of dsRNA and dsDNA are same as RNA-a in Fig. 22.2A. Flushed-ended dsRNA (29 bp) (lane 3) and dsDNA (30 bp) (lane 4) are described in Sections 3.2 and 3.3, respectively.

5. Wash the pellet with ice-cold 70% ethanol and repeat the centrifugation. 6. Air-dry the pellet, and dissolve it in 10 ml of formamide dye mix by incubating for at least 15 min at 37C. 7. After heating 5 min at 95C, resolve RNA on a 10% denaturing polyacrylamide gel.

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8. When BPB migrates about two-thirds of the gel, remove the top glass plate and cover the gel with plastic wrap on top of the bottom glass plate. 9. Inside a darkroom, overlay the covered gel on an X-ray film. 10. After 1–2 min exposure, develop X-ray film. 11. Place the developed X-ray film on a light box, overlay the gel with the glass plate facing the X-ray film, and mark the gel area containing RNA. 12. Cut out the marked gel area and transfer it to a microcentrifuge tube containing 0.4 ml of 1 RNA/DNA extraction buffer. 13. Thoroughly grind the gel piece using a micropestle, and incubate it on a nutator for 2 h at room temperature. 14. Centrifuge at 12,000 g for 5 min, and transfer the supernatant into a new tube. 15. Isolate 32P-labeled RNA following the procedures described in steps 2–5. 16. Dissolve the RNA pellets as follows. (a) To prepare ssRNA, dissolve the 38-mer RNA in 100 ml DNA/RNA dilution buffer, and go to step 17. (b) To prepare dsRNA, dissolve 38-mer and 98-mer RNAs in 50 ml DNA/RNA hybridization buffer and go to step 18. 17. Measure the radioactivity with 1 ml aliquot using the liquid scintillation counter. By adding extra volume of DNA/RNA dilution buffer, adjust the specific radioactivity of ssRNA solution to be either 50,000 cpm/ml (equivalent to 25 fmol ssRNA/ml) or 100,000 cpm/ml, and store 38-mer RNA in aliquots at –20C. 18. Combine 38-mer RNA with 98-mer RNA, and using a PCR instrument, incubate the mixture as follows: 95C for 5 min, 75C for 1 h, 68C for 3 h, 55C for 0.5 h, 45C for 0.5 h, 37C for 0.5 h, and 25C for 10 min. 19. Add equal volume of 2 DNA/RNA extraction buffer, adjust the final volume of the sample to 400 ml with 1 DNA/RNA extraction buffer, and isolate RNAs following the procedures described in steps 3–5. (a) To prepare a partial dsRNA (Fig. 22.2A), dissolve the RNA pellet in 10 ml of 1 Native dye mix and go to step 20. (b) To prepare flush-ended dsRNA, re-suspend the RNA pellet in 100 ml of DNA/RNA dilution buffer and go to step 22. 20. After pre-running an 8% native polyacrylamide gel for 30 min at 20 mA, load the RNA sample, and then follow the procedures described in steps 8–15 (Fig. 22.2B).

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21. The dsRNA is re-dissolved in 100 ml DNA/RNA dilution buffer and store them after adjusting the specific radioactivity to be either 50,000 or 100,000 cpm/ml as described in step 17. 22. Add 5 mg of RNase P (1 mg/ml), incubate at 37C for 90 min, and re-isolate the flush-ended dsRNA following the procedure described in step 19 (Fig. 22.2B). 23. Re-dissolve the flush-ended dsRNA as described in step 17. 3.3. Preparation of dsDNA Substrates for the Helicase Assay

1. Set up end-labeling reaction (25 ml) as follows: 2.5 ml 10 polynucleotide T4 Kinase buffer (provided by NEB), 0.5 ml (10 units) T4 polynucleotide kinase, 1 ml 0.1 M DTT, and 25 pmol of a synthetic DNA oligomer (38-mer, 5/30-mer, or 3/30-mer described in Section 2.3, step 6). 2. After 30 min incubation at 37C, continue the incubation for an additional 30 min at 65C and subsequently transfer the reaction mixture on ice. 3. Determine the labeling efficiency as follows: (a) Take 0.5 ml aliquot of each reaction and mix it with 100 ml carrier DNA solution in a new tube on ice. (b) Fill-up the tube with 10% TCA, and incubate it on ice for 20 min, during which, GF/C filter is thoroughly hydrated with de-ionized H2O. (c) Place the GF/C filter on meshed filtration funnel. (d) While applying a mild vacuum, collect the DNA precipitate by filtering the mixture through GF/C filter. (e) Wash the filter with 10 ml 1% TCA and 10 ml 95% ethanol. (f) Dry the filter and measure the radioactivity. (g) Calculate the specific radioactivity of DNA using the following formula: Specific radioactivity per 50 fmol of labeled DNA = Observed cpm 0.1. (h) By adding unlabeled DNA, adjust the specific radioactivity to be either 50,000 or 100,000 cpm per 50 fmol of ssDNA. 4. To the above reaction mixture, add 5 ml 1 M Tris–HCl, pH 8.0, 10 ml 5 M NaCl, 1 ml 10% SDS, 0.2 ml 0.5 M EDTA, and unlabeled complementary DNA as follows: (a) To prepare a partial dsDNA, 30 -tailed or 50 -tailed dsDNA (Fig. 22.3), add 200 pmol 98-mer ssDNA to the reaction mixture containing 32P-labeled 38-mer, 3/30-mer, or 5/ 30-mer DNA, respectively. (b) To prepare a flushed-ended dsDNA, add 200 pmol 5COM/30-mer to the reaction mixture containing 32 P-labeled 5/30-mer.

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Fig. 22.3. dsDNA substrates used for the determination of the directionality of helicasecatalyzed unwinding reaction. Helicase reactions were carried out with four different DNA substrates (A–D), as described in Section 3.8. Lane 1, DNA substrate alone; lane 2, boiled substrate; lane 3, 10 ng MLE without ATP; lanes 4–6, 2.5, 5, and 10 ng MLE with ATP, respectively.

5. After adjusting the final volume to 200 ml, the mixture is subject to hybridization reaction as described in step 18 of Section 3.2, and dsDNA formed is isolated following the procedures described in steps 19–21 of Section 3.2 (Figs. 22.2b and 22.3). 3.4. Preparation of Non-specific dsDNA Probes for EMSA

1. Six to ten standard PCR reactions (50 ml each), containing 1 ng pBS, 50 pmol T3 primer, and 50 pmol T7 primer, are incubated at 95C for 3 min, followed by 35 cycles of (95C for 30 s, 55C for 1 min, and 72C for 2 min). 2. Combine all reactions in one microcentrifuge tube and isolate PCR product by conventional phenol/chloform and chloroform extraction followed by ethanol precipitation. 3. Dissolve the DNA pellet in 50 ml TE buffer, re-purify the DNA using a spin column (Qiagen), and determine the DNA concentration using a UV spectrometer (1.0 OD260 equals 50 mg/ml). 4. Treat 1.1 mg (10 pmol) DNA with ClaI, (EcoRI + NotI) or (EcoRI + XbaI) to prepare a mixture of dsDNAs of varying length such as (65 þ 104 bp), (41 þ 50 þ 84 bp), or (34 þ 57 þ 84 bp) DNAs, respectively (Fig. 22.4A). Subsequently, re-purify the mixture of DNA fragments following the procedure described in steps 19 and 20 of Section 3.2, and re-suspend in 10 ml TE buffer.

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Fig. 22.4. Coupling of PCR with end-filling reaction to prepare dsDNA substrates. (A) A 167-bp long dsDNA was prepared by PCR using pBluescriptIIKS– and T3/T7 primers. Subsequent to digestion with the indicated restriction enzymes, end-filling reactions, provided with Klenow fragment and deoxynucleotides, yield radiolabeled dsDNAs, shown as filled-bars. (B) Two major 32P-labeled DNAs of predicted length, obtained from each end-filling reaction, were detected by autoradiography subsequent to electrophoresis on an 8% native polyacrylamide gel (30:1).

5. End-label the mixture of DNA fragments (10 pmol) in a fill-in reaction (50 ml), containing 5 ml 10 reaction buffer (provided by manufacturer), 2 ml 2 mM dATP/dCTP/dGTP, 15 ml (50 pmol) -32P-dCTP, 10 units Klenow enzyme, and 10 pmol DNA. After 30 min incubation at room temperature, add 2 ml 0.5 M EDTA and transfer on ice. 6. Determine the specific radioactivity of end-labeled DNA by TCA precipitation following the procedure described in Section 3.3, step 3. 7. Gel-purify the labeled DNAs as described in steps 19–21 of Section 3.2 (Fig. 22.4B). 3.5. Preparation of Specific dsDNA Probes for EMSA

1. ProxA (20 pmol) (1 ml) and ProxB (25 pmol) (1 ml) (see Section 2.3, step 7) are added to a mixture (34 ml) containing 5 ml 10 NEB buffer I (0.1 M Bis Tris propane-HCl, pH 7.0, 0.1 M MgCl2, and 10 mM DTT). 2. Incubate the above mixture at 94C for 5 min, 63C for 8 h, and then slowly cool down to room temperature.

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3. Supplement the mixture with 1 ml 0.1 M DTT, 2 ml 2 mM dATP/dGTP/dTTP, 1 ml 0.1 mM dCTP, 10 ml -32PdCTP, and 10 units of Klenow enzyme. 4. After 30 min incubation at room temperature, add 2 ml 0.5 M EDTA and transfer on ice. 5. Determine the specific radioactivity of end-labeled Prox DNA by TCA precipitation following the procedure described in Section 3.3, step 3. 6. The reaction mixture is loaded onto a G-50 column (5 ml) equilibrated with 1 TBS. 7. Elute the sample, collecting 200 ml fractions in microcentrifuge tubes. 8. Determine the distribution of 32P-labeled Prox DNA by determining the radioactivity of all fractions through Cerenkov counting (i.e., measurement of the radioactivity without using counting cocktail). It should be noted that Prox is reproducibly recovered in fractions 9–11. 9. Pool the peak fractions, and store in aliquots (50 ml) at –20C. 3.6. ATPase Assay

1. Prepare a master reaction mixture for ‘‘N’’ number of reactions as follows: (a) Mix ‘‘N+2’’ volumes of the following reagents: 0.3 ml 1 M HEPES–NaOH (20 mM)*, pH 7.4, 0.3 ml 0.1 M DTT (2 mM), 0.9 ml 50 mM MgCl2 (3 mM), 0.75 ml 1 mM ATP (50 mM), 0.3 ml (3 mCi) g-32P-ATP, 0.2 ml 1 mg/ml homopolymer RNA or DNA**, 0.15 ml 10 mg/ml bovine serum albumin (BSA) (0.1 mg/ml), and 7.1 ml de-ionized H2O. The final volume of the master mix should be (N+2) 10 ml. *; The final concentration of each component in the reaction mix (15 ml) is given in parentheses. **; If necessary, RNA or DNA homopolymer can be omitted in the master mix. (b) Dispense 10 ml aliquots into ‘‘N’’ number of microcentrifuge tubes. Add recombinant MLE (5–40 ng) and if needed, additional protein factors and RNA/DNA cofactors, and adjust the final reaction volume to 15 ml by adding de-ionized H2O. 2. After 0–60 min incubation at 37C, transfer the reaction mixture on ice, and aliquots (0.5 or 1 ml) of each reaction are spotted onto a PEI-F plate (about 15–18 samples per plate at about 2.5 cm from the bottom). 3. ATP and Pi are separated by chromatography for about 45 min in a TLC developing tank containing 75–100 ml TLC buffer (the buffer level should be kept lower than the line of spotted samples).

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4. Dry the TLC plate, and locate and quantify Pi and ATP using a Phosphorimager. Note that the specific radioactivity of ATP is 8,800 cpm/pmol. 3.7. Nucleotide UV-Crosslinking Assay

1. Prepare a master reaction mixture for ‘‘N’’ number of reactions as follows: (a) Mix ‘‘N+2’’ volumes of the following reagents: 0.4 ml 1 M HEPES–NaOH (20 mM)*, pH 7.4, 0.4 ml 0.1 M DTT (2 mM), 1.2 ml 50 mM MgCl2 (3 mM), 0.2 ml 1 mM GTP (10 mM), 0.2 ml (2 mCi) -32P-GTP, 0.2 ml 10 mg/ml bovine serum albumin (BSA) (0.1 mg/ml), and 7.4 ml de-ionized H2O. *; The final concentration of each component in the reaction mix (20 ml) is given in parentheses. (b) Dispense 10 ml aliquots into a 96-well assay plate. Add recombinant MLE (5–40 ng) and if needed, additional protein factors and RNA/DNA cofactors, and adjust the final reaction volume to 20 ml by adding de-ionized H2O. 2. After 10 min incubation on ice, irradiate for 10 min on ice with a UV-light source (254 nm) from 10 cm above the open plate. 3. Add 5 ml SDS sample buffer to each well, incubate for 5 min at 95C, and aliquots (10 ml) of each reaction are resolved by a 10% SDS-PAGE at 25 mA for 2.5 h. 4. After overnight fixation in a mixture of methanol and acetic acid (each 10%), the gel is dried on DE-81 paper under vacuum, and the GTP-MLE complex is visualized by autoradiography and quantified using a Phosphorimager.

3.8. Helicase Assay

1. Prepare a master reaction mixture for ‘‘N’’ number of reactions as follows: (a) Mix ‘‘N+2’’ volumes of the following reagents: 0.4 ml 1 M HEPES–NaOH (20 mM)*, pH 7.4, 0.4 ml 0.1 M DTT (2 mM), 1.2 ml 50 mM MgCl2 (3 mM), 2 ml 10 mM ATP (1 mM), 0.2 ml 10 mg/ml BSA (0.1 mg/ml), 1 ml 32Plabeled dsRNA or dsDNA substrate (50 fmol/reaction), 0.05 ml 40 U/ml RNasin (2 U/reaction), and 4.75 ml deionized H2O. The final volume of the master mix should be (N+2) 10 ml. *; The final concentration of each component in the reaction mix (20 ml) is given in parentheses. (b) Dispense 10 ml aliquots into ‘‘N’’ number of microcentrifuge tubes. Add recombinant MLE (5–40 ng) and if needed, additional protein factors such as topoisomerase II and RNA/DNA cofactors (Fig. 22.5), and adjust the final reaction volume to 20 ml by adding de-ionized H2O.

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Fig. 22.5. Influence of homopolymer RNA and topoisomerase II on MLE. (A) Helicase reaction was performed with a partial dsRNA substrate (25 fmol) (see Fig. 22.2B, lane 1), 10 ng MLE in the absence (lane 2) or presence of 60 fmol (lanes 3 and 5) or 300 fmol (lanes 4 and 6) homopolymer RNA. (B) Helicase reaction was provided with the indicated RNA homopolymer (300 fmol), 10 ng MLE, and increasing amounts (12.5 and 25 ng) of topoisomerase II. In (A) and (B), lane 1 shows the control helicase reaction containing dsRNA substrate alone.

2. After 30 min incubation at 37C, transfer the reaction mixture on ice, mix it with 5 ml of reaction termination buffer (see Note 3), and warm it to room temperature. 3. Aliquots (10 ml) of each reaction are loaded onto a 10% native polyacrylamide (30:1) gel and electrophoresed in 0.5 TBE at 19 mA until BPB migrates about two third of the gel (it usually takes 1.5–2 h). 4. The gel is then completely dried on top of DE-81 paper under vacuum at 80C. 5. Unwound ssRNA or ssDNA is visualized by autoradiography (Fig. 22.5). In addition, locate and quantify duplex substrates and unwound product using a Phosphorimager. 3.9. RNA/DNA EMSA

1. Prepare the master reaction mix containing ssRNA, dsRNA, non-specific dsDNA, or specific dsDNA (i.e., 54 bp Prox) following the procedure described in Section 3.8, step 1, in the presence or absence of ATP. For reactions containing Prox, 0.25 mg 1 Kb DNA ladder (Invitrogen) per reaction should be included in the master reaction mix. 2. After 30 min incubation at 37C, transfer the reaction mixture on ice, and mix it with 5 ml of native dye mix (see Note 3). 3. Aliquots (10 ml) of each reaction are loaded onto a 5 or 10% polyacrylamide (30:1)/5% glycerol composite gel and electrophoresed in 0.5 TBE at 19 mA until BPB migrates about two-thirds of the gel (it usually takes 2–3 h).

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4. The gel is dried on top of DE-81 paper under vacuum at 80C. 5. Visualize RNA- or DNA-MLE complexes by autoradiography and quantify them using a Phosphorimager (18, 19). 3.10. Immunoprecipitation (IP) and RT-PCR

1. Prepare two groups (1 and 2) of exponentially growing S2 cells on 150 mm culture plates at 80% confluency. 2. Carefully remove and discard culture medium from each well of the culture plate, and then collect and transfer the cells into two separate 50 ml conical tube on ice. 3. Centrifuge the tubes at 1,500 g for 5 min and wash the pellet once with ice-cold 1 PBS, and then carefully transfer the cell suspension to microcentrifuge tubes. 4. Two groups of cells are processed as follows. (a) For group 1, cells are quickly re-suspended in 1 ml Ultrapec reagent by pipetting, and total RNA is isolated following the procedure recommended by the manufacturer (see Note 4). Subsequently, proceed to step 20. (b) For group 2, cells are re-suspended in 1 extraction buffer, and proceed to step 5. 5. Homogenize the cell suspension using a micropestle and sonicate it three times, each 10 s with 1 min intervals on ice. 6. Remove the insoluble debris by centrifugation at 12,000 g for 5 min and transfer the clear total cell extract (TCE) into a new tube. 7. If necessary, repeat the centrifugation to clarify the TCE and determine the protein concentration using Bradford reagent (see Note 5). 8. Dilute aliquots (100 mg) of TCE to a final volume of 1 ml with 1 extraction buffer. 9. Add polyclonal antibodies (3 ml) specific for MLE, MSL1, or pre-immune sera (control), and the mixture is incubated on a nutator for 1 h at 4C. 10. While performing step 9, prepare protein A-sepharose as follows. (a) Take aliquots (100 ml) of protein A–sepharose suspension to microcentrifuge tubes* and mix them with 500 ml 1 extraction buffer. *Tube number should equal that of antibodies used. (b) Centrifuge the mixture at 3,000 rpm for 20 s. (c) Decant the supernatant and repeat the wash three times using 1 extraction buffer. (d) After the final wash, decant the supernatant and keep the protein A-sepharose on ice.

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11. Transfer the TCE/antibody mixtures to tubes containing protein A-sepharose, and incubate the mixture on a nutator for 30 min at 4C. 12. Centrifuge the mixture at 3,000 rpm for 20 s and remove the supernatant. 13. Wash the pellet with 1 ml 1 extraction buffer five times. 14. After the final wash, carefully remove the supernatant by pipetting. 15. The pellet is mixed with 40 ml 10 DNase I buffer and the final volume should be adjusted to 400 ml using DEPCtreated H2O. 16. Add 1 ml RNase-free DNase I, and the mixture is incubated for 20 min at 37C. 17. Isolate RNA by conventional phenol/chloroform and chloroform extraction method. 18. After ethanol precipitation, wash the RNA pellet with ice-cold 70% ethanol. 19. RNA is re-dissolved in DEPC-treated H2O to a final volume of 10 ml. 20. Prepare primary cDNA with an aliquot (5 mg) of total S2 cell RNA (Section 3.10, step 4(a)) or RNA obtained from the IP pellet (Section 3.10, step 19) using SuperScript II RT. 21. Constitute PCR reaction mixture (50 ml) using aliquots (2 ml) of cDNA, 1 PCR buffer, 0.25 mM dNTP mixture, 0.5 ml Taq DNA polymerase, roX2-5 primer (10 pmol), and roX2-3 primer (10 pmol). 22. PCR reaction is carried out as follows: 4 min at 94C, 20 or 30 cycles of (1 min at 94C, 1 min at 60C, 1 min at 72C), and 10 min at 72C. 23. PCR products are resolved on a 1.5% agarose gel. 24. After confirming their presence by ethidium bromide staining, the PCR products are subject to Southern blot analysis using a probe specific for roX2 (Fig. 22.6) (13, 24). 3.11. Reporter Gene Assay

1. S2 cells are grown in 6-well culture plate at 20% confluency. 2. Prepare a CaPO4-DNA mixture (100 ml/well) as follows: (a) To each microcentrifuge tube, add 1 mg Prox-luciferase reporter construct, 50 ng pMT/V5-b-galactosidase construct (an internal control vector), and varying amounts (0.4–1.2 mg) of either pMT-MLE or pMT-MSL1. (b) Adjust the total amount of DNA to 3 mg using pBS and the final volume to 40 ml using sterilized and de-ionized H2O.

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Fig. 22.6. In vivo association of MSL complex with roX2 RNA. Immunoprecipitation (IP) was performed using S2 cell lysate and the indicated antibodies. Subsequently, RT-PCR was performed with total RNA isolated from S2 cells or RNA obtained from the IP pellet. Lane 1, the unspliced product from the roX2 78.13 cDNA (10); lane 2, male larval total RNA (5 mg); lane 3, S2 cell total RNA (5 mg); lane 4, S2 cell lysate (100 mg); lane 5, MSL1 IP pellet; lane 6, IP pellet obtained with MSL1 pre-immune serum; lane 7, MLE IP pellet; lane 8, IP pellet obtained with MLE pre-immune serum; lane 9, pellet obtained with protein A-Sepharose alone without any antibody. Samples in lanes 1–4 were generated with 30 cycles of PCR and subsequently diluted 100- to 1,000-fold. Samples in lanes 5–9 were generated with 20 cycles of PCR. All samples were run on the same 1.5% agarose gel and Southern blotted. Two RT-PCR products for roX2 represent unspliced (roX2-1, 503 bp) and spliced form (roX2-2, 233 bp) of mRNA. Lanes 1–4 were exposed for 2 h and lanes 5–9 for 11 h.

(c) Add 50 ml 2 HBS and mix well by pipetting. (d) Add 6 ml 2 M CaCl2, mix by pipetting, and incubate for 30 min at room temperature. 3. Gently pipette the CaPO4-DNA precipitates up and down 3–4 times, and transfer them into the culture medium. 4. Rock the culture plate gently and incubate the transfected cells for 24 h. 5. Replenish with culture medium containing 0.5 mM CuSO4, and continue the incubation for an additional 24 h. 6. Carefully remove and discard 0.5 ml culture medium from each well of the culture plate, then collect and transfer the cells into microcentrifuge tubes on ice (see Note 6). 7. Centrifuge the tubes at 1,500 g for 5 min and wash the pellet once with ice-cold 1 PBS. 8. Quickly resuspend the pellet in 200 ml 1 luciferase lysis buffer by pipetting. 9. After 30 min incubation on ice, clear the lysate by centrifugation at 12,000 g for 15 min. 10. Transfer the clear lysate to fresh tubes and measure the b-galactosidase activity, a control to normalize the transfection efficiency, and the luciferase activity as reported previously (25) (see Fig. 22.7A).

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Fig. 22.7. Influence of MLE, MSL1, and MSL2 on the transcription activity of Prox2. (A) S2 cells, seeded in 6 well plate, were transfected with Prox2-luciferase (1 mg, lane 1) and increasing amounts (0.4–1.2 mg) of pMT/V5 vector expressing MLE (lanes 2 and 3), MSL1 (lanes 4 and 5), or MSL2 (lanes 6 and 7). On the following day, cells were treated with copper sulfate (0.5 mM), and 24 h later, total cell lysate was prepared and used to measure both luciferase and b-galactosidase activities, as described previously (19). Luciferase activities in lanes 1–7 were normalized to b-galactosidase and presented as RLU (relative luciferase unit) in comparison with that of control in lane 1. (B) RT-PCR was performed in a mixture (50 ml) provided with the indicated primers, as described previously (19). Following either 25 or 30 cycles of reaction, aliquots (5 ml) of PCR products were resolved on a 2% agarose gel and visualized by ethidium staining. Predicted size of RT-PCR product is 737 bp for MLE mRNA, 408 bp for dSRPK1 mRNA, and 322 bp for MSL1 mRNA. Two PCR products for roX2 RNA are described in Fig. 22.6. (C and D) RT-PCR was performed in the standard reaction mixture containing -32P-dCTP (30 mCi) and cDNA (1 ml) prepared from S2 cells expressing the indicated protein. RT-PCR products were isolated by spin column, and aliquots (10%) were resolved on a 2% agarose gel, followed by autoradiography (C). All RT-PCR products were quantified by Instant b-Imager, normalized to that of Drosophila SRPK1 (dSRPK1) and presented as relative RNA abundance in comparison with cognate RT-PCR product of control cells (D). Statistical analysis was performed using two-tailed Student’s t-test, comparing control and test samples. *p < 0.05.

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3.12. Multiplex PCR Assay

1. Transfect S2 cells following the procedures described in steps 1–4 of Section 3.11. 2. Replace the culture medium with media containing 0.5 mM CuSO4, and continue the incubation for an additional 48 h. 3. Carefully remove and discard 0.5 ml culture medium from each well of the culture plate, and then collect and transfer the cells into microcentrifuge tubes on ice. 4. Centrifuge the tubes at 1,500 g for 5 min and wash the pellet once with ice-cold 1 PBS. 5. Quickly resuspend the pellet in 1 ml Ultraspec reagent by pipetting. 6. Total RNA is isolated following the procedure recommended by the manufacturer (Biotecx Laboratories), and is dissolved in 85 ml DEPC-treated H2O and mixed with 10 DNase I buffer and 5 ml RNase-free DNase RQI. 7. After 30 min incubation at 37C, re-isolate RNA by phenol/ chloroform extraction and ethanol precipitation (see Note 4). 8. Prepare primary cDNA with an aliquot (1 mg) of total RNA using SuperScript II RT following the procedure recommended by the manufacturer. 9. Add an aliquot (1 ml) in a mixture (50 ml) containing 1 Advantage II PCR buffer, 0.25 mM dNTP mixture, 3 ml -32P-dCTP, 1 ml Advantage II polymerase, and a mixture of four PCR primer sets (10 pmol/primer) described in Section 2.7, step 8. 10. Perform PCR as follows: 1 min at 95C, 25 cycles of (95C for 15 s, 55C for 30 s, 68C for 1.5 min), and 7 min at 68C. 11. Purify PCR products using a spin column, and resolve aliquots (2.5–10 ml) on a 2% agarose gel in 0.5 TBE (Fig. 22.7B). 12. Visualize PCR products by ethidium bromide staining and autoradiography, and quantify them using a Phosphorimager (Fig. 22.7B–D).

4. Notes 1. This amount of -32P-GTP helps detect a longer ssRNA on a denaturing acrylamide but its contribution to the specific radioactivity of dsRNA formed is negligible. 2. If all the content is frozen due to prolonged incubation, thaw it by rubbing the tube a few seconds until it becomes a jelly.

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3. At this step, the reaction mixture can be stored at –70C for future analysis. 4. About 85–100 mg of total RNA is obtained from 1 107 S2 cells. 5. Protein concentration is usually 10–20 mg/ml. 6. S2 cells are easily detached from the culture plate, especially following the DNA transfection. Therefore, S2 cells can be efficiently recovered by pipetting. References 1. Park Y. and Kuroda M. I. (2001) Epigenetic aspects of X-chromosome dosage compensation. Science 293, 1083–1085. 2. Palmer M. J., Mergner V. A., Richman R., Manning J. E., Kuroda M. I., and Lucchesi J. C. (1993) The male-specific lethal-one (msl-1) gene of Drosophila melanogaster encodes a novel protein that associates with the X chromosome in males. Genetics 134, 545–557. 3. Bashaw G. J. and Baker B. S. (1995) The msl-2 dosage compensation gene of Drosophila encodes a putative DNA-binding protein whose expression is sex specifically regulated by sex-lethal. Development 121, 3245–3258. 4. Kelley R. L., Solovyeva I., Lyman L. M., Richman R., Solovyev V., and Kuroda M. I. (1995) Expression of msl-2 causes assembly of dosage compensation regulators on the X chromosomes and female lethality in Drosophila. Cell 81, 867–877. 5. Gorman M., Franke A., and Baker B. S. (1995) Molecular characterization of the male-specific lethal-3 gene and investigations of the regulation of dosage compensation in Drosophila. Development 121, 463–475. 6. Kuroda M. I., Kernan M. J., Kreber R., Ganetzky B., and Baker B. S. (1991) The maleless protein associates with the X chromosome to regulate dosage compensation in Drosophila. Cell 66, 935–947. 7. Hilfiker A., Hilfiker-Kleiner D., Pannuti A., and Lucchesi J. C. (1997) mof, a putative acetyl transferase gene related to the Tip60 and MOZ human genes and to the SAS genes of yeast, is required for dosage compensation in Drosophila. EMBO J. 16, 2054–2060. 8. Jin Y., Wang Y., Johansen J., and Johansen K. M. (2000) JIL-1, a chromosomal kinase implicated in regulation of chromatin

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structure, associates with the male specific lethal (MSL) dosage compensation complex. J. Cell Biol. 149, 1005–1010. Meller V. H., Wu K. H., Roman G., Kuroda M. I., and Davis R. L. (1997) roX1 RNA paints the X chromosome of male Drosophila and is regulated by the dosage compensation system. Cell 88, 445–457. Amrein H. and Axel R. (1997) Genes expressed in neurons of adult male Drosophila. Cell 88, 459–469. Kageyama Y., Mengus G., Gilfillan G., Kennedy H. G., Stuckenholz C., Kelley R. L., Becker P. B., and Kuroda M. I. (2001) Association and spreading of the Drosophila dosage compensation complex from a discrete roX1 chromatin entry site. EMBO J. 20, 2236–2245. Kelley R. L., Meller V. H., Gordadze P. R., Roman G., Davis R. L., and Kuroda M. I. (1999) Epigenetic spreading of the Drosophila dosage compensation complex from roX RNA genes into flanking chromatin. Cell 98, 513–522. Meller V. H., Gordadze P. R., Park Y., Chu X., Stuckenholz C., Kelley R. L., and Kuroda M. I. (2000) Ordered assembly of roX RNAs into MSL complexes on the dosage-compensated X chromosome in Drosophila. Curr. Biol. 10, 136–143. Park Y., Mengus G., Bai X., Kageyama Y., Meller V. H., Becker P. B., and Kuroda M. I. (2003) Sequence-specific targeting of Drosophila roX genes by the MSL dosage compensation complex. Mol. Cell 11, 977–986. Oh H., Park Y., and Kuroda M. I. (2003) Local spreading of MSL complexes from roX genes on the Drosophila X chromosome. Genes Dev. 17, 1334–1339. Park Y., Kelley R. L., Oh H., Kuroda M. I., and Meller V. H. (2002) Extent of chromatin spreading determined by roX RNA

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intrinsically associated with U2AF. J. Biol. Chem. 268, 13472–13478. Lee C. G., Hague L. K., Li H., and Donnelly R. (2004) Identification of toposome, a novel multisubunit complex containing topoisomerase IIalpha. Cell Cycle 3, 638–647. Zhou K., Choe K. T., Zaidi Z., Wang Q., Mathews M. B., and Lee C. G. (2003) RNA helicase A interacts with dsDNA and topoisomerase IIalpha. Nucleic Acids Res. 31, 2253–2260. Park Y., Oh H., Meller V. H., and Kuroda M. I. (2005) Variable splicing of noncoding roX2 RNAs influences targeting of MSL dosage compensation complexes in Drosophila. RNA Biol. 2, 157–164. Nakajima T., Uchida C., Anderson S. F., Lee C. G., Hurwitz J., Parvin J. D., and Montminy M. (1997) RNA helicase A mediates association of CBP with RNA polymerase II. Cell 90, 1107–1112.

Chapter 23 Biochemical Characterization of Human Upf1 Helicase Zhihong Cheng, Gaku Morisawa, and Haiwei Song Abstract We present here the biochemical characterization of human Upf1 helicase core (hUpf1c). hUpf1c is overexpressed as a GST fusion protein in Escherichia coli and purified using chromatographic methods. In vitro ATP binding and single-stranded RNA (ssRNA) binding activities are measured using dot-blot technique. Measurement of RNA-dependent ATPase activity is performed by thin layer chromatography (TLC). The ATP-modulated ssRNA binding activity is examined by surface plasma resonance (SPR). The binding of double-stranded DNA (dsDNA) to hUpf1c is checked by electrophoretic mobility shift assay (EMSA, gel shift assay). Key words: Upf1, helicase, ATPase, RNA binding, nonsense-mediated mRNA decay, superfamily 1, NMD.

1. Introduction Nonsense-mediated mRNA decay is an evolutionally conserved mRNA quality-control mechanism that selectively degrades aberrant mRNAs containing premature termination codons in order to prevent the accumulation of truncated proteins, which are often non-functional or potentially deleterious in the cells (1–3). Three conserved proteins Upf1, Upf2, and Upf3 constitute the core NMD machinery (4–9) with Upf1 as the key member in this protein set. Upf1 acts in concert with the peptide release factors eRF1/eRF3 to recognize aberrant translation termination events and, together with Upf2 and Upf3, triggers degradation of mRNA in a subsequent step (10–12). In addition to its role in NMD, Upf1 also regulates mRNAs in a NMD-independent manner. For example, Upf1 is recruited by the RNA-binding protein Staufen to the downstream of a stop M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_23, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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cordon to degrade some mRNAs in a Staufen-mediated decay (13). Human Upf1 (hUpf1) has been shown to regulate the degradation of histone mRNAs in mammalian cells (14). Moreover, hUpf1 is involved in nonsense-mediated altered splicing (15). Upf1 contains two conserved functional regions, an N-terminal Cys-rich domain and a C-terminal helicase domain, which belong to superfamily 1 (SF1) of DNA/RNA helicases (16, 17). It has been shown that Upf1 exhibits RNA binding, RNA-dependent ATP hydrolysis, and 50 to 30 ATP-dependent RNA helicase activities, which are critical for its role in NMD (18–20). The crystal structure of the human helicase core has been determined (21). Structural data combined with mutational analysis reveals a mechanism by which ATP binding modulates the RNA binding to Upf1 (21). In this chapter, biochemical characterization of human Upf1 helicase core domain (hUpf1c) is described. The cDNA encoding hUpf1c is cloned into the pGEX-6P-1 vector and expressed as a GST-fusion protein in E. coli. GST affinity, ion exchange, and size exclusion chromatography are used to purify hUpf1c. ATP and single-stranded RNA (ssRNA) binding assays are carried out based on the unique property of PVDF membrane on protein-specific absorption. Effects of nucleotides on ssRNA binding to hUpf1c are measured by Surface Plasma Resonance (SPR), which is the label-free study of interactions between biomolecules (22). The sensorgram of SPR reflects the mass change in real time near the sensor chip surface, on which one molecule is immobilized; the other exists in the nearby micro fluid (23). Thin layer chromatography has the capability of differentiating small molecules like ATP, ADP, and phosphate, enabling researchers to measure ATPase activity (http://inst.sfcc.edu/chemscape/catofp/chromato/tlc/tlc.htm). Finally, double-stranded DNA (dsDNA) binding to hUpf1c is examined by electrophoretic mobility shift assay (EMSA), which is based on the observation that protein:DNA complexes migrate more slowly than free DNA molecules when subjected to non-denaturing polyacrylamide or agarose gel electrophoresis. Because the rate of DNA migration is shifted or retarded upon protein binding, the assay is also referred to as a gel shift or gel retardation assay.

2. Materials 2.1. Cloning and Overexpression

1. Human cDNA library (ATCC). 2. PCR primers (Sigma, Proligo).

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3. pGEX-6p-1 vector (Amersham). 4. Turbo Pfu DNA polymerase (Stratagene). 5. Restriction enzyme and ligase (New England Biolabs). 6. TAE buffer (1X): 40 mM Tris, 1.14% acetic acid, 1 mM EDTA. 7. 1.5% agarose gel in 1X TAE buffer. 8. QIAquick PCR purification kit, Gel Extraction kit and QIAprep Spin Miniprep kit (QIAGEN). 9. DH-5a competent cell (Invitrogen), BL-21 DE3 expression strain (Novagen). 10. 80% Glycerol autoclaved. 11. LB medium: 10 mg/ml tryptone, 5 mg/ml yeast extract, 10 mg/ml NaCl, 100 mg/ml ampicillin, pH 7. 12. Isopropyl-b-D-thiogalactopyranosid (IPTG), 1 M in stock. 13. Sample loading buffer (3X): 200 mM Tris-Cl, pH 6.8, 6.5% (w/v) SDS, 0.16% (v/v) b-mercaptoethanol, 0.6 mg/ml bromophenol blue. 14. Running buffer (1X): 3.03 mg/ml Tris base, 14.4 mg/ml glycine, 1 mg/ml SDS. 15. Gel staining buffer: 45% methanol, 10% acetic acid, 0.25% Coomassie Brilliant Blue R-250. 16. De-staining buffer: 5% methanol and 7.5% acetic acid. 2.2. Purification

1. Hen egg lysozyme (Sigma). 2. LB medium: 10 mg/ml tryptone, 5 mg/ml yeast extract, 10 mg/ml NaCl, 100 mg/ml ampicillin, pH 7. 3. PBS buffer: 8 g NaCl, 0.2 g KCl, 1.44 g Na2HPO4, 0.24 g KH2PO4 in 1 l, pH 7.4. 4. Lysis buffer: 20 mM Tris, 500 mM NaCl, 2 mM DTT, 2 mM benzamidine, 0.1 mM PMSF, pH 6.0, 1 mg/ml lysozyme. 5. Elution buffer: 20 mM L-glutathione reduced (GSH) in lysis buffer. 6. Coomassie Protein Assay kit (Pierce). 7. PreScission protease (Amersham). 8. Desalting buffer: 20 mM HEPES, 100 mM NaCl, 2 mM DTT, pH 7.0. 9. Ion-exchange buffer A: 20 mM HEPES, 100 mM NaCl, 2 mM DTT, pH 7.0. 10. Ion-exchange buffer B: 20 mM HEPES, 1000 mM NaCl, 2 mM DTT, pH 7.0.

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11. Gel filtration buffer: 20 mM HEPES, 200 mM NaCl, 2 mM DTT, pH 7.0. 2.3. In Vitro ATP Binding

1. PVDF membrane for protein blotting (Bio-rad). 2. 5000 Ci/mmol [g-32P] ATP (Amersham). 3. Blocking buffer: 20 mM HEPES, 50 mM KAc, 2.5 mM Mg(Ac)2, 2 mM DTT, 3% BSA, pH 7.0. 4. Binding buffer: 20 mM HEPES, 50 mM KAc, 2.5 mM Mg(Ac)2, 2 mM DTT, 1.5% BSA, pH 7.0. 5. Cling wrap. 6. Autoradiography film cassette and Hyperfilm (Amersham).

2.4. In Vitro ssRNA Binding

1. PVDF membrane for protein blotting (Bio-rad). 2. 2 mg/ml BSA protein. 3. 5000 Ci/mmol [g-32P] ATP (Amersham). 4. Single-stranded RNA 50 CGCCCGAGGCTGTGCCGT30 (Dharmacon). 5. T4 kinase kit (Invitrogen). 6. G-25 Spin column (Amersham). 7. Blocking buffer: 20 mM HEPES, 50 mM KAc, 2.5 mM Mg(Ac)2, 2 mM DTT, 3% BSA, pH 7.0. 8. Binding buffer: 20 mM HEPES, 50 mM KAc, 2.5 mM Mg(Ac)2, 2 mM DTT, 1.5% BSA, pH 7.0. 9. Cling wrap. 10. Autoradiography film cassette and Hyperfilm (Amersham).

2.5. In Vitro ATP Hydrolysis

1. 500 nM 15-mer poly (C) purchased from Dharmacon. 2. 5000 Ci/mmol [g-32P] ATP (Amersham). 3. 500 mM EDTA. 4. Reaction buffer: 50 mM MES, pH 6.0, 50 mM KAc, 2.5 mM Mg(Ac)2, 1 mM DTT, 0.1 mg/ml BSA. 5. Nucleotide ATP (Sigma) dissolved in distilled water. 6. PEI-cellulose plate (Sigma). 7. Development buffer: 0.3 M K2HPO4, pH 7.6. 8. Glass development tank (Sigma). 9. Cling wrap. 10. Autoradiography film cassette and Hyperfilm (Amersham).

2.6. Effects of Nucleotides on ssRNA Binding to hUpf1c

1. BIAcore 2000 instrument (BIAcore). 2. 50 Biotin-labeled ssRNA TGTCATTCGAGTACAGTCTGT TCAGCTAGTCTCC (CureVac).

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3. Steptavidin-coated Sensor Chip SA (BIAcore). 4. Coupling buffer: 10 mM HEPES, pH 7.0, 150 mM NaCl, 0.005% (v/v) Tween-20. 5. 0.3 M NaCl and 2.5 M NaCl. 6. 1 mg/ml biotin dissolved in 0.3 M NaCl (Sigma). 7. Binding buffer: 20 mM HEPES, pH 7.0, 100 mM NaCl, 2 mM MgCl2, 2 mM DTT, 0.002% (v/v) Tween-20. 8. Nucleotide ATP (Sigma) dissolved in distilled water. 2.7. Gel Shift Assay of dsDNA Binding to hUpf1c

1. T4 Polynucleotide Kinase (New England Biolabs). 2. 3000 Ci/mmol [g-32P] ATP (Amersham). 3. 500 mM EDTA. 4. G-25 Spin column (Amersham). 5. Single-stranded 28 nt DNA 50 AAACAAAA CTAGCACCGT AAAGCAAGCT30 and 18 nt DNA 50 AGCTTGCTTTAC GGTGCT30 (Sigma). 6. Binding buffer: 20 mM Tris, pH 8.0, 50 mM KCl, 3 mM MgCl2, 1 mM DTT. 7. TBE buffer (5X): 30.28 g Tris base, 1.86 g EDTA, and 13.25 g boric acid dissolved in 1 l water. 8. Vertical electrophoresis apparatus (Bio-rad). 9. Gel dryer (Bio-rad). 10. Autoradiography film cassette and Hyperfilm (Amersham).

3. Methods 3.1. Cloning and Overexpression

1. Gene fragment encoding hUpf1c (residues 295–914) is amplified by PCR using human cDNA library as the template. General PCR parameters are used (see Note 1). 2. 5 ml of the PCR product is run on a 1.5% agarose gel with 1X TAE buffer as running buffer. The PCR product is purified using a QIAquick PCR purification kit (QIAGEN). 3. Restriction enzyme digestions of plasmid pGEX-6p-1 and PCR product are performed at 37C for 3–5 h. The digested DNA fragments were excised from the gel and purified using a QIAquick Gel Extraction kit (QIAGEN). 4. Ligation is performed at 15C overnight. Five microliters of the ligation product is then transformed into 100 ml of DH5a competent cells. Positive clones are screened by colony PCR or double digestion. The sequences of positive clones are

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confirmed by DNA sequencing. The recombinant plasmid is transformed into BL-21 star (DE3) expression strains for overexpression of hUpf1c. 5. The resulting expression strain is inoculated into 2 ml of LB medium supplemented with 100 mg/ml ampicillin and incubated overnight at 37C. Five hundred microliters of cells mixed with 80% glycerol were stored at –80C. For trial expression, 100 ml of cells is transferred into 10 ml of fresh LB medium and cultured at 37C. When the optical density at 600 nm (OD600) reaches to about 0.5–0.6, 0.5 mM of isopropyl-ß-D-thiogalactopyranosid (IPTG) is added to the cell culture. Further 2 h incubation at 37C is required to get sufficient amount of overexpressed hUpf1c. Two hundred microliters of non-induced cells and 100 ml of induced cells are pelleted and boiled in the SDS-PAGE loading buffer. The samples are separated by SDS-PAGE using a 12% polyacrylamide gel to check the protein expression level (see Note 2). 3.2. Purification

1. The strain overexpressing hUpf1c is inoculated into 150 ml of LB medium supplemented with 100 mg/ml ampicillin and grown at 37C overnight. One hundred and twenty milliliters of the overnight culture is added into 4 l of LB medium and incubated in shaker (220 rpm) at 37C. When OD600 reaches to 0.5–0.6, 0.1 mM IPTG is added into the cell culture and further cell growth is performed overnight at 18C. The cells are harvested by centrifugation (4000 rpm), and washed by the PBS buffer. The resulting cell pellets are stored at –80C for subsequent protein purification (see Note 3). 2. All steps of protein purification must be performed at 4C. The frozen cell pellets are thawed at room temperature and re-suspended in the lysis buffer containing 1 mg/ml Hen egg lysozyme for at lease 30 min. The lysozyme-treated cells are subjected to sonication with eight bursts of 15 s using a Soniprep 150 (SANYO). At least 90 s incubation on ice is required after each sonication burst to cool down the temperature of the cells. Cell debris is removed by centrifugation for 1 h using J6-HC centrifuge (Beckman Coulter) at 18,000 rpm. The supernatant is filtered (0.45 mm) and mixed with 2 ml of glutathione sepharose 4B (Amersham) for 30 min at 4C. 3. The resulting GST beads are centrifuged at 780 g for 10 min and washed with 50 ml of the lysis buffer. At least three cycles of centrifugation/washing are required to remove the non-specific bound proteins to the GST beads. During the centrifugation, elution buffer is prepared by adding 20 mM GSH into the lysis buffer (see Note 4).

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4. GST-hUpf1c is eluted with 15 ml of the elution buffer. Protein concentration is determined by the Bradford method using the Coomassie Protein Assay kit (Pierce) with BSA as the standard. PreScission protease (Amersham) is added to the eluted fraction at a ratio of 50:1 (w/w) between target protein and protease. The reaction mixture is incubated at 4C overnight to cleave off the GST tag. 5. The protease-treated protein sample is loaded onto a HiPrep desalting column (Amersham) equilibrated with the desalting buffer. The desalted protein sample is then reloaded onto a regenerated glutathione sepharose 4B column to remove the cleaved GST tag and non-cleaved GST-hUpf1c. 6. The flow-through from the second glutathione sepharose 4B column is loaded onto a MonoS HR 10/10 column (Amersham) pre-equilibrated with ion exchange buffer B and subsequently buffer A. The hUpf1c is eluted with a 100 ml linear gradient of NaCl (0.1–1.0 M) using the same buffers (A and B). The fractions containing hUpf1c are verified by SDS-PAGE and pooled for further purification. 7. The pooled fractions from MonoS column are loaded onto a Superdex-200 column (Amersham) pre-equilibrated with the gel filtration buffer. The fractions containing pure hUpf1c are verified by SDS-PAGE (Fig. 23.1a) and concentrated to 10 mg/ml using vivaspin 20 concentrators (30,000 MWCO, Vivascience). The concentrated hUpf1c is aliquoted and stored at –80C. 3.3. In Vitro ATP Binding

1. Cut a proper size of PVDF membrane; slowly lower the membrane into 100% methanol and shake briefly for 10 s. After the membrane become translucent, rinse the membrane with deionized water and equilibrated in a binding buffer for at least 10 min. Keep the membrane fully wet all the time. If the membrane becomes partially dry, allow it to dry completely and repeat step 1. Do not touch membrane directly by hands; use forceps instead. 2. Soak two thick tissue papers into water and place them on a sheet of cling wrap. Slowly place the PVDF membrane onto the tissue paper and remove the air bubbles between them (see Note 5). Keep the tissue papers wet by adding water. Dotblot equal amounts (15 mg) of BSA and hUpf1c onto the membrane. After protein samples are absorbed into membrane, transfer the membrane into the blocking buffer and incubate for 1 h at room temperature. 3. Transfer the membrane into a binding buffer and add 50 mCi of 5000 Ci/mmol [g-32P] ATP (Amersham) for incubation at low rotation speed for 30 min.

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Fig. 23.1. Purification and biochemical characterization of hUpf1c. a. Ten percent SDS-PAGE of the fractions (F4–F10) from Superdex-200 column. Ten microliters of protein samples in each fraction are loaded for SDS-PAGE. b. ATP binding activity of hUpf1c (15 mg). c. RNA-dependent ATPase activity of hUpf1c: 200 nM protein in the presence of 500 nM 15-mer poly(C). d. Surface plasma resonance analysis of ssRNA binding to 100 nM hUpf1c in the absence or presence of 2 mM ATP. e. dsDNA binding activity to hUpf1c. Except for the control sample (lane 1), 1 pmol (lane 2) or 5 pmol (lanes 3–7) of purified hUpf1c are used for binding assay. At a high salt concentration, 50 mM (lane 4), 100 mM (lane 5), and 200 mM (lane 6), sodium chloride appears to inhibit Upf1-DNA complex formation. Non-labeled double-stranded DNA (15 pmol) is also included as a competitor in the reaction (lane 7).

4. After incubation, the membrane was washed twice with 10 ml of the binding buffer each time. The signal was measured by exposing the membrane to Hyperfilm and quantified using a Phosphoroimager (Molecular Dynamics). An example of the ATP binding is shown in Fig. 23.1b. 3.4. ATP Hydrolysis

1. In vitro ATPase assay is performed in 20 ml of reaction mixture. The mixture contains reaction buffer, 200 nM hUpf1c, 500 nM 15-mer poly(C), 10 mM cold ATP, and 1 mCi of 5000 Ci/mmol [g-32P] ATP (Amersham). The reaction condition should be optimized through several trial experiments (see Note 6).

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2. The reaction mixture is incubated for 1 h at 37C. During the incubation, prepare a proper size of PEI-cellulose plate and the development buffer. Make the glass tank ready for development. 3. After incubation, add 1 ml of 500 mM EDTA into the mixture to terminate the reaction and spot 1 ml of reaction mixture on the PEI-cellulose plate. The plate is placed vertically in the glass tank with the spots of samples 1 cm above the development buffer. 4. After the edge of development buffer reaching to the top of plate, take the plate out and cover the plate by a plastic wrap. The plate is exposed to Hyperfilm and quantified using a Phosphoroimager. An example result is shown in Fig. 23.1c. 3.5. In Vitro ssRNA Binding Assay

1. PVDF membrane is prepared as described in step 1 of in vitro ATP binding assay. 2. Label ssRNA with [g-32P] ATP. The labeling reaction mixture of 25 ml contains 1X reaction buffer and 10 units of T4 kinase (Invitrogen), 5 pmol ssRNA, and 50 mCi of 5000 Ci/ mmol [g-32P] ATP (Amersham), and is incubated at 37C for 10 min. 3. The reaction mixture is then passed through G-25 spin column (Amersham) to remove unused [g-32P] ATP. 4. As in step 2 of in vitro ATP binding assay, dot-blot equal amounts (15 mg) of BSA and hUpf1c onto the membrane, transfer the membrane into the blocking buffer and incubate it for 1 h at room temperature. 5. Transfer the membrane into the binding buffer, add in 1 pmol of 32P-labeled RNA substrate, and incubate for additional 1 h with low speed shaking. 6. After incubation, the membrane is washed with 20 ml of the binding buffer three times and then measured in the same manner as in step 4 of ATP binding assay.

3.6. Effects of Nucleotides on the ssRNA Binding to hUpf1c

1. Undock the maintenance chip or the previously used sensor chip and remove it from the instrument. Insert and dock the new Sensor chip SA, set up the instrument buffer tubing into the coupling buffer (see Note 7). Perform the prime to exchange all of the solutions in the system with the coupling buffer. 2. The coupling buffer is flowed across the sensor chip SA until the trace leveled off. During the equilibration, prepare 100 nM of biotin-labeled RNA.

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3. Inject 20 ml of 100 nM biotin-labeled RNA in 0.3 M NaCl at the flow rate of 5 ml/min. 4. After immobilization, 100 ml of 1 mg/ml biotin is flowed across the flow cell 2 and reference flow cell 1 at the flow rate of 5 ml/min to block the unbinding site of the sensor chip. 5. Set up the instrument buffer tubing into the binding buffer and perform the prime to exchange all of the solutions in the system with the binding buffer. Keep the instrument in standby mode when not in use. 6. Inject 150 ml of 100 nM wild-type protein in the absence of ATP into the binding buffer across the sensor chip at the flow rate of 50 ml/min, followed by the dissociation time of 180 s. 7. The bound protein sample is removed by injecting 50 ml of regeneration buffer 2.5 M NaCl to regenerate the sensor chip for next cycle of binding assay (see Note 8). 8. Repeat steps 6 and 7 by injecting protein sample in the presence of 2 mM ATP. 9. The data curves were analyzed using the BIAevaluation software. An example result is shown in Fig. 23.1d. 3.7. Gel Shift Assay of dsDNA Binding to hUpf1c

1. DNA duplex is prepared by annealing 18- and 28-nt singlestrand oligodeoxyribonucleotide. 2. Double-stranded DNA (3 pmol) is labeled at the 50 end with 10 mCi [g-32P] ATP (3000 Ci/mmol) and 10 units of T4 polynucleotide kinase for 30 min at 37C. Stop the reaction by adding 1 ml of 0.5 M EDTA and reaction mixture is passed through G-25 spin column to remove unincorporated [g-32P] ATP. 3. Binding reaction is performed in 10 ml volume of the binding buffer (20 mM Tris, pH 8.0, 3 mM MgCl2, 1 mM DTT, 50 mM KCl). 4. Purified hUpf1c and 50 fmol of labeled double-stranded DNA was added to the binding buffer and incubated for 15 min at room temperature. Different concentrations of hUpf1c, sodium chloride, and nucleic acid are used for comparative study. 5. Fill an electrophoresis apparatus with 0.5X TBE and preelectrophorese the 4.5% non-denaturing gel (contains 0.5 TBE and 5% glycerol) for 30 min at 100 V and samples are resolved in a gel for 1 h. 6. Gel is dried and exposed to Hyperfilm. An example result is shown in Fig. 23.1e.

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4. Notes 1. Turbo Pfu DNA polymerase should be used to avoid random mutations. In some cases, optimizing the concentration of Mg2+ may improve the quality of PCR product. 2. High concentration of IPTG is used in trial expression to shorten the time for induction. 3. In case of proteins’ poor solubility, low temperature (such as at 18C) induction is preferred to improve protein solubility and yield. 4. It is essential to adjust the pH back to its original value after adding GSH to the elution buffer as GSH will decrease the elution buffer’s pH to around 3. 5. It is critical to remove air bubbles between the PVDF membrane and tissue papers. 6. The concentration of each reactant or substrate should be optimized by a couple of trial assays by varying the concentrations of proteins, cold ATP, RNA, as well as incubation time and temperature. 7. Operate the instrument following the manual provided by the manufacturer. All buffers used in this assay should be filtered using a 0.22-mm filter and degassed prior to use. 8. Optimize the regeneration buffer by injecting different concentrations of NaCl into the sensor chip and select the lowest concentration of NaCl that can remove the bound protein completely.

Acknowledgments This work is financially supported by the Biomedical Research Council of A*STAR (Agency for Science, Technology and Research). References 1. Baker K. E. and Parker R. (2004) Nonsensemediated mRNA decay: terminating erroneous gene expression. Curr. Opin. Cell Biol. 16, 293–299. 2. Conti E. and Izaurralde E. (2005) Nonsense-mediated mRNA decay: molecular insights and mechanistic variations across

species. Curr. Opin. Cell Biol. 17, 316–325. 3. Lejeune F. and Maquat L. E. (2005) Mechanistic links between nonsense- mediated mRNA decay and pre-mRNA splicing in mammalian cells. Curr. Opin. Cell Biol. 17, 309–315.

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4. Cui Y., Hagan K. W., Zhang S., and Peltz S. W. (1995) Identification and characterization of genes that are required for the accelerated degradation of mRNAs containing a premature translational termination cordon. Genes Dev. 9, 423–436. 5. He F., Brown A. H., and Jacobson A. (1997) Upf1p, Nmd2p, and Upf3p are interacting components of the yeast nonsense-mediated mRNA decay pathway. Mol. Cell Biol. 17, 1580–1594. 6. Hodgkin J., Papp A., Pulak R., Ambros V., and Anderson P. (1989) A new kind of informational suppression in the nematode Caenorhabditis elegans. Genetics 123, 301–313. 7. Gatfield D., Unterholzner L., Ciccarelli F. D., Bork P., and Izaurralde E. (2003) Nonsense-mediated mRNA decay in Drosophila: at the intersection of the yeast and mammalian pathways. EMBO J. 22, 3960–3970. 8. Lykke-Andersen J., Shu M. D., and Steitz J. A. (2000) Human Upf proteins target an mRNA for nonsense-mediated decay when bound downstream of a termination codon. Cell 103, 1121–1131. 9. Serin G., Gersappe A., Black J. D., Aronoff R., and Maquat L. E. (2001) Identification and characterization of human orthologues to Saccharomyces cerevisiae Upf2 protein and Upf3 protein (Caenorhabditis elegans SMG-4). Mol. Cell Biol. 21, 209–223. 10. Amrani N., Ganesan R., Kervestin S., Mangus D. A., Ghosh S., and Jacobson A. (2004) A faux 30 -UTR promotes aberrant termination and triggers nonsensemediated mRNA decay. Nature 432, 112–118. 11. Kashima I., Yamashita A., Izumi N., Kataoka N., Morishita R., Hoshino S., Ohno M., Dreyfuss G., and Ohno S. (2006) Binding of a novel SMG-1-Upf1-eRF1-eRF3 complex (SURF) to the exon junction complex triggers Upf1 phosphorylation and nonsense-mediated mRNA decay. Genes Dev. 20, 355–367. 12. Sheth U. and Parker R. (2006) Targeting of aberrant mRNAs to cytoplasmic processing bodies. Cell 125, 1095–1109. 13. Kim Y. K., Furic L., Desgroseillers L., and Maquat L. E. (2005) Mammalian staufen1

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recruits Upf1 to specific mRNA 30 UTRs so as to elicit mRNA decay. Cell 120, 195–208. Kaygun H. and Marzluff W. F. (2005) Regulated degradation of replica- tondependent histone mRNAs requires both ATR and Upf1. Nat. Struct. Mol. Biol. 12, 794–800. Mendell J. T., ap Rhys C. M., and Dietz H. C. (2002) Separable roles for rent1/ hUpf1 in altered splicing and decay of nonsense transcripts. Science 298, 419–422. Applequist S. E., Selg M., Raman C., and Jack H. M. (1997) Cloning and characterization of HUPF1, a human homolog of the Saccharomyces cerevisiae nonsense mRNAreducing UPF1 pro- tein. Nucleic Acids Res. 25, 814–821. Culbertson M. R. and Leeds P. F. (2003) Looking at mRNA decay pathways through the window of molecular evolution. Curr. Opin. Genet. Dev. 13, 207–214. Czaplinski K., Weng Y., Hagan K. W., and Peltz S. W. (1995) Purification and characterization of the Upf1 protein: a factor involved in translation and mRNA degradation. RNA 1, 610–623. Weng Y., Czaplinski K., and Peltz S. W. (1996) Genetic and biochemical characterization of mutations in the ATPase and helicase regions of the Upf1 protein. Mol Cell Biol. 16, 5477–5490. Bhattacharya A., Czaplinski K., Trifillis P., He F., Jacobson A., and Peltz S. W. (2000) Characterization of the biochemical properties of the human Upf1 gene product that is involved in nonsensemediated mRNA decay. RNA 6, 1226–1235. Cheng Z., Muhlrad D., Lim M., Parker R., and Song H. (2007) Structural and functional insights into the human Upf1 helicase core. EMBO J. 26, 253–264. Lieberg B., Cylinder C., and Lundstrom I. (1983) How surface plasmon resonance for gas detection and biosensing. Sens. Actuators 4, 299–304. Jason-Moller L., Murphy M., and JoAnne Bruno J. (2006) Overview of biscoe systems and their applications. Curr. Protoc. Protein Sci. 19.13.1–19.13.14.

Chapter 24 Assays of the Helicase, ATPase, and Exoribonuclease Activities of the Yeast Mitochondrial Degradosome Michal Malecki, Piotr P. Stepien, and Pawel Golik Abstract The mitochondrial degradosome (mtEXO) is the main enzymatic complex in RNA degradation, processing, and surveillance in Saccharomyces cerevisiae mitochondria. It consists of two nuclear-encoded subunits: the ATP-dependent RNA helicase Suv3p and the 30 to 50 exoribonuclease Dss1p. The two subunits depend on each other for their activity; the complex can therefore be considered as a model system for the cooperation of RNA helicases and exoribonucleases in RNA degradation. All the three activities of the complex (helicase, ATPase, and exoribonuclease) can be studied in vitro using recombinant proteins and protocols presented in this chapter. Key words: RNA helicase, RNA degradation, degradosome, mtEXO, mitochondria, yeast.

1. Introduction RNA turnover is one of the key processes in the regulation of gene expression and plays an important role in removing aberrant forms of RNA resulting from defective synthesis or maturation of RNA molecules (RNA surveillance) (1–5). In most physiological conditions, the level of each RNA species is the result of a balance between transcription and degradation. We recently demonstrated the critical importance of maintaining this balance in the yeast mitochondrial system (6). Mechanisms of RNA degradation in different genetic systems are very divergent and involve several classes of ribonucleases (both 30 to 50 and 50 to 30 exoribonucleases as well as endoribonucleases) and various other activities (1, 2, 5, 7–11). In general, cooperation of ribonucleases and RNA helicases appears to be a common theme in prokaryotic (7), eukaryotic (10, 11), and organellar (1) systems. M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_24, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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RNA turnover in yeast mitochondria is carried out by a relatively simple enzymatic complex called the mitochondrial degradosome or mtEXO, consisting of only two subunits: a 30 to 50 exoribonuclease encoded by the nuclear gene DSS1 (YMR287C) belonging to the RNR (RNase II-like) family and an NTP-dependent RNA helicase related to the DExH/D (Ski2p) superfamily encoded by the nuclear gene SUV3 (YPL029W) (12–14). The activity of this complex is critical for the functioning of the mitochondrial genetic system—inactivating either of the two genes results in respiratory incompetence and a loss of mitochondrial genome stability, with a plethora of phenotypes indicating severe perturbations in all RNA-related processes (6, 12, 15–18). The yeast mitochondrial degradosome displays a remarkably tight functional interdependence of the two subunits (13). The activity of the Dss1p exoribonuclease alone is barely detectable, but is greatly enhanced and becomes entirely ATP-dependent in the presence of Suv3p, making the mtEXO complex an ATPdependent exoribonuclease, which is unique for this group of enzymes. This is evident even with short single-stranded oligoribonucleotide substrates devoid of any secondary structure, suggesting that the role of Suv3 RNA helicase is not restricted to the unwinding of the secondary-structure elements in the substrate that could impede the enzyme’s progress. We, therefore, proposed a model in which the Suv3p helicase acts as a molecular motor feeding the substrate to the catalytic center of the Dss1p ribonuclease (13). The Suv3 protein alone does not display any detectable RNA helicase activity and becomes a 30 to 50 directional helicase requiring a free 30 single-stranded substrate only when Suv3p is in complex with Dss1p. The ATP-dependent RNA-duplex unwinding activity of the Suv3p helicase can therefore only be studied within the whole mtEXO complex. Such interdependence may be a key to explain the frequent failure of demonstrating the helicase activities of DExH/DEAD-box proteins in vitro. It appears that the activity of Suv3p depends on the presence of Dss1p, but not on its activity, as RNA/DNA and even DNA/DNA duplexes are also unwound, even though the DNA strands are not degraded by Dss1p (13). In contrast to the helicase activity, the ATPase activity of Suv3p does not depend on the presence of Dss1p; its background activity in the absence of RNA is, however, greatly reduced when the protein is in the complex (13). It is therefore clear that in order to study the activities of the Suv3p helicase in vitro it is necessary to analyze the entire mtEXO complex, not the isolated protein alone. In this chapter, we describe the methods used in the study of the ATPase, helicase, and exoribonuclease activities of the mitochondrial degradosome complex.

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2. Materials 2.1. Recombinant Proteins and Materials Common to All Protocols

1. All the assays are carried out with the reconstituted recombinant mtEXO complex or the Suv3 and/or Dss1 proteins. The methods for obtaining the recombinant proteins and reconstituting the complex are beyond the scope of this chapter and have been published elsewhere (13, 19). The recombinant proteins are stored at –80C in 10% glycerol, 0.5 M NaCl, 20 mM Tris–HCl (pH 8.0) in small aliquots. Bovine Serum Albumin (BSA) can be used as a negative control. 2. Mid-size vertical SDS-PAGE gel apparatus (10 10 cm glass plates with 0.7 mm spacers) with a suitable power supply. 3. 6 loading dye: 10 mM Tris–HCl (pH 7.6), 0.03% bromophenol blue, 0.03% xylene cyanol FF, 60% glycerol, 60 mM EDTA. 4. 10 TBE buffer: 0.9 M Tris–HCl (pH 8.0), 0.9 M borate, 20 mM EDTA. 5. 10 reaction buffer (RB): 100 mM Tris–HCl (pH 8.0), 250 mM KCl, 100 mM MgCl2. 6. 0.1 M dithiotreitol in water (DTT, store at –20C in small aliquots). 7. General-use autoradiography equipment (we use a digital PhosphorImager).

2.2. Substrate Preparation and Helicase Activity Assays

1. Synthetic dephosphorylated oligoribonucleotides: D – 50 CAAACUCUCUCUCUCUCAAC 30 5 W – 50 AGAGAGAGAGGUUGAGAGAGAGAGAGUUUG 30 3 W – 50 GUUGAGAGAGAGAGAGUUUGAGAGAGAGAG 30 B – 50 GUUGAGAGAGAGAGAGUUUG 30 Deoxyribonucleotide equivalents of the above oligos can also be used in helicase activity assays, as RNA/DNA and DNA/DNA hybrids are also unwound by the yeast mitochondrial degradosome (13). 2. T4 Polynucleotide Kinase (PNK) (New England Biolabs, Ipswich, MA) with the supplied reaction buffer. 3. Radiolabeled g-P32ATP (3000–5000 Ci/mmol). 4. 1:1 acid phenol:chlorophorm mixture. 5. For RNA precipitation: glycogen (MBI Fermentas, Vilnius, Lithuania), 96% ethanol, 3 M Na-acetate (pH 5.2). 6. RNase-free water obtained by DEPC treatment or purchased.

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7. 4 annealing buffer: 80 mM Tris–HCl (pH 8.0), 2 M NaCl, 4 mM EDTA. 8. 30% 19:1 acrylamide/bisacrylamide solution in water; N,N,N’,N’-tetramethyl-ethylenediamine (TEMED); 10% ammonium persulfate solution in water (APS) prepared immediately before use. 9. Autoradiography X-ray film, exposure cassette, developing machine with appropriate chemicals (Agfa). 10. Plastic wrap (Saran), aluminum foil, sharp blades, handheld Geiger counter. 11. Gel extraction buffer: 0.2 M Tris–HCl (pH 7.5), 0.3 M NaCl, 25 mM EDTA, 2% SDS. 12. E. coli tRNA. 13. Proteinase K. 14. 20 mM ATP solution. 2.3. ATPase Activity Assays

1. Plastic-backed PEI-cellulose F TLC plates (Merck). 2. Radiolabeled -P32ATP or g-P32ATP (3000–5000 Ci/ mmol). 3. 1:1 mixture of 0.5 M formic acid and 2 M LiCl. 4. ATP, ADP and AMP solutions (100 mM). 5. 0.5 M EDTA.

2.4. Exoribonuclease Activity Assays

1. Synthetic oligoribonucleotide (5 W) (see Section 2.2, step 1) and materials for T4 PNK labeling with g-P32ATP as described in Section 2.2 2. Denaturing gel stock solutions: 20% 19:1 acrylamide/bisacrylamide solution in 8 M urea and 1 TBE; 8 M urea in 1 TBE. 3. N,N,N’,N’-tetramethyl-ethylenediamine (TEMED); 10% ammonium persulfate solution in water (APS) prepared immediately before use. 4. 20 mM ATP. 5. 2 denaturing loading buffer: 95% formamide, 0.025% SDS, 0.025% bromophenol blue, 0.025% xylene cyanol FF, 0.025% ethidium bromide, 0.5 mM EDTA. 6. (Optional): T7 Transcription Kit (MBI Fermentas, Vilnius, Lithuania), radiolabeled -P32 UTP.

2.5. RNA–Protein Binding Filter Assays

1. Synthetic oligoribonucleotide (5 W) (see Section 2.2, step 1) and materials for T4 PNK labeling with g-P32ATP as described in Section 2.2

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2. Nylon and nitrocellulose membranes (Whatman Nytran N and Protran, respectively, Sigma-Aldrich, St Louis, MO). 3. 0.1 M EDTA. 4. 0.5 M KOH. 5. 1 M KCl. 6. pH indicator strips. 7. 5 binding buffer: 50% glycerol, 25 mM EDTA, 125 mM KCl, 50 mM Tris–HCl (pH 8.0). 8. Dot-blotter (Bio-Dot1SF Microfiltration Apparatus, BioRad, Hercules, CA) with a suitable vacuum source.

3. Methods All experiments involving RNA were carried out in siliconized tubes (Sigma, St Louis, MO) to avoid sticking of nucleic acids to the walls. Most of the protocols involve working with significant amounts of radioactivity, use proper safety equipment, wear protective clothing, and monitor the work area and equipment. Consult with your lab’s radiation safety officer before attempting these procedures for the first time. 3.1. Helicase Activity Assays

In standard helicase assays we use three types of double-stranded RNA substrates. Each of them has a 20 nt double-stranded core structure, optionally with 10 nt single- stranded overhangs on the 50 or the 30 end (Fig. 24.1). The ‘‘bottom’’ strand (D) in all of the substrates was the same, and usually this oligonucleotide was radiolabeled during substrate preparation.Each ofthe top strand oligonucleotides 5 W, 3W, or B, is then annealed to the radiolabeled D oligonucleotide in order to produce the 50 overhang (S5), 30 overhang (S3), or blunt-ended (SB) substrates, respectively (Fig. 24.1a). Reaction products and substrates during preparation are separated on native polyacrylamide gels. As the wild-type mtEXO complex is an active exoribonuclease, time-course assay is required to visualize the strand separation before RNA degradation, as the helicase reaction products are rapidly degraded. Alternatively, a DNA oligonucleotide can be used as the labeled strand, with only minor decrease in the helicase activity (13). Only the double-stranded substrate with the 30 overhang (S3), produced by annealing oligonucleotides D and 3 W is unwound by the helicase activity of the mtEXO complex; the remaining two duplexes are not good substrates and should only be used as negative controls (13) (see also Fig. 24.1c). The Suv3p protein alone does not display detectable duplex-unwinding activity; the presence of the Dss1p ribonuclease subunit is required (13).

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A

B

S5

5W 5’ AGAGAGAGAGGUUGAGAGAGAGAGAGUUUG 3’ D 5’ CAACUCUCUCUCUCUCAAAC 3’

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3W 5' GUUGAGAGAGAGAGAGUUUGAGAGAGAGAG 3' D 5’ CAACUCUCUCUCUCUCAAAC 3’

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C S5 t(min.)

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SB 5

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0.5

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Fig. 24.1. Preparation of substrates and time-course assay of the yeast mitochondrial degradosome helicase activity. (a) Structures of different duplex substrates used in the assay. Oligoribonucleotide D was labeled at the 50 end using T4 PNK. (b) Preparative electrophoresis of substrates after annealing. Note different migration of substrates S5 and S3 in comparison to substrate SB suggesting that the observed bands are indeed doublestranded molecules (see Note 12). (c) A typical time-course assay for the helicase activity of the mtEXO complex using 0.1 mg of recombinant proteins. Reaction products were analyzed on a 15% native polyacrylamide gel. Substrates S5 and SB are not unwound (and only partially degraded), while substrate S3 is unwound after 30 s, and subsequently rapidly degraded. 3.1.1. Substrate Preparation

1. The first step is to obtain the radioactively labeled substrate oligonucleotide (D) (see Note 1). This is achieved by the 50 labeling protocol using the T4 PNK enzyme (New England Biolabs, Ipswich, MA) according to the manufacturer’s instructions. Prepare a 20 mL reaction with 2 mL of 10 PNK buffer (NEB), 1 mL of T4 PNK, 2 pmole of the RNA oligonucleotide, and 2 mL (20 mCi) of radiolabeled gP32ATP. The reaction is carried out at 37C for 45 min. 2. After reaction, remove proteins by phenol:chlorophorm extraction: add equal volume of acidic phenol:chlorophorm solution (see Note 2), vortex vigorously for about 1 min and spin down at maximum speed in a tabletop centrifuge for 5 min. Carefully collect the upper phase (see Note 3). 3. Transfer the water phase to a new eppendorf tube and precipitate RNA with ethanol (see Note 4). To precipitate RNA, add 1/10 of sample volume of 3 M Na-acetate (pH 5.2), 5 mg of glycogen, and two volumes of ice-cold 96% ethanol (see Note 5); incubate for at least 1 h at –20C. 4. After incubation, spin down at maximum speed for 10 min; the pellet is usually hardly visible. Remove ethanol and wash the pellet with 70% ice-cold ethanol (100–200 mL), centrifuge for 1 min at maximum speed, carefully remove ethanol, and air-dry the pellet for 1–2 min (see Note 6).

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5. Dissolve the pellet in 33 mL of RNase-free water and divide the solution into three tubes (11 mL each). 6. Add 4 mL of 4 annealing buffer to each tube and add a proper amount of each unlabeled second-strand oligonucleotide (3 W, 5 W, or B) to obtain a ratio of 1:3 or higher (see Note 7). The final volume should be 16 mL. 7. Heat the samples at 80C for 10 min and then let them cool down slowly to room temperature. You can use a heat block set to 80C, switch it off after 10 min, and wait until the temperature goes down to about 25C (see Note 8). 8. In the meantime prepare a 15% native polyacrylamide gel. To prepare the gel, mix equal volumes of the 30% 19:1 acrylamide/bisacrylamide solution in water and 2 TBE. Add proper amounts of 10% APS (100 mL per 10 mL of gel) and TEMED (8 mL per 10 mL of gel) immediately before pouring. The glass plates should be carefully washed and rinsed with RNase-free water before use. With the 10 10 cm plates and 0.7 mm spacers that we routinely use, about 10 mL of acrylamide solution is sufficient for one gel. Use a comb that will allow at least 20 mL of well volume. 9. Prepare the running buffer (1 TBE). 10. Add the 6 loading dye (see Note 9) to the samples cooled to room temperature and load on the gel. Leave empty wells between each sample—this will make the subsequent steps of band excision easier. Run the gel at 200 V for 1 h (this is for 10 cm gels; adjust to match your equipment). 11. Autoradiography on film (not digital) is necessary for detection at this stage. After electrophoresis, disassemble the apparatus trying to keep the gel intact on one of the glass plates. Prepare the autoradiography cassette by lining it with a piece of aluminum foil. Cover the gel on top of a glass plate with plastic wrap (‘‘Saran’’), and place on the aluminum foil in the exposure cassette (see Note 10). Attach corners of the glass plate to the silver foil with sticky tape to immobilize the gel in the cassette. Subsequent steps will be carried out in the darkroom. 12. Working under the safety light (adapted to your film), cut a sheet of film approximately the size of the gel, and put it on the top of the gel. Mark the film on the edges with a waterproof marker so that you leave markings both on the film and on the aluminum foil (see Note 11). This is done so that you can reposition the developed film on the gel exactly as it was positioned during exposition. Carefully close the cassette and expose for about two minutes. Develop the film.

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13. Using markings on the film and the aluminum foil place the developed film again on the gel, locate the bands corresponding to the labeled product, and cut the corresponding area of the gel with a clean sharp blade, and place in eppendorf tubes (see Note 12 and Fig. 24.1b). 14. Freeze the tubes with cut sections in liquid nitrogen, then grind them carefully with a sterile pipette tip (see Note 13). 15. Add 800 mL of gel extraction buffer to the crumbled gel and rotate overnight at 4C. 16. Spin the samples for 5 min at 4C at maximum speed, collect supernatant as carefully as possible; centrifugation at 4C will also remove some of the SDS from the sample. Try to avoid aspirating any solid pieces; if this happens, repeat the centrifugation step. Check the supernatant and the remaining gel with a Geiger counter to estimate the elution efficiency. 17. Precipitate the RNA by adding glycogen and 2 volumes of icecold 96% ethanol (see Note 14). Incubate the sample at – 20C for an hour, then spin down, and wash the pellet with 70% ethanol as in point 4. Check the pellet radioactivity with a Geiger counter. 18. Calculate the amount of water or 10 mM Tris–HCl (pH 7.5) required to resuspend the samples. You should aim to obtain enough radioactivity per mL to ensure efficient detection of reaction products with your equipment. We usually use substrates at about 50–100 cps/mL (see Note 15). 3.1.2. Time-Course Assay for the Helicase Activity

1. Prepare a 15% native polyacrylamide gel (see Section 3.1.1, step 8) and the running buffer (1 TBE). 2. Prepare the loading dye by supplementing it with carrier tRNA (see Note 16) and proteinase K (1 mg of tRNA and 10 mg of proteinase K per reaction), then aliquot in siliconized eppendorf tubes, one tube for each time point of each assay. As in this assay proteins are in large excess over the RNA substrate, adding unlabeled carrier RNA and proteinase K is necessary to avoid the formation of large protein-substrate complexes that will not migrate into the gel. 3. Prepare the reaction mix for each reaction consisting of 2 mL of 10 reaction buffer (RB), 0.2 mL of 0.1 M DTT, and 1 mL of radiolabeled substrate (50–100 cps, see Section 3.1.1, step 18) in a total volume of 19 mL. Supplement each reaction with 1 mL of 20 mM ATP or water (see Note 17). Reactions without ATP serve as negative controls. Add 0.1 mg of recombinant mtEXO complex; as a negative control, use samples without any proteins or with 0.1 mg BSA. Start timing the reactions. Reactions are carried out at 30C (see Note 18).

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4. At each time point, take out 4 mL of each reaction and immediately add to the tube containing the loading dye with carrier RNA and proteinase K. 5. After the time-course is completed, let the last sample stand in RT for 10 min; then load the samples on the gel. As a positive control to visualize the products of strand separation, prepare one reaction without the proteins and denature by heating to 80C for 5 min and then rapidly cooling on ice for 5 min. Run the gel at 200 V (for a 10 cm gel) for 1 h. 6. Perform the autoradiographic detection procedure appropriate for your equipment. We recommend using a digital phosphor imager system, but photographic film can also be used. We recommend drying the gel before detection (see Note 19). Typical results are presented in Fig. 24.1c. 3.2. ATPase Activity Assays

In the mtEXO protein complex the Suv3p subunit has an ATP hydrolysis activity, hydrolyzing ATP to ADP. This activity is induced by RNA and short single-stranded DNA; the presence of Dss1p subunit does not increase the maximum ATPase activity, it does, however, decrease the basal activity in the absence of the inducing nucleic acid (13). The ATP hydrolysis reaction is monitored using thin layer chromatography on PEI-cellulose. In the conditions used in this assay, ATP remains at the starting point of the chromatography plate while ADP, AMP, and Pi products migrate in the abovementioned order. Radiolabeled -P32ATP or g-P32ATP can both be used as substrates. When -P32ATP is used, radioactive ADP is detected as the reaction product; if the substrate is g-P32ATP, radioactive inorganic phosphate is detected. The substrates are commercially available radiolabeled nucleotides diluted to obtain radioactivity optimal for subsequent detection (300–500 cps/mL). As the resulting ATP concentration is very low, unlabeled ATP can be added to a desired concentration in, for example, enzyme kinetics studies. As the ATPase activity is induced by RNA, unlabeled RNA can be added to the assay as described in the protocol. 1. Prepare substrates for the reaction: dilute commercially available radiolabeled -P32ATP (g-P32ATP can be used as well, see discussion above) to obtain radioactivity in the 300–500 cps/ mL range (about 500 depending on the initial activity and age of the preparation). This usually results in a final concentration of ATP in the reaction in the range of 200 pM. We usually supplement the reactions with unlabeled ATP to the final concentration of 1 mM. Obviously, other final ATP concentrations can be used, for example, in kinetics measurements. 2. Prepare the chromatography standards. Use commercially available unlabeled ATP, ADP, and AMP at a concentration of 100 mM (see Note 20). Verify that at the chosen amount

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the standards are visible on chromatography plates in the UV light before starting the experiment; increase the amount if necessary. 3. Prepare plates for chromatography. We use plastic-backed PEICellulose F from Merck. Cut out a piece about 6–7 cm high and 9 cm wide—this will give sufficient space for about seven samples, if more are desired the plate should be made proportionally wider. The samples are deposited about 1.5 cm from the bottom edge (see Note 21), leave about 1 cm space between each sample. It is convenient to mark the spots for sample loading on the reverse (plastic) side of the plate with a waterproof marker (the markings will be visible on the other side through the plate). 4. Prepare the reaction mix. For one reaction (total volume of 20 mL) add 2 mL of 10 reaction buffer (RB), 0.2 mL of 0.1 M DTT, and 1 mL of diluted radiolabeled substrate (300– 500 cps). These ingredients can be premixed for convenience. Supplement each reaction with 1 mg of inducing RNA (tRNA, total RNA or oligoribonucleotides can be used) or water. Add 0.1 mg of the recombinant mtEXO complex or the Suv3 protein; use samples without proteins or with BSA as negative controls. Incubate the reaction at 30C for 30 min. The reactions can also be run in a time-course format (e.g., in kinetics studies), in such case remove an aliquot at each time point and stop the reaction by adding EDTA to 25 mM. 5. Prepare the chromatography solvents: a 1:1 mixture of 2 M formic acid and 0.5 M LiCl. Dedicated chromatography chambers are not necessary; if they are not available any vessel of a suitable size can be used, for example, a plastic culture dish. Prepare enough solvent for the vessel used. 6. Pre-run the plate with water. Fill the vessel with water (ddH2O) just to cover the bottom (water and solvent levels must be below the spots where the samples will be deposited). Put your plate with the bottom edge in the water and support it so that it remains nearly vertical. Water will move up the plate by capillary action, when it gets close to the upper edge of the plate (this should take about 5 min) you can begin to prepare the samples (see Note 22). 7. Stop the reactions by adding EDTA to 25 mM and placing the samples on ice. 8. Remove the pre-run plate from the vessel and put it on the bench with the silica side up. Load 1 mL of each sample on the places marked with spots; try not to touch the silica layer with the tip during loading (see Note 23). Then load the unlabeled ATP, ADP, and AMP standards (1 mL each). Remove water from the vessel and add the solvent mixture. Place the plate back in the vessel.

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9. Chromatography takes 10–20 min and can be monitored under the UV light by the migration of the markers. After the separation is complete, the plate should be dried—we use a 50C oven for 10 min. 10. Wrap the plate with transparent plastic foil (Saran) and visualize by autoradiography on film or on a phosphor imager screen. Typical results are shown in Fig. 24.2a.

A

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Fig. 24.2. Assays for the ATPase, exoribonuclease and RNA-binding activities of the yeast mitochondrial degradosome. (a) Chromatographic assay of the ATPase activity of the mtEXO complex. -P32ATP was used as substrate with 0.1 mg of recombinant proteins in the presence of RNA, DNA, or no nucleic acid (–). BSA was used as a negative control. Direction of chromatography is marked by an arrow. (b) Time-course assay of the exoribouclease activity using 50 labeled 30 nt oligoribonucleotide 5 W as substrate (S, 2 mM) with 0.1 mg of recombinant proteins. The reactions were separated on a 15% denaturing polyacrylamide gel. The product (P) is a short residual 4 nt fragment that is left after mtEXO digestion of the substrate. (c) Double filter assay of the RNA-binding capability of the mtEXO complex. The 5 W RNA oligonucleotide labeled on the 50 end was used at the concentration of 87.5 pM (1.75 fmol per 20 mL reaction) with a series of twofold dilutions of the mtEXO complex. Results for increasing protein concentration from 9.45 to 605 nM, along with the negative control (BSA) are shown.

3.3. Exoribonuclease Activity Assays

The Dss1 protein is the exoribonuclease of the mtEXO complex. Its activity is greatly enhanced by the helicase activity provided by the Suv3p subunit—the activity of Dss1p alone is barely detectable (13). It digests RNA in the 30 to 50 direction, producing nucleoside monophosphates. There are many possible assays that can be performed to analyze various aspects of the exoribonuclease activity of mtEXO; most of them are, however, beyond the scope of this volume. In this protocol, we will only introduce a method to perform a time-course assay of the mtEXO exoribonuclease

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reaction using a short oligonucleotide substrate (30 nt in this example), labeled at the 50 end, and separation of the products on a denaturing polyacrylamide gel. Like other enzymes in the RNB family, mtEXO leaves a short residual undigested core of four nucleotides (13), which is easily detected using the described protocol. Longer substrates can be obtained by in vitro transcription of appropriate DNA constructs using -P32UTP labeling with subsequent detection of degradation by-products and released UMP. In such cases, PEI-cellulose thin layer chromatography can also be used to monitor the reaction in conditions described in protocol 3.2 (undigested RNA will remain at the bottom of the chromatogram, while the released UMP will migrate upwards). Different formats of the exoribonuclease assay have been described in our previous papers (13, 19). 1. Label the substrate. Use an oligoribonucleotide substrate like oligo 5 W used in previous assays. Label the oligo at the 50 end with T4 PNK and g-P32ATP exactly like it is described in Section 3.1.1, step 1. After the completion of the reaction, add 2X loading dye for denaturing gels, heat to 65C for 10 min, and immediately put on ice for 5 min. Denaturation should be performed just prior to electrophoresis; the labeled undenatured substrate can be stored at –20C for a few days. 2. Prepare a 15% denaturing polyacrylamide gel by mixing two volumes of 20% 19:1 acrylamide/bisacrylamide solution in 8 M urea and 1 TBE with one volume of 8 M urea in 1 TBE. Add proper amounts of 10% APS (100 mL per 10 mL of gel) and TEMED (8 mL per 10 mL of gel) immediately before pouring. With the 10 10 cm plates and 0.7 mm spacers that we routinely use, about 10 mL of acrylamide solution is sufficient for one gel. Run the gel for 1 h at 200 V (for 10 cm gels). 3. Locate and purify the radioactively labeled oligonucleotide exactly as described in Section 3.1.1, steps 11–18. 4. Prepare another 15% denaturing polyacrylamide gel as described in point 2 above and pre-run it. 5. Prepare the 2 loading dye for denaturing gels supplemented with 1 mg of carrier RNA (tRNA or total yeast RNA) per reaction and aliquot it into siliconized eppendorf tubes (4 mL for each tube). Carrier RNA is added to avoid the formation of protein-substrate complexes that will remain in the wells (this can be a problem even with denaturing gels) and to prevent further hydrolysis of the substrate (which should be negligible due to the presence of EDTA in the loading buffer). 6. Prepare the reaction mix. For one reaction (total volume of 20 mL) add 2 mL of 10 reaction buffer (RB), 0.2 mL of 0.1 M DTT, 1 mL of 20 mM ATP, and 1 mL of radiolabeled substrate

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(50–100 cps). Add 0.1 mg of recombinant mtEXO complex; as a negative control, use samples without any proteins or with 0.1 mg BSA. Start timing the reactions. Reactions are carried out at 30C. 7. At each time point, take out 4 mL of each reaction and immediately add to the tube containing the loading dye with carrier RNA and place on ice. 8. After the time-course is completed, denature the samples by heating to 65C for 10 min and immediately transfer to ice and incubate for further 5 min. Load the samples on the prepared and pre-run 15% denaturing polyacrylamide gel and electrophorese for about 1 h at 200 V (for a 10 cm gel). 9. After electrophoresis dry the gel, wrap with transparent plastic foil (Saran), and visualize by autoradiography on film or on a phosphor imager screen. Typical results are shown in Fig. 24.2b. Decreasing amounts of substrate and the production of the 4 nt residual core left after exoribonucleolytic digestion should be readily detectable. 3.4. RNA–Protein Binding Filter Assays

For investigating the ability of the mtEXO complex to bind various RNA molecules, we use a variant of the double filter assay developed by Wong and Lohman (20) and adapted by Tanaka and Schwer (21). In our experiments we generally follow the protocol described by Vincent and Deutsher (22). The method, in general, is based on the separation of free RNA molecules from RNA bound to proteins. Radiolabeled RNA is incubated with the proteins and subsequently passed through a nitrocellulose filter and then through a nylon filter. Proteins with bound RNA should be retained on the first (nitrocellulose) filter, while the free unbound RNA should be retained on the second (nylon) filter. Quantitative autoradiography of both filters allows the estimation of the amount of bound vs. unbound RNA, and the calculation of the bound fraction. Varying the amount of protein (conveniently by a series of twofold dilutions) and plotting the bound fraction against the protein concentration makes estimation of the binding constant (Kd) possible. In this protocol, we provide the method using a short oligoribonucleotide substrate (5 W) labeled on the 50 end; longer polynucleotide substrates obtained by in vitro transcription of appropriate DNA constructs using -P32UTP labeling can also be used with this method (13). 1. Label the oligoribonucleotide substrate (5 W) and purify it as described in Section 3.3, steps 1–3. 2. Prepare the solutions for filter presoaking: 0.1 M EDTA; 1 M KCl; 0.5 M KOH.

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3. Prepare the 1 binding buffer (BB) using the 5 stock, add DTT to 1 mM (see Note 24). 4. Cut both filter membranes to a proper size determined by the dot-blotting equipment used (we use the Bio-Dot1SF Microfiltration Apparatus, Bio-Rad, Hercules, CA; but any similar device should work). 5. Presoak the membranes (see Note 25) as follows. For the nylon membrane: 10 min in 0.1 M EDTA, followed by three times in 1 M KCl for 10 min each, then 1 min in 0.5 M KOH and finally rinse several times with H2O until the pH returns to neutral (see Note 26). For the nitrocellulose membrane: 10 min in 0.5 M KOH and then rinse several times with H2O until the pH returns to neutral. After presoaking incubate both filters in the binding buffer at 4C for at least one hour before proceeding with the experiment. 6. Prepare the reaction mixtures. For one reaction (total volume of 20 mL) add 4 mL of 5 binding buffer, 0.2 mL 0.1 M DTT, and a proper amount of protein (generally from about 5 nM to about 1200 nM, 200–600 nM should give strong binding to oligoribonucleotide substrates). Add the radioactive substrate (we use 1.75 fmol per 20 mL reaction, which gives 87.5 pM final RNA concentration). 7. Incubate the reactions for about 10 min at 30C. Exact timing is not very important as the sample-loading step usually takes much longer when many samples are processed in parallel. 8. Assemble the dot blotter equipment, from the bottom: sealing gasket, two (or more depending on the equipment) sheets of Whatman 3 MM paper soaked in the binding buffer, the nylon membrane, then the nitrocellulose membrane, and finally the sample loading template. Secure with screws according to the manufacturer’s instructions. Attach the vacuum pump and verify if the apparatus is properly assembled and sealed by loading one well with buffer and checking if it is quickly passed through the membranes. 9. Load the samples one at a time. Immediately before loading each sample, wash the well with 100 mL of binding buffer, and as soon as the wash passes through the membranes, load the entire 20-mL sample of the binding reaction (see Note 27). Repeat the washing step with another 100 mL of binding buffer. Proceed to the next sample. With many samples the whole procedure will take considerable amounts of time, but it is critical that the initial wash, loading of the sample, and the final wash are done quickly in succession. Delays at this stage can contribute to a high nonspecific background.

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10. After loading the samples, wait until all the buffer has passed through the membranes, disassemble the dot-blotter, and airdry the membranes. Wrap the dried membranes with transparent plastic foil (Saran) and visualize by autoradiography on film or on a phosphor imager screen (see Note 28). Typical results are shown in Fig. 24.2c. 11. Bound RNA fraction is estimated as the product of the signal on the nitrocellulose filter and the total signal on both filters. Kd can be estimated by a nonlinear regression fit of the data to the formula B = [P]/(Kd + [P]), where B is the bound RNA fraction and [P] is the protein concentration (see Note 29).

4. Notes 1. We recommend ordering synthetic dephosphorylated oligonucleotides, otherwise a dephosphorylation step is needed prior to 50 end labeling 2. As the volume of the sample is small (20 mL); if you decide to perform phenol:chlorophorm extraction, you can dilute the sample with water to a convenient volume (generally few hundred mL)—it will make the procedure much easier and minimize the loss of material. 3. If the labeled ATP is marked with a color dye, it can make this step easier as the dye will partition in the organic phase (not all dyes used by different suppliers will necessarily exhibit this behavior). 4. As the buffer used in the next step (RNA annealing) has a very high salt concentration, the precipitation step at point 3 can be skipped and the annealing reaction can be performed directly with the oligonucleotide obtained after phenol:chlorophorm extraction. 5. In each RNA precipitation step we use glycogen and siliconized tubes to avoid sticking of nucleic acids to tube walls, but even then it remains a problem which causes significant loses of material—after precipitation, up to 50% of total radioactivity can remain on the tube walls after dissolving the pellet. 6. The pellet formed during precipitation of radiolabeled RNA oligonucleotides is usually hardly visible, but its presence can be verified using a Geiger counter. Do not discard the supernatants from the precipitation steps immediately; when monitoring the pellet and supernatants with a Geiger counter, you can notice when the pellet becomes accidentally unstuck and is removed with the supernatant. Recover it by reprecipitation.

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7. Estimating the amount of labeled oligonucleotide is difficult (it depends on the labeling and purification yield); we usually use 2 pmole of each unlabeled second-strand oligonucleotide for annealing reaction. It is not critical, as the subsequent gel purification will get rid of any not-annealed strands. 8. If the design of the heating block allows it, you can remove the block from the heater and place it on the bench to accelerate the cooling. 9. The 6 loading buffer can be prepared using the formula given in Section 2 but we used commercial DNA loading buffer supplied by MBI Fermentas, Vilnius, Lithuania. For the RNA work, choose a fresh unopened tube of the buffer and store it at –20C. 10. At this point you should use a Geiger counter to check the efficiency of your preparation; electrophoresis removes the unincorporated nucleotides, so the signal from the gel should correspond to the amount of labeled oligonucleotide. It should be strong enough to exceed the scale of most handheld Geiger counters, if the signal is weak there is no point in continuing the procedure and the labeling and annealing steps should be repeated. 11. Before this step, you should check if the ink of your waterproof marker survives the film development—mark a spare piece of film, run it through the developer, and see if the markings are still visible. 12. Using different second-strand unlabeled oligonucleotides of various lengths (like our oligonucleotides B and 5 W or 3 W) in the same experiment provides an additional control of the annealing reaction. The resulting double-stranded products will migrate differently on the gel (Fig. 24.1b), depending on the size of the second strand; if annealing fails, the labeled oligonucleotide will always migrate at the same level. In our experience the annealing step is very reliable and further controls are not necessary. 13. Before grinding the frozen gel slice wait until it is slightly thawed; a hard frozen piece of polyacrylamide can jump uncontrollably in the test tube when you attempt to grind it. 14. As the elution buffer contains 0.3 M NaCl, no further addition of salt is required for precipitation. SDS should not precipitate under these conditions. 15. When estimating the amount of water needed for resuspension of the pellet, start with half the intended volume and check the radioactivity of 1 mL of the solution using a Geiger counter. A significant portion of radioactivity can remain in insoluble form. (see also Note 5).

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16. You can use different carrier RNAs for this purpose; we used commercial E. coli tRNA or total yeast RNA preparations with success. Without the carrier RNA, the majority of reaction products remain in the wells associated with proteins and do not migrate into the gel. The same suggestion also applies to denaturing polyacrylamide gels used in exoribonuclease assays. 17. For performing multiple assays in parallel, the most convenient way is to prepare a master premix (without ATP, RNA substrate, and proteins) for all the samples. ATP or water (control) can be placed in the reaction tubes (1 mL of 20 mM ATP/water each). Then add an appropriate amount of the radioactive RNA substrate to the master premix and aliquot to each of the reaction tubes (18 mL each). Finally add 1 mL of the recombinant enzyme, vortex, and start the incubation at 30C. 18. You can use ‘‘trap’’ RNA (0.5 pmole of unlabeled oligoribonucleotide D) in each reaction to avoid strand reannealing. It is not required when working with the wild-type mtEXO complex, as the exoribouclease activity will quickly degrade the released strands anyway. It can be useful when working with mutants deficient in the exoribonuclease activity or with substrates that do not undergo degradation by mtEXO (like DNA molecules). 19. Drying high-percentage polyacrylamide gels can sometimes be problematic (gel crumbling) depending on the equipment available. If this happens you can try to make exposure with wet gels using X-ray film (using wet gels in phosphor imager cassettes is not recommended). This will work if the signal is strong enough to allow detection with short exposure time (not more than a few hours), otherwise the bands will become blurred due to diffusion. 20. If ADP and AMP solutions are not available, the standards can be prepared by leaving an aliquot of the ATP solution at room temperature overnight. Spontaneous hydrolysis should produce enough ADP and AMP to serve as chromatography standards (verify by chromatography and UV light detection before the actual experiment). 21. The exact distance from the edge is not critical, it is important to make sure the samples are deposited above the level of the solvents in the vessel. 22. The important part is to make sure that the chromatographic plate does not become dry during the experiment. After the pre-run, it is therefore critical to load the samples quickly and replace the plate in the vessel with the proper solvent mixture before it dries out. This is why it is convenient to get the samples ready for loading during the pre-run.

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23. The silica on the plate is very fragile when wet and touching it with a tip can result in scratches that will disrupt the proper development of the chromatogram. 24. The main difference between the binding and reaction buffers is the lack of magnesium and presence of EDTA in the former. The reason is to avoid degradation of the RNA substrate by the exoribonuclease activity of mtEXO. In the absence of divalent cations, the exoribonuclease should not be active. 25. Presoaking the membranes is a critical step in this protocol and its omission will lead to a very high nonspecific background, particularly on the nitrocellulose filter which without presoaking will bind free RNA not bound to proteins. 26. We use pH indicator strips to check the neutrality; touch the strip to the filter in the area where no samples will be loaded. 27. When loading the sample try to place it at the bottom of the well without touching the membrane. Try to avoid air bubbles and sticking of the sample to the walls of the well. If this happens, touch the sample with the tip and bring it to the bottom of the well. It is important that the sample is passed through the membranes quickly and in its entirety. 28. Mark the membranes (e.g., by making notches) to avoid mixing them during the experiment and analysis. 29. If the experiment is performed with Kd estimation in mind, it is important to ensure that both ends of the binding curve are included in the range of concentrations used: the bound RNA fraction for the highest protein concentrations should approach 95–100% and should not increase with further protein concentration increase. On the other end, the bound RNA fraction for the lowest protein concentrations should approach that of a negative control (e.g., BSA). When the experiment is performed properly, this should not exceed 1%.

Acknowledgments This work was supported by the Ministry of Science and Higher Education of Poland through The Faculty of Biology, Warsaw University Intramural Grants BW#1720/46 and BW#1680/40, the CoE BioExploratorium project: WKP_1/1.4.3/1/2004/ 44/44/115/2005, and by grants 2P04A 002 29 and N N301 2386 33 from the Ministry of Science and Higher Education of Poland.

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References 1. 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. 2. Meyer S., Temme C., and Wahle E. (2004) Messenger RNA turnover in eukaryotes: pathways and enzymes. Crit. Rev. Biochem. Mol. Biol. 39, 197–216. 3. Mitchell P. and Tollervey D. (2000) mRNA stability in eukaryotes. Curr. Opin. Genet. Dev. 10, 193–198. 4. Mitchell P. and Tollervey D. (2001) mRNA turnover. Curr. Opin. Cell Biol. 13, 320–325. 5. Newbury S. F. (2006) Control of mRNA stability in eukaryotes. Biochem. Soc. Trans. 34, 30–34. 6. Rogowska A. T., Puchta O., Czarnecka A. M., Kaniak A., Stepien P. P., and Golik P. (2006) Balance between transcription and RNA degradation is vital for Saccharomyces cerevisiae mitochondria: reduced transcription rescues the phenotype of deficient RNA degradation. Mol. Biol. Cell 17, 1184–1193. 7. Carpousis A. J. (2002) The Escherichia coli RNA degradosome: structure, function and relationship in other ribonucleolytic multienzyme complexes. Biochem. Soc. Trans. 30, 150–155. 8. Mitchell P., Petfalski E., Shevchenko A., Mann M., and Tollervey D. (1997) The exosome: a conserved eukaryotic RNA processing complex containing multiple 30 !50 exoribonucleases. Cell 91, 457–466. 9. Zuo Y. and Deutscher M. P. (2001) Exoribonuclease superfamilies: structural analysis and phylogenetic distribution. Nucleic Acids Res. 29, 1017–1026. 10. Cordin O., Banroques J., Tanner N. K., and Linder P. (2006) The DEAD-box protein family of RNA helicases. Gene 367, 17–37. 11. Rocak S. and Linder P. (2004) DEAD-box proteins: the driving forces behind RNA metabolism. Nat. Rev. Mol. Cell Biol. 5, 232–241. 12. Dziembowski A., Piwowarski J., Hoser R., Minczuk M., Dmochowska A., Siep M., van der Spek H., Grivell L., and Stepien P. P. (2003) The yeast mitochondrial degradosome. Its composition,

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interplay between RNA helicase and RNase activities and the role in mitochondrial RNA metabolism. J. Biol. Chem. 278, 1603–1611. Malecki M., Jedrzejczak R., Stepien P. P., and Golik P. (2007) In vitro reconstitution and characterization of the yeast mitochondrial degradosome complex unravels tight functional interdependence. J. Mol. Biol. 372, 23–36. 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. Dmochowska A., Golik P., and Stepien P. P. (1995) The novel nuclear gene DSS-1 of Saccharomyces cerevisiae is necessary for mitochondrial biogenesis. Curr. Genet. 28, 108–112. Dziembowski A., Malewicz M., Minczuk M., Golik P., Dmochowska A., and Stepien P. P. (1998) The yeast nuclear gene DSS1, which codes for a putative RNase II, is necessary for the function of the mitochondrial degradosome in processing and turnover of RNA. Mol. Gen. Genet. 260, 108–114. Golik P., Szczepanek T., Bartnik E., Stepien P. P., and Lazowska J. (1995) The S. cerevisiae nuclear gene SUV3 encoding a putative RNA helicase is necessary for the stability of mitochondrial transcripts containing multiple introns. Curr. Genet. 28, 217–224. Stepien P. P., Margossian S. P., Landsman D., and Butow R. A. (1992) The yeast nuclear gene suv3 affecting mitochondrial post-transcriptional processes encodes a putative ATP-dependent RNA helicase. Proc. Natl. Acad. Sci. U.S.A. 89, 6813–6817. Malecki M., Jedrzejczak R., Puchta O., Stepien P. P., and Golik P. (2008) In vivo and in vitro approaches for studying the yeast mitochondrial RNA degradosome complex. Methods Enzymol. 447, 463–488. Wong I. and Lohman T. M. (1993) A double-filter method for nitrocellulose-filter binding: application to protein-nucleic acid interactions. Proc. Natl. Acad. Sci. U.S.A. 90, 5428–5432.

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21. Tanaka N. and Schwer B. (2005) Characterization of the NTPase, RNA-binding, and RNA helicase activities of the DEAH-box splicing factor Prp22. Biochemistry 44, 9795–9803.

22. Vincent H. A. and Deutscher M. P. (2006) Substrate recognition and catalysis by the exoribonuclease RNase R. J. Biol. Chem. 281, 29769–29775.

Chapter 25 Characterization of the Helicase Activity and Anti-telomerase Properties of Yeast Pif1p In Vitro Jean-Baptiste Boule´ and Virginia A. Zakian Abstract Pif1p is the prototype member of a family of helicases that is highly conserved from yeast to humans. In yeast, Pif1p is involved in many aspects of the preservation of genome stability. In particular, Pif1p is involved in the maintenance of mitochondrial DNA and in the direct inhibition of telomerase at telomeres and double-stranded breaks. Here we describe methods to purify Pif1p and study in vitro its enzymatic properties and functional interaction with telomerase. Key words: Yeast, Pif1p, helicase, telomerase, oligonucleotide substrate-based radiometric assays.

1. Introduction This chapter focuses on the characterization of the Saccharomyces cerevisiae Pif1p helicase. Pif1p is the prototype member of the PIF1 family of helicases, which is conserved from yeast to human (1, 2). Two isoforms of the enzyme are expressed in yeast, owing to alternative usage of two start codons from the same mRNA. Translation from the first start codon leads to the synthesis of a mitochondria-directed isoform, while translation from the second AUG codon leads to the synthesis of the nuclear isoform (3). Genetic studies have shown that the nuclear form of Pif1p plays an important role in counteracting the activity of telomerase, the specialized reverse transcriptase that elongates the end of eukaryotic chromosomes. Through this activity, Pif1p prevents gross chromosomal rearrangements that are due to the addition of telomerase-mediated de novo telomere addition at double strand breaks (4). In vivo and in vitro data suggest that this action is M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_25, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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achieved through a direct interaction between Pif1p and telomerase (5, 6). Using oligonucleotide-based radiometric assays, Pif1p has been shown to unwind preferentially RNA–DNA hybrids over DNA substrates (7). This preference suggests that Pif1p inhibits telomerase by unwinding the RNA–DNA substrate formed by the telomerase RNA, TLC1, and the telomeric DNA end. Importantly, the interaction between Pif1p and telomerase is conserved in evolution, since Pif1p has been shown to interact with mouse and human telomerase (8–10). This chapter focuses on in vitro methods to purify recombinant yeast Pif1p and to characterize its activity by classical oligonucletide substrate-based radiometric assays. We also describe methods to study in vitro its functional interaction with yeast telomerase.

2. Materials 2.1. Overexpression of Recombinant His-Tagged Pif1p in Bacteria

1. Luria Bertani (LB) (per liter): 10 g Bacto-tryptone, 5 g yeast extract, 10 g NaCl. Adjust pH to 7.5 with 1 N NaOH. Autoclave. 2. Isopropyl-b-thiogalactoside (IPTG): 1 M solution in ddH2O. Filter-sterilize and store at –20C in 1 mL aliquots. 3. Kanamycin sulfate stock solution: make stock at 50 mg/mL in ddH2O. Filter sterilize. Store at 4C. 4. Chloramphenicol stock solution: make stock at 50 mg/mL in 100% ethanol. Store at –20C.

2.2. Purification of Recombinant His-Tagged Pif1p 2.2.1. Affinity Chromatogaphy

1. Buffer A: 30.5 mM Na2HPO4, 19.5 mM NaH2PO4, 300 mM NaCl, pH 7.0. Filter sterilize. 2. 1 M imidazole: pH 7.0. Adjust pH with 1 N NaOH and filter sterilize. 3. Protease inhibitor cocktail tablets, EDTA free (Roche Applied Science). 4. 45 mM sterile syringe filter; 10 mL sterile plastic syringe. 5. Sonicator and thin probe. 6. Talon polyhistidine-Tag purification resin (Clontech) (see Note 1). 7. Low pressure chromatography column (5 mL capacity or above, e.g., Millipore Vantage-L chromatography column or equivalent). 8. Peristaltic pump (Gilson Minipuls, or equivalent) and silicone tubing.

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9. Low protein binding tubes (e.g., Nunc MiniSorp). 10. Dialysis tubing (e.g., Pierce Snakeskin, 10 kD cut off) and clips (Pierce). 11. Anti-His-Tag monoclonal antibody (Novagen). 2.2.2. Cation Exchange Chromatography

1. Buffer A2: 45.5 mM Na-acetate, 4.5 mM acetic acid, 50 mM (NH4)2SO4, 50 mM Mg-acetate, 200 mM NaCl, 5% glycerol, pH 5.6. Filter sterilize. 2. Buffer B2: Same composition as buffer A2 but containing NaCl 1 M, pH 5.6. Filter sterilize. 3. Buffer C: HEPES 50 mM pH 7.8, (NH4)2SO4 50 mM, Mgacetate 50 mM, 200 mM NaCl, DTT 1 mM, glycerol 5%. Filter sterilize. 4. Pif1p storage buffer: 0.5 buffer C containing 50% glycerol. 5. FPLC system (GE healthcare Akta system or equivalent). 6. Centrifugal filter concentrator, 10 kD cut off (Millipore Centricon YM-10 or equivalent).

2.3. Preparation of Telomerase Activity from Yeast Protein Extracts 2.3.1. Overexpression of Telomerase Core Subunits

1. Complete SCGLA medium (per liter): 6.7 g of yeast nitrogen base without amino acids, 1 g glucose, 30 g glycerol, 20 g lactic acid, 20 mg of each adenine, uracil, histidine, tryptophan, proline, arginine, and methionine; 30 mg of each leucine, isoleucine, tyrosine, and lysine. Adjust pH to 5–6 with NaOH 10 N and autoclave. 2. YPGLA medium (per liter): 10 g yeast extract, 20 g peptone, 2 g glucose, 30 g glycerol, 20 g lactic acid, 20 mg adenine. Adjust to pH 5–6 and autoclave. 3. D(þ)-galactose (Sigma). 4. Diethylpyrocarbonate (DEPC)-treated ddH2O. Add 0.5 mL DEPC to 500 mL ddH2O. Mix well. Incubate at 37C overnight. Sterilize by autoclaving. 5. Buffer L: 40 mM Tris–HCl, pH 8.0, 500 mM sodiumacetate, 2.2 mM MgCl2, 0.2 mM EDTA, 0.2% Triton X-100, 0.4% Igepal CA-630, 20% glycerol. Prepare using RNase-free reagents and glassware (see Note 2).

2.3.2. Telomerase Activity Fractionation and Storage

As telomerase activity is RNase-sensitive, all reagents, solutions, and equipment must be handled in an RNase-free environment (see Note 2). 1. Automated mortar and Pestle (e.g., Retsch RM100). 2. RNaseZAP solution (Ambion). 3. 50 mL sterile conical polypropylene tubes. 4. Sterile single-usage plastic pipettes.

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5. RNase inhibitor (e.g., Promega RNAsin). 6. DEAE sepharose fast flow (GE healthcare). 7. TMG-500 buffer: 10 mM Tris–HCl, pH 8.0, 1.1 mM MgCl2, 500 mM Na-Acetate, 0.1 mM EDTA, 0.1% Triton X-100, 0.2% Igepal CA-630, 10% glycerol, 0.1 mM phenylmethanesulphonylfluoride (PMSF). 8. TMG-900 buffer: same as above, but containing 900 mM Na-Acetate. 9. TMG-30 buffer: same as above, but containing 30 mM Na-Acetate. 10. PD-10 desalting column (GE healthcare). 11. Centrifugal filter concentrator (e.g., 5 mL centricon YM-30, Millipore). 12. Glycerol (Sigma). 2.4. Characterization of Pif1p Activity Using Radiolabeled Oligonucleotide Substrates 2.4.1. Preparation of Oligonucleotide Substrates

1. T4 polynucleotide kinase (New England Biolabs). 2. [g-32P]-ATP (> 5000 Ci/mmol). 3. Annealing buffer (5 ): 10 mM Tris–HCl, pH 7.5, 10 mM MgCl2. 4. Ficoll loading buffer (6 ): 17% Ficoll, 0.05% bromophenol blue, 0.05% xylene cyanol. 5. Polyacrylamide gel oligonucleotides.

electrophoresis

(PAGE)-purified

6. Microspin G-25 columns (GE healthcare). 7. 12% polyacrylamide gel (20:1 acrylamide:bis-acrylamide ratio): To prepare 100 mL of gel, mix 28.5 mL 19:1 acrylamide:bis-acrylamide 40% solution (Bio-Rad), 1.45 mL acrylamide 40% solution (Bio-Rad), 10 mL TBE 10 , 60 mL ddH2O. Filter and degas. Per mini-gel, mix 10 mL of gel mix with 100 mL of APS 10% and 10 mL of N,N,N 0 ,N 0 tetramethylethylenediamine (TEMED). Poor the solution to make a 1.5-mm thick 10 8 cm mini-gel. 8. Microcon Ultrafree-MC centrifugal filters (0.22 mm, Millipore). 9. D-Tube midi Dialyzer tubes (Novagen). 10. Microcon YM-10 centrifugal filter unit (Millipore). 11. Horizontal agarose gel slab unit and power supply. 12. Sterile razor blades. 13. Kodak autoradiography film and cassette. 14. Scintillation cocktail and 20 mL scintillation vials. 2.4.2. Pif1p Activity Assay

1. Helicase reaction buffer (5 ): 100 mM Tris, pH 7.5, 200 mM NaCl, 500 mg/mL BSA, 10 mM dithiotreitol (DTT).

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2. Helicase stop/load buffer: Ficoll loading buffer (see Section 2.4.1) supplemented with 50 mM EDTA and 1 mM single stranded DNA oligonucleotide (see Note 3). 3. 12% polyacrylamide gel (20:1 acrylamide:bis-acrylamide ratio): Per gel, mix 40 mL of gel mix with 0.2 mL of APS 10% and 40 mL of TEMED. Poor the solution to make a 1.5-mm thick 18 16 cm gel. 4. Vertical slab gel electrophoresis unit and power supply. 5. Kodak autoradiography film (BioMax MR) and film cassette. 6. Radiolabeled oligonucleotide helicase substrate (see Section 2.4.1). 7. 3 MM and DE81 chromatography papers (Whatman). 2.5. Functional Interaction Assay Between Pif1p and Yeast Telomerase 2.5.1. Telomerase Assay

1. Telomerase Reaction (TR) buffer (10 ): 200 mM Tris, pH 8.0, 200 mM NaCl, 10 mM DTT, 10 mM spermidine. 2. 50 mM MgCl2. 3. dNTP mix: 0.5 mM of each dATP, dGTP, and dCTP, 50 mM dTTP, 40 mM ATP. 4. [a-32P]-TTP (>5000 Ci/mmol). 5. TR stop buffer: 20 mM Tris, pH 8.0, 1 mM EDTA, 0.5% SDS, 250 mg/mL PCR-grade proteinase K, 50 pM [g-32P]labeled control oligonucleotide (see Note 4). 6. Glycogen. 7. (NH4)-Acetate 4 M. Filter sterilize. 8. Formamide loading buffer: 10 mM NaOH, 95% formamide, 0.05% bromphenol blue, 0.05% xylene cyanole.

2.5.2. Electrophoresis and Detection of Reaction Products

1. 16% urea-polyacrylamide sequencing gel. To prepare 1 L of gel mix, add successively in a beaker 100 mL TBE 10 , 400 mL 19:1 acrylamide:bis-acrylamide 40% solution (BioRad) and 420 g urea. Mix by stirring on medium heat until urea is dissolved. Let cool to room temperature, poor into a 1 L graduated cylinder, and complete to 1 L with ddH2O. Mix, filter through a paper filter and degas. To prepare one gel, mix 100 mL of 16% gel mix, 0.5 mL APS 10%, and 100 mL TEMED. 2. Nucleic acid sequencing unit (Sigma IBI model STS-45 is recommended) and power supply.

2.5.3. Telomerase Displacement Assay

All buffers and material should be RNase-free (see Note 2). 1. Dynabeads M-280 streptavidin magnetic beads (Invitrogen). 2. PAGE-purified biotinylated telomeric oligonucleotide.

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3. Magnet. 4. Buffer S1: 0.1 M NaOH, 0.05 M NaCl. 5. 0.1 M NaCl. 6. Buffer B/W: 10 mM Tris–HCl (pH 7.5), 1 mM EDTA, 2 M NaCl. 7. TR buffer: see Section 2.5.1.

3. Methods 3.1. Overexpression of Recombinant Pif1p in Bacteria

3.2. Purification of Recombinant Pif1p 3.2.1. Affinity Chromatography

1. These instructions assume the use of a bacterial strain expressing the Pif1p nuclear isoform (amino acids 40–859) fused to a 6-histidine tag. As an example, we use the BL21(DE3) derivative strain Rosetta (Novagen) expressing the PIF1 ORF (minus the first 117 base pairs) cloned into the pET28(b) vector (Novagen), in order to express, upon IPTG induction, Pif1p fused at its N-terminus to a 6-histidine tag (see Note 5). Fresh colonies are grown overnight on a LB plate containing 30 mg/mL kanamycin and 34 mg/mL chloramphenicol. Inoculate a colony in 50 mL LB supplemented with kanamycin and chloramphenicol and grow at 37C overnight with agitation (150 rpm). Inoculate 5 mL of the overnight culture in 1 L of LB and grow cells at 37C with agitation until OD600 reaches 0.6–0.8 (see Note 6). Cool the bacterial culture for 30 min by placing it in an ice bucket. After the culture has cooled down, add IPTG at 1 mM final concentration and incubate the culture at 18–23C with agitation for another 15 h (see Note 7). Pellet the bacteria by centrifugation at 4000 g and freeze the pellet at –80C, or keep on ice and proceed to purification (see Section 3.2). All steps should be performed at 4C. 1. Resuspend cell pellet in 1/20th culture volume of cold buffer A containing protease inhibitors (one Complete EDTA-free protease inhibitor tablet per 30 mL buffer A). Cells are broken by a single passage in a French press at 10,000 PSI. The lysate is subsequently cleared by centrifugation for 30 min at 16,000 g at 4C. At this stage, the supernatant is still viscous due to presence of bacterial nucleic acids. The supernatant is transferred to a beaker and sonicated on ice using a thin probe at the following settings: 40% amplitude, pulse 40% (see Note 8). Continue until the lysate is no longer viscous. Filter the supernatant through a 45-mM syringe filter to remove remaining aggregates.

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2. Load the supernatant on a column containing 5 mL packed Talon resin equilibrated in buffer A using a peristaltic pump (see Note 9). Load at a constant flow rate of about 3 mL/min or slower. Faster flow rates tend to decrease binding of recombinant Pif1p to the resin. After loading the supernatant, wash the column successively with 3 volumes of buffer A supplemented with 30 mM imidazole and 3 volumes of buffer A. Recombinant Pif1p is eluted from the Talon column with 3 column volumes buffer A supplemented with 200 mM imidazole (see Note 10). Collect fractions from flow through, washes and elution steps in low protein-binding tubes (e.g., MiniSorp tubes) for analysis. Most of recombinant Pif1 should elute between one and two column volumes. 3. The elution profile of Pif1p should be monitored by analyzing aliquots of the colleted fractions by SDS-polyacrylamide gel electrophoresis followed by Western blotting using an anti His-tag antibody and/or Coomassie staining (An example is shown in Fig. 25.1).

Fig. 25.1. Pif1p purification. Coomassie staining of an 8% SDS-PAGE gel showing the supernatant after IPTG induction (SN), the protein pool after elution from the Talon resin (His), or the protein pool after cation exchange (CE) chromatography.

4. Pool fractions containing Pif1p and proceed to cation exchange chromatography. 3.2.2. Cation Exchange Chromatography

1. The pooled fractions from the affinity purification are poured into dialysis tubing (10 kD cut off) and dialyzed against 1 L buffer A2 at 4C overnight. Provide gentle agitation using a stirring bar. 2. Remove sample from dialysis tubing. The sample usually contains some precipitate. Pellet precipitate briefly at 4C at 10,000 g and filter the supernatant through a 0.2-mm filter fitted on a 10-mL syringe. The volume of the sample can be reduced if desired using centrifugal concentrators.

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3. The following steps assume the use of a FPLC system with gradient elution capabilities, to which is connected a 2-mL cation exchange column. The column is first rinsed with water and then equilibrated in buffer A2 until the OD280 and the conductivity signals are stable. The sample injection loop should also be rinsed with buffer A2 during this step. Load the sample on the column at a rate of 1–1.5 mL/min and then wash the column with 5 volumes of buffer A2. Start collecting 2-mL fractions and elute proteins from the column with a 0–100% gradient in buffer B2 (the gradient should be 10 column volumes in length). Using these conditions, Pif1p will elute around 300 mM NaCl. Check elution fractions by analyzing 20-mL aliquots by SDS-PAGE followed by Coomassie staining. Pool fractions containing pure Pif1p based on visual estimation of the stained gel. 4. Concentrate pooled fractions using a centrifugal concentrator with a cut of 10 kD to a volume of around 500 mL. Add 4 mL of cold buffer C and concentrate the sample again. Repeat this step three times and concentrate the sample to less than 500 mL. Transfer the concentrated protein to a clean eppendorf tube and place on ice. Add 0.8 volume of glycerol, mix well, and distribute in aliquots of 10–50 mL (or desired volume). The aliquots are then flash frozen in liquid N2 and stored at –80C (see Note 11). 3.3. Preparation of Telomerase Activity from Yeast Protein Extracts 3.3.1. Overexpression of Telomerase Core Subunits

This protocol assumes the use of a yeast strain containing a 2-mm plasmid allowing the expression of EST2 and TLC1 ORFs under the control of GAL promoters. For example, we use a pESC plasmid (stratagene) containing the EST2 and TLC1 genes placed under the control of he GAL1 and GAL10 promoters, respectively. The yeast strain used for expression was the protease deficient strain BCY123 est1 type II survivor strain (described in (5)). 1. Starting from a fresh colony on plate, grow a 50-mL culture in SCGL media lacking appropriate amino acids to saturation. Use this culture to inoculate 1 L of SCGL media (minus appropriate amino-acids) and grow at 30C with agitation to an OD600 of 1. Add 1 L of YPGLA media and incubate at 30C with agitation for another 3 h. Add 20 g galactose (2% final concentration) to induce expression of EST2 and TLC1 and incubate for another 16 h at 30C. 2. Pellet cells by centrifugation at 3000 g. Resuspend the pellet in 1 volume L buffer and pour slowly in liquid N2. Keep frozen noodles at –80C or proceed directly to telomerase fractionation.

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Since telomerase is a ribonucleoprotein, all steps of the purification should be performed at 4C and in an RNase-free environment (see Note 2). With these considerations in mind, telomerase activity can be fractionated from yeast cells using the following method, adapted from (8). 1. This protocol assumes the use of a RM100 automated mortar and pestle. Breakage of the cells is performed while cells are still frozen with liquid N2 (see Note 12). Pre-cool the mortar and pestle with liquid N2. To avoid hazardous splashes, pour only little amount of liquid N2 at a time. The mortar should be cold during the entire procedure. Add the frozen noodles obtained as described in Section 3.3 to the mortar and grind them at the finest setting of the RM100 for approximately 15 min. The cell lysate will appear as a fine white powder. Add a small amount of liquid N2 every 2–3 min to ensure that the cells stay frozen. After 15 min, check a small amount of the frozen powder under a microscope to estimate roughly lysis efficiency. Most of the cells should appear as dark debris (a lysis efficiency of about 70% or above can be easily achieved using this technique). Continue grinding if necessary. 2. Add the frozen powder to a clean 50 mL polypropylene tube and add 120 units RNase inhibitor (e.g., RNAsin, Promega) and one protease inhibitor tablet. Thaw the lysate at 4C (see Note 13). Pour the lysate in DEPC-treated corex tubes and centrifuge at 4C at 10,000 g for 30 min. Transfer supernatant in a fresh tube and place on ice. 3. Incubate the supernatant with DEAE sepharose preequilibrated in TMG-500 buffer at 4C for 30 min on a rotating wheel. Use 1 mL bed volume for 10 mL supernatant. 4. Pellet resin by centrifugation at 800 g for 1 min. Discard supernatant and replace with the same volume of TMG-500 buffer. Resuspend by gentle pipetting and gently rotate 10 min at 4C. Repeat three times. 5. Telomerase is eluted by incubation with the high salt TMG900 buffer. After the last wash, add 1 mL TMG-900 per mL of resin and incubate 15 min on the rotating wheel at 4C. Pellet resin by centrifugation at 3000 g for 10 min and save the supernatant containing telomerase at 4C. 6. The elution fraction is then desalted by passage through a PD-10 column equilibrated in TMG-30 buffer according to the manufacturer’s instructions. Concentrate the eluate using a centrifugal concentrator (10 kD cut off) to a volume of approximately 100 mL or less. Add an equal volume of glycerol. Mix well by pipetting, distribute in aliquots of 10 mL RNase-free microfuge tubes and freeze in liquid N2. Store at –80C.

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3.4. Characterization of Pif1p Activity Using Radiolabeled Oligonucleotide Substrates

3.4.1. Preparation of Oligonucleotide Substrates

Detailed methodological reviews describing radiometric assays and their application to helicase mechanistic studies exist (9–11). A favorite general reference that details synthesis of radiolabeled nucleic acids substrates and gel-based analysis of reaction products can be found in this book series (11). However, since optimal assay conditions vary among different helicases, we will briefly describe the assay system that we developed to analyze Pif1p helicase enzymatic properties using radiometric assays (5, 7). 1. Radiolabel the top (helicase-displaced)-strand oligonucleotide by adding the following to a microfuge tube: 1 mL 10 T4 polynucleotide kinase buffer, 10 pmol of oligonucleotide (5 mL, 2 mM), 3 mL [g-32P]-ATP, 1 mL T4 polynucleotide kinase. Incubate the mixture for 1 h at 37C. Inactivate the kinase by heating 5 min at 95C. Place on ice for 5 min and spin down briefly. 2. For the annealing reaction, add directly to the previous reaction: 5 mL of complementary oligonucleotide (loading strand), 2 mL 10 annealing buffer, 3 mL ddH2O. Place on a 95C heating block for 5 min, then remove the block from the thermostat and allow cooling to room temperature over a period of 2–3 h. 3. To remove unincorporated [g-32P]-ATP, load the 20 mL reaction on a prespun MicroSpin G-25 column. Centrifuge at 735 g for 1 min and collect the eluate. 4. Measure carefully the volume of the eluate and calculate the specific activity assuming 95% recovery. Add 6 Ficoll loading buffer to a final 1 concentration and load substrate on a 12% non denaturing mini-gel set up at 4C. Electrophorese at 5 V/cm until the bromophenol blue dye reaches two-thirds of the gel (or appropriate time). Disassemble and wrap gel in saran wrap. Expose briefly to an autoradiography film, develop, and cut out the gel area containing the substrate with a sterile razor blade. 5. To electroelute the substrate, place the gel piece in a dialysis midi D-Tube, fill with 1 TBE, close the tube, and place it in an electrophoresis tank containing cold 1 TBE (most horizontal agarose gel slabs will be convenient for this). Electroelute the substrate for 1 h at 80 V at 4C. 6. Invert current for 40 s, remove power supply, and disassemble. Filter the content of the dialysis tube through an Ultrafree-MC centrifugal filter and concentrate the eluate to approximately 30 mL. Measure the activity of the sample, and calculate the concentration of the radiolabeled substrate according to the previously determined specific activity. Prepare a dilution of the substrate at 10 fmol/mL and store at –20C behind a shield until use.

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1. Reaction mixtures are set up in 10 mL aliquots. Place a microfuge tube on ice and add in the following order: 2 mL of 5 helicase reaction buffer, 1 mL 50 mM Mg2+, 10 fmol radiolabeled DNA substrate (1 mL, 10 fmol/mL), 4 mL H2O, and 1 mL of Pif1p dilution in Pif1p storage buffer (see Note 14). Place the tube at 35C (or desired temperature). Start the reaction by adding 1 mL of 40 mM ATP and incubate for 15 min (or desired time) (see Note 15). 2. Stop the reaction by adding 2 mL of helicase stop buffer. 3. Load the reaction products on a 13 18 cm non-denaturing 12% polyacrylamide gel (20:1 acrylamide:bis-acrylamide ratio). Run the gel by electrophoresis at 150 V for 2 h or appropriate time at 4C (see Note 16). 4. Disassemble the gel tray and recover the gel on DEAE paper (see Note 17). It is convenient to double the DEAE paper with a piece of 3 MM paper in order to ease handling the gel. Cover with plastic wrap and expose against an autoradiography film at –80C. 5. Detection and quantification are best carried out on a dried gel using phosphorimaging systems, such as the Molecular Dynamics Storm device, and the Image quant software (GE healthcare).

3.5. Functional Interaction Assay Between Pif1p and Yeast Telomerase 3.5.1. Telomerase Assay

Yeast telomerase activity can be monitored with an oligonucleotide extension assay, using an oligonucleotide whose sequence mimicks the end of a yeast telomere. Similarly to results from other labs, we find that telomerase extends efficiently short oligonucleotides (around 15 nucleotides in size) but displays poor polymerization efficiency on longer oligonucleotides (above 30 nucleotides). It is also advantageous that the 30 end of the telomeric primer has with a unique sequence complementary to TLC1 RNA. We had the best success with the following sequence: TEL15: 50 -TGTGGTGTGTGTGGG-30 , which anneals on TLC1 at the template position 475C (8). 1. Enzymes and solutions should be kept cold during the assembly of the reaction. 2. Prepare a dilution of Pif1p in Pif1p storage buffer such that 1 mL will contain the desired amount of enzyme. 3. Assemble the telomerase reaction by adding to a microfuge tube, in the following order: 1 mL 10 TR buffer, 1 mL Telomeric primer at 1 mM, 1 mL dNTP mix, 1 mL 40 mM ATP, 2 mL [a-32P]-dTTP, and 0.3 mL RNAsin. Place the tube in a water bath at 30C for 2 min to equilibrate the reaction in temperature, and then add 1 mL Pif1p dilution and 3 mL telomerase extract. Incubate the tube for 45 min or desired time.

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4. Stop the reaction by adding 200 mL TR stop buffer containing the radiolabeled control oligonucleotide that will serve as a quantitative loading control. Prepare enough TR stop buffer containing the loading control to distribute to all samples from the same stock. Incubate for another 45 min at 30C. 5. Extract the reaction twice with phenol/chloroform. 6. Precipitate the reaction products by adding 1 volume of 4 M (NH4)2-Acetate, 30 mg glycogen, and 2.5 volumes of cold 100% Ethanol. Place the tubes at –80C for 30 min. Centrifuge at 20,000 g for 30 min at 4C. 7. Due to the presence of glycogen, a clear white pellet should be visible at the bottom of the tube after centrifugation. Remove supernatant carefully and add 300 mL of 70% ethanol. Centrifuge again, and air-dry the pellet. Add 5 mL of formamide loading buffer and heat denature the samples at 96C for 5 min. Telomerase reaction products are now ready to be separated on a sequencing gel. 3.5.2. Electrophoresis and Detection of Reaction Products

The telomerase reaction products are best resolved on a polyacrylamide-urea sequencing gel. We had the best results with 16% polyacryamide gels run on a STS45 IBI sequencing gel unit, but the method can be adapted for other equipment. 1. Before pouring the gel, plates need to be cleaned with a water soluble detergent and rinsed extensively with distilled water. Plates are then rinsed briefly with ethanol 95% and air-dried. Avoid touching the surface of the plates with hands after cleaning. Plates should then be coated with a glass plate coating solution (e.g., gel slick solution, FMC bioproducts) to facilitate removing of the gel. Distribute the coating solution evenly using a paper towel on both plates, air dry and assemble the gel unit. Prepare the 100 mL of sequencing-gel mix and pour promptly into the plates. Insert the comb (see Note 18) and let the gel polymerize for an hour. Assemble the gel unit, and pre-run the gel in 1 TBE for 1 h at 75 W constant power setting (around 1600 V on the STS45 unit). 2. Disconnect the power supply and load the samples, leaving the wells on the side of the gel empty if possible. Reconnect the power supply and run the gel until the bromophenol blue dye has reached four-fifths of the gel. 3. Disassemble the gel unit and delicately remove one plate, leaving the gel stuck to the bottom plate. Apply two sheets of 3 MM paper to the gel and carefully remove the gel from the glass plate by having it stick to the paper, starting from a corner. Cover with plastic wrap and expose at –80C against an autoradiography film. A 24-h exposure is usually sufficient. An example of the signal observed is shown in Fig. 25.2.

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Fig. 25.2. Effects of Pif1p on telomerase activity in vitro. Autoradiograph of a sequencing gel showing telomerase extension products of a TEL15 oligonucleotide in the presence of telomerase alone (lane 1), in presence of RNase A (lane 2), or two concentrations of Pif1p (lane 3, 1 mM Pif1p; lane 4, 0.2 mM Pif1p). LC: loading control.

4. If quantitation is desired, the gel can be exposed against a phosphor screen to be scanned on a phosphorImager. Sixteen percent urea-gel cannot be dried before exposure. Therefore, to expose the gel against a phosphor screen, thaw the gel at room temperature for 10 min and place it in a phosphorImager cassette. Cover with a clean plastic wrap (to prevent the phosphor screen from contact with the wet gel) and expose for 4–5 h. Longer exposure is not recommended, as the gel likely will start deteriorating. 5. Given the unique alignment between TLC1 and the TEL15 oligonucleotide, the sequence of the extension products is predicted to be 50 -TGTGGTG (8). Using the ImageQuant software, measure the intensity of each addition product as well as the intensity of the loading control. If several lanes are to be compared, the intensity of the signal given by the loading control with serve as a reference for normalization of the signal observed in each lane.

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6. Calculate the percentage of each addition product synthesized during the reaction according to the following rule, which takes into account the number of radiolabeled nucleotide for each product size: The total amount of products being synthetized (in arbitrary units) is given by the following formula: T=I1þI2þI3/ 2þI4/2þI5/2þI6/3þI7/3 where In is the quantification of band at position +n. Therefore, the percentage of product synthesized of a specific size (+n) is given by the ratio In/T, divided by the number of radiolabeled nucleotide in the product of this size (e.g., %(þ5) = I+5/T/2) (see Notes 19 and 20). 3.5.3. Telomerase Displacement Assay

1. Prepare 100 mL (or the desired volume) of M-280 Streptavidin beads by washing them successively in 5 volumes buffer S1, 5 volumes 0.1 M NaCl, and 5 volumes buffer W/B. Resuspend the beads in one volume buffer W/B. To remove buffer between each step, place tube on magnet for 1–2 min to separate beads from buffer and remove buffer by gentle pipetting. 2. Using a 100 mM stock of 50 -biotinylated TEL15 telomeric oligonucleotide (or the desired oligonucleotide), add the amount of oligonucleotide to the equilibrated beads to give a final oligonucleotide concentration of 1 mM. incubate at room temperature for 15 min with occasional mixing by gently taping the tube. Remove buffer and wash the beads 2–3 times with 1 buffer W/B and resuspend beads in 1 volume 1 TR buffer. 3. Assemble on ice the following reaction in a microfuge tube: 3 mL of beads (coated with the telomeric oligonucleotide), 4 mL of 1 TR buffer, and 3 mL of telomerase extract. Incubate at room temperature for 10 min. In the meantime, assemble in a separate tube on ice the following helicase mix: 1 mL 10 TR buffer, 1 mL ATP (40 mM stock), 1 mL Pif1p (at a concentration of 10 mM if possible), 1 mL non-biotinylated TEL15 oligonucleotide (10 mM stock), and 6 mL H2O. Separate beads from reaction mix containing unbound telomerase using a magnet and replace with the Helicase mix. Incubate the reaction at 30C for 10 min (or desired time). 4. Again, separate beads from supernatant and carefully remove supernatant by pipetting. Wash beads by gentle pipetting with 100 mL 1 TR buffer and resuspend beads in 10 mL 1 TR buffer. Add to both tubes (beads and reaction supernatant) 4 mL of 3 Laemmli buffer and boil for 5 min. Centrifuge briefly and load on an 8% SDS-PAGE protein gel.

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5. Following electrophoresis, reveal bound fraction (bead fraction) and unbound fraction (supernatant) by Western blotting using an anti-Est2p antibody (or other appropriate antibody if using, for example, a tagged version of Est2p) (see Note 21).

4. Notes 1. Other type of IMAC resins can be adapted to fit this protocol, although binding, wash, and elution buffer components (pH, salt, and imidazole concentration) should be optimized for the specific resin. 2. To prevent RNase contamination, wear latex gloves at all times and change them regularly. All glassware should be DEPC-treated. Work surfaces should be cleaned with an RNase inhibitor solution, e.g., RNaseZAP (Ambion). All buffers should be DEPC-treated and autoclaved prior to use. Tris-containing buffers can not be DEPC-treated as DEPC will react with primary amines. Therefore, RNasefree Tris and DEPC-treated stock solutions should be used to make these buffers. 3. The presence of excess unlabeled oligonucleotide prevents reannealing of the unwound strand to its complementary strand. 4. To prevent confusion between the loading control and the addition products, we use a 60-mer single-stranded oligonucleotide of random sequence as a loading control. 5. Although the pET system is convenient and has become a widely used standard, other inducible systems for heterologous expression in bacteria can be used. 6. Given the low level of Pif1p overexpression, we usually induce large volumes (5–10 L). The volume of LB in each flask should be no more than a third of the flask volume. For example, use a 6 L flask for a 2 L culture. 7. Induction at lower temperatures greatly increases Pif1p solubility. The optimal induction temperature and time will depend on the expression system and should be determined experimentally. 8. To prevent heating of the supernatant, sonication should be paused every minute for 30 s to cool the probe. One round of 100 pulses is usually enough to sonicate 100 mL of supernatant.

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9. Optimal column volume will depend on the culture volume and on the level of Pif1p expression in the system used. We find that 5-mL columns give reproducible yields and quality in Pif1p purified from 10-L cultures. 10. As a rule of thumb, 5 column volumes of washing buffer should be enough to remove proteins interacting nonspecifically with the Talon resin. If another resin is used for affinity chromatography, optimal volumes for washing the column before elution should be determined experimentally. 11. We find that the enzyme is stable at –80C for a year. It is not recommended to freeze/thaw the enzyme as it makes the helicase activity decrease rapidly. If an aliquot is thawed, it can be kept at –20C for 2–3 months without a significant drop in activity. However, we have observed precipitation at –20C. Therefore, aliquots kept at –20C should be checked for precipitation and protein concentration should be recalculated before each use. 12. Since liquid N2 can cause serious burns, safety glasses and protective gloves should be worn at all times during this procedure. 13. Frozen lysates can take a long time to thaw (> 30 min for a 20-mL lysate). Thawing can be initiated by warming the lysate between the hands and then on a rotating wheel at 4C until thawing is complete. 14. Similarly to what has been reported for other helicases, we find that the Pif1p concentration necessary to observed efficient unwinding exceeds several fold the concentration of the substrate. We routinely perform Pif1p helicase assays using 100 nM enzyme and 1 nM substrate. 15. For a control reaction, set up a reaction using Pif1p storage buffer instead of the helicase. This is the ‘‘no enzyme’’ control. Another control is the heat denatured substrate, which is achieved by heating the reaction mix containing the labeled substrate and no enzyme at 95C for 2 min. 16. The final polyacrylamide percentage of the gel and the optimal electrophoresis time depends on the sizes of the intact nucleic acids substrate and the unwound radiolabeled product. These conditions should be optimized depending on the size of the substrate used. 17. Use of DEAE paper is recommended when the gel is going to be dried before quantification, since, unlike 3 MM paper, it will bind and retain small nucleic acids. 18. Use a 5-mm well-dented comb, not a sequencing shark-tooth comb.

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19. To visualize the displacement of telomerase, a variation of this protocol can be performed. During the course of the reaction, an excess of a ‘‘chasing’’ telomeric oligonucleotide of different size is incorporated in the reaction. For example, we use a 30-mer oligonucleotide containing the TEL15 sequence extended by 15 random nucleotides from its 50 end. This oligonucleotide is utilized more efficiently than a 30-mer oligonucleotide containing only telomeric sequence, as discussed in Section 3.5.1. The reaction is started as described, but a 10-fold excess chasing oligonucleotide is added after 15 min into the reaction. Since yeast telomerase stays associated with its product (12), telomerase will only elongate the chasing oligonucleotide if telomerase is released from its elongation product. 20. The effect of Pif1p on telomerase activity can be calculated in term of telomerase nucleotide processivity, defined as the probability P with which telomerase adds more than one nucleotide without dissociating from its product. Telomerase nucleotide processivity is then defined for each product of size þn by P+n ¼ ((x > n) Ix)/T. This calculation assumes that in the presence of Pif1p, an already elongated product is not re-elongated by secondary association with telomerase, provided that TEL15 primer is present in large excess compared to telomerase. 21. Alternatively, quantification of telomerase core enzyme in each fraction can be achieved by detection of the TLC1 RNA, either by qRT-PCR or Northern blotting.

Acknowledgments This work was supported by grants from the National Insitutes of Health to VAZ.

References 1. Bessler J. B., Torres J. Z., and Zakian V. A. (2001) The Pif1p subfamily of helicases: region specific DNA helicases. Trends Cell Biol. 11, 60–65. 2. 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. 3. Schulz V. P. and Zakian V. A. (1994) The Saccharomyces PIF1 DNA helicase inhibits

telomere elongation and de novo telomere formation. Cell 76, 145–155. 4. Myung K., Chen C., and Kolodner R. D. (2001) Multiple pathways cooperate in the suppression of genome instability in Saccharomyces cerevisiae. Nature 411, 1073–1076. 5. Boule´ J.-B., Vega L. R., and Zakian V. A. (2005) The yeast Pif1p helicase removes telomerase from telomeric DNA. Nature 438, 57–61.

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6. Zhou J. Q., Monson E. M., Teng S. C., Schulz V. P., and Zakian V. A. (2000) The Pif1p helicase, a catalytic inhibitor of telomerase lengthening of yeast telomeres. Science 289, 771–774. 7. Boule´ J.-B. and Zakian V. A. (2007) The yeast Pif1p DNA helicase preferentially unwinds RNA DNA substrates. Nucleic Acids Res. 35, 5809–5818. 8. Forstemann K. and Lingner J. (2001) Molecular basis for telomere repeat divergence in budding yeast. Mol. Cell Biol. 21, 7277–7286. 9. Bachrati C. Z. and Hickson I. D. (2006) Analysis of the DNA unwinding activity of

RecQ family helicases. Methods Enzymol. 409, 86–100. 10. Brosh R. M., Jr., Opresko P. L., and Bohr V. A. (2006) Enzymatic mechanism of the WRN helicase/nuclease. Methods Enzymol. 409, 52–85. 11. Brosh R. M., Jr. and Sharma S. (2006) Biochemical assays for the characterization of DNA helicases. Methods Mol. Biol. 314, 397–415. 12. Prescott J. and Blackburn E. H. (1997) Functionally interacting telomerase RNAs in the yeast telomerase complex. Genes Dev. 11, 2790–2800.

Chapter 26 A Method to Confer Salinity Stress Tolerance to Plants by Helicase Overexpression Narendra Tuteja Abstract High salinity stress adversely affects plant growth and limits agricultural production worldwide. To minimize these losses it is essential to develop stress-tolerant plants. Several genes, including the genes encoding for helicases, are induced in response to salinity stress. Helicases are ubiquitous motor enzymes that catalyze the unwinding of energetically stable duplex DNA (DNA helicases) or duplex RNA secondary structures (RNA helicases) in an ATP-dependent manner. Helicase members of DEAD-box protein family play essential roles in cellular processes that regulate plant growth and development. Overexpression of one helicase in plant by using Agrobacterium tumefaciens-mediated transformation system confers salinity stress tolerance. To develop the salinity stress tolerant transgenic plants several sequential steps are required including cloning the helicase gene into plant transformation vector, transformation of the gene into Agrobacterium followed by Agrobacterium-mediated transformation of the gene into plant, selection and regeneration of the transgenic plants, confirmation of transgenic plants by PCR or GUS assay, and finally analysis of transgenic plants (T0 and T1 generations) for salinity stress tolerance. Key words: Agrobacterium, DEAD-box protein, DNA and RNA helicases, leaf disk assay, plant transformation, salinity stress tolerance, transgenic plant.

1. Introduction Among abiotic stress, the high salinity stress is the major cause for reducing the crop yield (1). High salinity exerts its negative impact mainly by disrupting the ionic and osmotic equilibrium of the cell (1, 2). Various genes are upregulated in response to high salinity stress signal. The products of these genes are involved, either directly or indirectly, in plant protection. Some of the genes encoding osmolytes, ion channels, receptors, components of calcium-signaling, M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_26, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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some other regulatory signaling factors, or enzymes are able to confer salinity-tolerant phenotypes when transferred to sensitive plants (2). Since abiotic stress affects the cellular gene expression machinery, it is possible that molecules involved in nucleic acid metabolism including helicases are likely to be affected (3). Helicases are motor proteins that catalyze the unwinding of duplex nucleic acids in an ATP-dependent manner and thereby play important role in most of the basic genetic processes including replication, repair, recombination, transcription, and translation (4). Most helicases share a core region (200–700 amino acids) of highly conserved nine sequence motifs (designated Q, I, Ia, Ib, II, III, IV, V, and VI). The rapidly growing DEAD-box protein family of helicases is conserved from bacteria to humans (5, 6). In the plant genome several helicase genes are present but only a few have been biochemically characterized (3, 4). Some recent reports indicate a role of helicases in salinity stress-regulated processes (7–9). To develop the salinity-tolerant plants, we have delivered PDH45 gene (pea DNA helicase 45) into the tobacco plants by using Agrobacterium tumefaciens-mediated transformation system (7, 10). Agrobacterium tumefaciens is a phytopathogenic bacterium (gram-negative) and has been widely used for plant transformation (11). The resulting transgenic plants grow normally in the presence of salt without yield loss (7).

2. Materials 2.1. Cloning the PDH45 Gene into Plant Transformation Vector

1. PDH45 cDNA: The PDH45 cDNA was cloned by pea cDNA library screening with degenerate oligodeoxynucleotide corresponding to DEAD-box motif of the helicase as described (10) (see Note 1). 2. PGMT-T Easy vector (Promega Life Science). 3. pBluescript (SK+) vector (Stratagene). 4. pBI-121 (Plant transformation vector) (Clontech, Palo Alto, CA, USA). 5. Primers to amplify ORF of PDH45 gene: Forward primer, PDH45-F1 [50 -ATGGCGACAACTTCTGTGG-30 ] starting from the translation initiation site ATG. Reverse primer, PDH45-R1 [50 -GAGCTCGAGTTATATAAGATCACCAATATTC-30 ], designed to create an XbaI site at the 30 end (italicized above) next to the translation termination codon (underlined). 6. Tobacco seeds/plants: Nicotiana tabacum cv. Xanthi (see Note 2). 7. Agrobacterium tumefaciens: Bacterial strain LBA4404.

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8. Escherichia coli (DH5): (Invitrogen life technologies) (see Note 3). 2.2. Transformation of Agrobacterium

1. YEM medium: 0.04% yeast extract, 1% mannitol, 0.01% NaCl, 0.02% MgSO4.7H2O, and 0.05% K2HPO4, pH 7.0.

2.3. AgrobacteriumMediated Plant Transformation

1. Seed surface sterilization solution: 1% bleach plus 0.1% Tween-20.

2.4. AgrobacteriumMediated Transformation

1. Benzyl amino purine (BAP).

2. MS-Basal media: 3.44 gm MS-salt (Sigma), 3% sucrose, and 1X Gamborg’s B5-vitamins (Sigma, USA), pH 5.8, and 0.6% agar.

2. Naphthan acetic acid (NAA). 3. Carbenicillin. 4. Thiamine.

2.5. Confirmation of Transgenic Plants by PCR or GUS Assay

1. CTAB extraction buffer: 2% CTAB, 1.4 M NaCl, 20 mM EDTA, pH 8.0, 100 mM Tris–HCl, pH 8.0, 100 mM bME (see Note 4) 2. Phenol mixture: phenol:chloroform:isoamyl alcohol (25:24:1). 3. Chloroform mixture: chloroform:isoamyl alcohol (24:1). 4. TE buffer: 10 mM Tris–HCl, pH 8.0, and 1 mM EDTA. 5. GUS-R1 primer: 50 -TCATTGTTTGCCTCCCTGCTGC-30 , as the reverse primer. 6. X-Gluc solution: 2 mM 5-bromo-4-chloro-3-indolyl glucuronide (X-Gluc, Biosynth. Inc.) in 50 mM of Na-phosphate buffer, pH 7.0.

3. Methods 3.1. Cloning the PDH45 Gene into Plant Transformation Vector (pBI-121)

The strategy of cloning the PDH45 gene into pBI-121 vector is described as a flow diagram in Fig. 26.1, and the detailed method is described below. 1. The complete ORF of PDH45 cDNA (1.2 kb) was PCR amplified using the gene specific forward and reverse primers. 2. The amplified fragment (1.2 kb) was purified and cloned into pGEMTeasy vector to create a pGEMT-PDH45[ORF] construct. 3. The EcoRI fragment from pGEMT-PDH45[ORF] was isolated and ligated into EcoRI site of vector pBluescript (SK+) to create a pBluescript-PDH45[ORF] construct. 4. The XbaI fragment from pBluescript-PDH45[ORF] was isolated and ligated into XbaI site of vector pBI-121 to create a pBI-121-PDH45[ORF] sense construct. This vector contains PDH45 and GUS (uidA) under a single CaMV-35S promoter;

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however, a stop codon has been inserted in between the PDH45 gene and the reporter gene to avoid translational fusions. It also carries the NPTII (Kanamycin) gene as a selectable marker. The pBI-121-PDH45[ORF] construct contains two HindIII sites, one internal in the PDH45 gene and one in the backbone of the vector just at the beginning of CaMV-35S promoter region, which will help in checking the orientation (sense or antisense) of the gene (Fig. 26.1).

Fig. 26.1. Diagramatic representation of the flow chart of generation of pBI-121-PDH45 sense construct for Agrobacteriummediated transformation of PDH45 gene into tobacco plant.

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5. The above ligated product (pBI-121-PDH45[ORF]) was transformed into Escherichia coli (DH5) competent cells and spread on LB plates containing 50 mg/ml kanamycin. Plasmid DNA was prepared from few transformed colonies and subjected to HindIII digestion followed by agarose (1%) gel electrophoresis. The product sizes of 13 kb (vector backbone) and 1.2 kb (CaMV-35S promoter + 50 end portion of PDH45 gene) fragments should give a PDH45 construct in sense orientation. 3.2. Transformation of the PDH45 Gene into Agrobacterium

1. A single colony of Agrobacterium tumefaciens was inoculated in YEM medium and kept at 28C with vigorous shaking for 2 days. 2. Competent cells of Agrobacterium were prepared by inoculating 1 ml of a full-grown culture into fresh 50 ml YEM medium and further grown at 28C till OD600 of 0.5–0.6 was reached. 3. The culture was chilled on ice and cells were harvested by centrifugation at 3,000 g for 10 min at 4C. 4. The pellet was resuspended in 2 ml of 20 mM CaCl2 (chilled) and 0.1 ml aliquots were dispensed in pre-chilled eppendorf tubes, frozen in liquid nitrogen, and stored at –80C. 5. Transformation of Agrobacterium with recombinant pBI121-PDH45[ORF] plasmid constructs in sense orientation was done by adding 0.5–1 mg of the recombinant plasmid to 0.1 ml of Agrobacterium competent cells, mixed gently, and immediately frozen in liquid nitrogen for 2 min. 6. Subsequently, cells were thawed by incubating the eppendorf tube at 37C for 5 min. 7. Thereafter, 1 ml of YEM was added to the eppendorf tube and the tube was kept for incubation at 28C for 6 h with slow shaking. 8. The revived cells were plated on YEM-agar plate containing 50 mg/ml of kanamycin and 50 mg/ml of streptomycin and incubated at 28C. 9. Transformed colonies appeared after 2–3 days and were analyzed either by PCR or by colony hybridization. 10. Agrobacterium tumefaciens carrying the recombinant pBI121 sense clones were grown in YEM containing 50 mg kanamycin and used for Agrobacterium-mediated plant transformation.

3.3. AgrobacteriumMediated Transformation of PDH45 Gene into Plant

1. Tobacco seeds were first washed with the seed surface sterilization solution for 15–20 min and then washed with 70% ethanol for 1 min followed by washing with sterile water (at least ten times).

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2. Sterilized seeds were plated on MS-Basal media and allowed to grow under moderate light, temperature, and humidity conditions till plants with healthy leaves were produced (see Note 4). 3. Wild-type tobacco plants were maintained in jam bottles by taking the stems and cutting them at the internode regions such that a single node remains intact. 4. An oblique cut was given to the internode below the node and a horizontal cut was given to the internode above the node. 5. The stem was then placed, oblique side facing downwards on approximately 50 ml MS-Basal media. 6. Healthy, unblemished leaves from young plants were harvested for preparing leaf samples. 7. Leaf discs of uniform size were made by cutting leaves into small circle or square (1 cm2)and placed them in preculture medium (MS + BAP + NAA) for 1–2 days under photo period of 16/8 h (16 h day light and 8 h dark). 8. The explants (above leaf discs) were then immersed in a 1:10 diluted grown recombinant Agrobacterium culture at OD600 = 0.5 in YEM medium (see Note 5). 9. Incubation was carried out for 5–10 min and then the incubated leaf discs were blot dried and placed on the same medium (MS + BAP + NAA) minus antibiotics with abaxial side facing upwards. 10. These explants were allowed to grow (co-cultivate) with Agrobacterium for 3–4 days. 3.4. Selection, Regeneration, and Growth of Transgenic Plants Overexpressing PDH45

1. After co-cultivation steps, the explants were briefly rinsed in MS-Basal solution containing 500 mg/ml carbenicillin, blot dried and plated onto MS-Basal-agar plates containing 300 mg/ml kanamycin, 500 mg/ml carbenicillin, 1.0 mg/ml BAP, 0.10 mg/ml NAA, and 1.0 mg/ml thiamine. 2. After 3–4 weeks, the shoots developed and defined stems were visible. They were cleanly cut from the explant and callus and placed upright on rooting medium (MS-basalagar medium containing 500 mg/ml carbenicillin and 100 mg/ml kanamycin) for root formation. Only one shoot was taken from each explant to ensure that no siblings are propagated. 3. When the roots emerged, the plants were taken out, rinsed with sterile water to remove any piece of agar, and transferred to sterile vermiculite containing pots for hardening. To provide high humidity the pots were covered with plastic bags and grown in tissue culture room (see Note 6).

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4. After 7–10 days the plastic bags were opened slowly in order to reduce the humidity gradually until the plants were acclimatized to the ambient humidity. Once the plants hardened, they were transferred to potted soil and then to glasshouse for further growth. 3.5. Confirmation of Transgenic Plants by PCR or GUS Assay

The integration of the trans-gene into the transgenic plants can be confirmed by PCR, GUS assay, Northern blot, Southern blot, or by Western blot analysis. Here only PCR and GUS assay are described.

3.5.1. PCR

PCR is one of the first approaches to confirm the integration of the PDH45 gene into the transgenic plants. For this we need gene specific primers, GUS-specific primer and the genomic DNAs from all the putative transgenic lines. Genomic DNA was isolated by grinding leaf tissue followed by extraction using the CTAB (N-acetyl-N,N,N-trimethylammonium bromide) method of Murray and Thompson (12) with minor modifications as described below. 1. Small pieces of leaf tissue (1 1 cm) were frozen in liquid nitrogen in Eppendorf tubes and homogenized in (500 ml) CTAB extraction buffer. 2. The extract was incubated at 60C for 20 min. 3. To this 500 ml of phenol mixture was added and mixed by vortexing for 30 s followed by centrifugation at 10,000 g for 5 min at room temperature. 4. The aqueous phase (top phase) was transferred to another tube, and extracted once with 500 ml of chloroform mixture in Eppendorf tube. 5. 0.6 volume of isopropanol was added to the final aqueous phase that precipitated the genomic DNA that was spooled out. 6. Genomic DNA was then washed thrice with 70% ethanol, dried in vacuum, dissolved in TE buffer containing 10 mg/ml RNase A and incubated at 37C for 1 h. 7. This was followed by extraction again with phenol mixture and aqueous phase was transferred to a fresh tube (genomic DNA). 8. Thereafter, the genomic DNA was precipitated by adding 0.3 M sodium acetate (final concentration), pH 5.2, and 2.5 volumes of ethanol. The tube was kept at –20C freezer for 1–2 h. 9. The pellet was collected by centrifugation at 10,000 g for 20 min at 4C and washed with 70% ethanol, vacuum dried and dissolved in TE. The DNA is ready for PCR as a template.

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10. PCR was performed using the PDH45-F1 and PDH45-R1 primers. The PCR was also performed using PDH45-F1 primer as the forward primer and GUS-R1, 50 TCATTGTTTGCCTCCCTGCTGC-30 , as the reverse primer. The pBI-PDH45 was used as a positive control template. 3.5.2. Histochemical GUS Assay

This method was used for screening the putative transgenic plants for the expression of b-glucuronidase (GUS). 1. Small pieces (2–3 cm) of leaf tissues from wild-type and transgenic plants were collected and rinsed in 50 mM Na-phosphate buffer, pH 7.0. 2. Then the tissue was stained with X-Gluc solution, followed by brief vacuum infiltration. 3. The stained tissues were placed at 37C overnight in dark. 4. After staining, tissues were rinsed extensively in 70% ethanol to remove chlorophyll before examination. 5. If the GUS gene is expressing then the tissues will be blue or blue-green in color, which confirmed that the tans-gene has been integrated in to the transgenic plants. 6. After visualization of the blue color in the leaf tissues, the GUS activity can also be measured fluorimetrically using 1 mM MUG as substrate as described by Jefferson (13).

3.6. Analysis of T0-Transgenic Plants to High Salinity Stress Tolerance by Leaf Disk Assay

In general, the morphological and growth characteristics of T0 generation transgenic tobacco plants were similar to the untransformed plants (wild type). The helicase overexpressing transgenic plants (T0 generation) were first checked by leaf disk assay for salinity stress tolerance as described below and the results are shown in Fig. 26.2a. 1. Healthy and fully expanded leaves (of similar age) from wild types (WT) and transgenic plants (around 60 days old) were briefly washed in deionized water (see Note 4). 2. Leaf disks of about 1 cm diameter were punched out and floated in 6 ml solution of NaCl (200 mM) or sterile distilled water (control) in petri dishes (see Note 5). 3. The petri dishes were kept at room temperature for 72 h (14). 4. After 72 h the leaf disks were visually examined for bleaching of the green color of the leaf (Fig. 26.2a) (see Note 7). 5. Please note that the WT leaf will not tolerate the salt therefore these will bleach faster and will give yellow color to the leaf disk as compared to green color (nonbleach) of the WT leaf disk floating on the water. On the other hand if the transgenic plants are salt tolerant then

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Fig. 26.2. (a) Leaf disk assay of PDH45 sense (overexpressing) transgenic lines and wildtype plant under 300 mM salt (NaCl) concentration. Relative bleaching of leaf disks from PDH45 sense and wild type are shown. (b) Growth of WT and transgenic plants in soil pot supplied with 200 mM NaCl solution. Note that WT plant could not sustain growth under salinity stress while transgenic plants grew normally up to maturity without yield loss.

the leaf disks of these plants will not bleach (remain green in color) or will bleach less (green-yellow color) as compared to the WT. 6. The chlorophyll a and b content was then measured spectrophotometrically after extraction with 80% acetone (15). 7. Overall, the results clearly showed that PDH45 overexpressing lines can tolerate the salinity stress. 3.7. Analysis of T1-Transgenic Plants Germination and Growth Under Salinity Stress 3.7.1. Analysis of T1-Transgenic Progeny

When the seeds from the T0 sense plants of PDH45 were plated onto kanamycin-containing medium, the segregation ratio was found to be in agreement with the Mendelian ratio, i.e., 3:1 (Kanr/Kans). The T1 seedlings from each line were further confirmed for the presence of the transgene by PCR and the GUS assay (as described in Sections 3.5.1 and 3.5.2). Leaf disk senescence assay for salinity stress tolerance from T1-transgenic plants and WT tobacco were also performed as described above (see Section 3.6). Overall the results were similar to the T0-transgenic plants, which clearly show that overexpressing PDH45 resulted in tolerance salinity stress. Morphologically there was not much difference in T1 generation of PDH45 tobacco transgenics and wildtype plants in terms of height, chlorophyll content, flowering and seed weight per pod.

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3.7.2. Germination of T1-Seeds and Growth of the Plants Under Salinity

The T1-transgenic lines of G and Gb and WT tobacco seeds germinated and grew normally in water. To assess the effect of high salt on seeds germination/growth of overexpressing PDH45 plants (T1) and the kanamycin-positive T1 seedlings were characterized. In the presence of salinity (100 and 200 mM NaCl) the seeds of WT plants showed no germination (or only very slow germination), while seeds of PDH45 overexpressing plants showed normal germination and the plants did not develop any sign of stress. Statistically similar results were obtained for the seven transgenic lines. Finally the transgenic plants were transferred to the pots, which continued to grow till maturity directly in the presence of salt and the plants showed normal growth under salinity stress (Fig. 26.2b).

4. Notes 1. The sequence of the degenerate oligonucleotide primer used for pea cDNA library screening is as follows: 50 -ACTAGT(A/ G/C/T)CT(AGCT)GA(T/C)GA(G/A)GC(A/G/C/ T)GA-30 . Please note that this oloigonucleotide has to be first purified electrophoretically before use as a probe. 2. The tobacco seeds first need to be surface sterilized before use (see Section 3.3). 3. The E. coli strain DH5 is the most common strain used for transformations in research laboratories. Its full genotype is as follows: F-endA1 glnV44 thi-1 relA1 gyrA96 deoR nupG lacZdeltaM15 hsdR17. This bacterial strain is resistance to nalidixic acid, therefore, E. coli strain DH5 should first be grown in LB agar plates containing 10 mg/ml nalidixic acid before use. 4. Unless stated otherwise, all solutions should be prepared in deionized water that has a resistivity of 18.2 M -cm and total organic content of less than five parts per billion. This standard is referred to as ‘‘water’’ in this text. 5. The explants should be handled very carefully using blunt ended forceps. 6. After 5–6 days few pin size holes can be made on the plastic bags. 7. The treatment was carried out in continuous white light at 25–2C.The experiment was repeated minimum three times with different transgenic lines.

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Acknowledgments Work on plant stress tolerance in NT’s laboratory is partially supported by Department of Science and Technology, Government of India, and Department of Biotechnology, Government of India. I am thankful to Dr. Renu Tuteja, Mr. Hung Quang Dung, and Dr. Hoi Xuan Pham for their help in the preparation of the article. References 1. Tuteja N. (2007) Mechanisms of high salinity tolerance in plants. Methods Enzymol. 428, 419–438. 2. Mahajan S. and Tuteja N. (2005) Cold, salinity and drought stresses: an overview. Arch. Biochem. Biophys. 444, 139–158. 3. Vashisht A. A. and Tuteja N. (2006) Stress responsive DEAD-box helicases: a new pathway to engineer plant stress tolerance. J. Photochem. Photobiol.: Biology. 84, 150–160. 4. Tuteja N. and Tuteja R. (2004) Prokaryotic and eukaryotic DNA helicases: essential molecular motor proteins for cellular machinery. Eur. J. Biochem. 271, 1835–1848. 5. Rocak S. and Linder P. (2004) DEAD-box proteins: the driving forces behind RNA metabolism. Nat. Rev. Mol. Cell Biol. 5, 232–241. 6. Tuteja N. and Tuteja R. (2004) Unraveling DNA helicases: motif, structure, mechanism and function. Eur. J. Biochem. 271, 1849–1863. 7. Sanan-Mishra N., Pham X. H., Sopory S. K., and Tuteja N. (2005) Pea DNA helicase 45 overexpression in tobacco confers high salinity tolerance without affecting yield. Proc. Natl. Acad. Sci. U.S.A. 102, 509–514. 8. Vashisht A., Pradhan A., Tuteja R., and Tuteja N. (2005) Cold and salinity stressinduced pea bipolar pea DNA helicase 47 is involved in protein synthesis and stimulated by phosphorylation with protein kinase C. Plant J. 44, 76–87.

9. Liu H. H., Liu L., Fan S. L., Song M. Z., Han X. L., Liu F., and Shen F. F. (2008) Molecular cloning and characterization of a salinity stress-induced gene encoding DEAD-box helicase from the halophyte Apocynum venetum. J. Exp. Bot. doi:10.1093/jxb/erm355 10. Pham X. H., Reddy M. K., Ehtesham N. Z., Matta B., and Tuteja N. (2000) A DNA helicase from Pisum sativum is homologous to translation initiation factor and stimulates topoisomerase I activity. Plant J. 24, 219–229. 11. Horsch R. B., Fry J. E., Hoffman N. L., Eichholtz D., Rogers S. G., and Fraley R. T. (1985) A simple and general method for transferring gene into plants. Science 227, 1229–1231. 12. Murray M. G. and Thompson W. F. (1980) Rapid isolation of high molecular-weight plant DNA. Nucleic Acid Res. 8, 4321–4325. 13. Jefferson R. A., Kavanagh T. A., and Bavan M. W. (1987) GUS fusions: betaglucuronidase as a sensitive and versatile gene fusion marker in higher plants. EMBO J. 6, 3901–3907. 14. Fan L., Zheng S., and Wang X. (1997) Antisense suppression of phospholipase D alpha retards abscisic acid—and ethylene— promoted senescence of postharvest Arabidopsis leaves. Plant Cell 9, 2183–2196. 15. Lichtenthaler H. K. (1987) Methods Enzymol. 148, 350–366.

Chapter 27 A Method to Inhibit the Growth of Plasmodium falciparum by Double-Stranded RNA-Mediated Gene Silencing of Helicases Renu Tuteja Abstract Malaria in human is caused by four Plasmodium species, with Plasmodium falciparum responsible for the most severe form of the disease. Global resistance to multiple antimalarial drugs is becoming a major challenge in worldwide efforts to control malaria. It is essential to identify new targets. One possible target is helicases, which are important ubiquitous unwinding enzymes required for nucleic acid metabolism and the maintenance of genomic stability. Helicases are motor proteins that use the energy derived from their intrinsic nucleic acid-dependent NTPase activity to unwind the duplex nucleic acid substrate. In this chapter, we study the functional role of helicases in malaria parasite by using specific dsRNA against PfH45, one of the parasite helicases. We describe the methods for Plasmodium falciparum culture, the amplification of specific helicase gene, the construction of specific dsRNA, and the analysis of the effect of dsRNA on parasite growth. Using this approach, we show that helicases are indispensable enzymes, which are required for growth and most probably survival of the malaria parasite Key words: dsRNA, DNA-dependent NTPase, helicase, malaria parasite, molecular motor, nucleic acid unwinding, Plasmodium falciparum.

1. Introduction Unwinding of double-stranded DNA into single-stranded intermediates required for various fundamental life processes is catalyzed by helicases, a family of mono-, di-, or hexameric motor proteins fueled by nucleoside triphosphate hydrolysis (1, 2). These enzymes use the free energies of binding and hydrolysis of ATP to drive the unwinding of double-stranded nucleic acids, and their function is usually ‘‘coupled’’ to the macromolecular

M.M. Abdelhaleem (ed.), Helicases, Methods in Molecular Biology 587, DOI 10.1007/978-1-60327-355-8_27, ª Humana Press, a part of Springer ScienceþBusiness Media, LLC 2010

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machines of gene expression. These enzymes are believed to transduce free energy available from NTPase activity to unwind the duplex and translocate along the nucleic acid lattice. Helicases play essential roles in many important biological processes such as DNA replication, repair, recombination, transcription, splicing, and translation. RNA helicases of the DEAD-box and related DExD/H proteins form a very large superfamily of proteins conserved from bacteria and viruses to humans and play a central role in the control of RNA metabolism (1, 2). They have seven to eight conserved motifs, the characteristics of which are used to subgroup members into individual families (3, 4). They are associated with all processes involving RNA molecules, including transcription, editing, splicing, ribosome biogenesis, RNA export, translation, RNA turnover, and organelle gene expression. Analysis of the three-dimensional structures obtained through the crystallization of viral and cellular RNA helicases reveals a strong structural homology to DNA helicases (3, 4). Malaria is one of the important and most widespread parasitic diseases caused by protozoa of the genus Plasmodium. Each year approximately 300–500 million people become infected with malaria and 2–3 million people die as a result (5). A full set of helicases was identified in the original genome sequence of Plasmodium falciparum during annotation (http://www.plasmodb.org) (6), but detailed analysis using a bioinformatics approach revealed that the genome contains at least 22 full-length putative DEAD-box helicases, as well as a few other putative helicases (7, 8). Recently it has also been reported that helicases are feasible novel drug targets for malaria (9). We have cloned and characterized important helicases from Plasmodium falciparum such as PfDH60 and PfH45 (10, 11). PfH45 contains all the conserved domains of the DEAD-box family and is homologous to eukaryotic translation-initiation factor 4A (eIF4A). It has a role in translation and it is expressed in all the intraerythrocytic developmental stages of the parasite (11). The parasite culture treated with dsRNA against PfH45 exhibited 60% growth inhibition and this inhibitory effect was due to interference with expression of the cognate messenger and downregulation of synthesis of PfH45 protein in the parasite culture (11). Our previous studies have shown that PfH45 is a multifunctional protein; therefore, inhibition of its synthesis leads to downregulation of all its associated activities, which most likely results in parasite growth inhibition. These results also suggest that PfH45 is essential for the growth and survival of the parasite (11). Few other helicases have also been characterized from Plasmodium falciparum (8, 12). The method for studying the role of helicases in parasite growth and survival using the dsRNA approach is presented in the following sections.

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2. Materials 2.1. Parasite Culture

1. Roswell Park Memorial Institute (RPMI) 1640 medium (Gibco/BRL, Bethesda, MD) supplemented with 10% human serum. 2. Human serum (type O+ve /AB+ve) is obtained by collecting fresh human blood without an anticoagulant from different (at least three) donors. The blood is stored overnight at 4C and then centrifuged at 2000 rpm for 10 min at 4C. The serum obtained is pooled and heat inactivated at 56C for 30 min in a circulating water bath. The serum is filter sterilized through a 0.45-mm filter and stored in single use aliquots at –20C. 3. Gentamycin sulfate (Sigma) is dissolved in tissue-culture water at 1 mg/ml concentration, stored in aliquots at –80C, and then added to tissue-culture dishes as required. 4. Albumax I (GIBCO): dissolve 5% (w/v) in 0.05% hypoxanthine in RPMI 1640 medium by gentle stirring for 1–2 h and filter sterilized through a 0.22-mm filter and stored in single use aliquots at –20C. 5. 5% solution of sorbitol: dissolve 5 g sorbitol (Sigma) powder in 100 ml tissue-culture water, filter sterilized through a 0.22mm filter and stored in single use aliquots at 4C. 6. 0.15% solution of saponin: dissolve 150 mg saponin (Sigma) powder in 100 ml tissue-culture water, filter sterilized through a 0.22-mm filter and stored in single use aliquots at 4C. 7. Complete medium using serum: add 5.8 ml of 3.6% sodium carbonate solution, 100 ml of gentamycin sulfate solution and 10 ml of Type O+ve /AB+ve human serum per 100 ml of RPMI 1640 solution. 8. Complete medium using Albumax is prepared by adding 5.8 ml of 3.6% sodium carbonate solution, 100 ml of Gentamycin sulfate solution and 10 ml of 5% (w/v) Albumax solution per 100 ml of RPMI 1640 solution.

2.2. Staining of Parasite Smears

1. Glass slides.

2.3. Genomic DNA Isolation

1. Phosphate buffered saline (PBS. 10X): 1.37 M NaCl, 27 mM KCl, 100 mM Na2HPO4, 18 mM KH2PO4 (adjust to pH 7.4 with HCl if necessary) and autoclave before storage at room temperature. Prepare working solution by dilution of one part with nine parts water.

2. Methanol and Giemsa stain (Sigma).

2. Lysis buffer: 50 mM Tris–HCl, pH 8.0, 5 mM EDTA, 100 mM NaCl.

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3. 10% SDS. 4. Proteinase K (20 mg/ml). 5. Phenol equilibrated with 0.1 M Tris–HCl, pH 7.0. 6. Chloroform saturated with isoamyl alcohol 24:1 ratio. 7. RNase (10 mg/ml) (New England Biolabs). 8. 3 M sodium acetate, pH 5.2. 9. Absolute and 70% alcohol. 10. TE buffer: 10 mM Tris–HCl, pH 8.0, 1 mM EDTA, pH 8.0. 2.4. RNA Isolation

1. Trizol reagent (Invitrogen). 2. PBS. 3. Isopropanol. 4. 75% ethanol. 5. RNase free water. 6. RNA isolation kit (Qiagen).

2.5. Reverse Transcription and Polymerase Chain Reaction (PCR)

1. Superscript II reverse transcriptase (Invitrogen) and reaction buffer. 2. dNTP mix (Pharmacia). 3. Oligo dT (Invitrogen), Random hexamers (Invitrogen) or gene-specific primer. 4. RNase H (New England Biolabs, Ipswich, MA, USA). 5. Taq polymerase (New England Biolabs) and Taq buffer. 6. Gene-specific sense and antisense primers. 7. Thermocycler.

2.6. Cloning of PCR Product and Preparation of dsRNA

1. pGEMT easy vector (Promega, Madison, WI, USA). 2. Competent E. coli cells. 3. In vitro transcription kit (RiboMAX express system, Promega).

3. Methods 3.1. Parasite Culture

1. Thaw the cryopreserved parasite culture (Plasmodium falciparum) at 37C in a circulating water bath by gently swirling and immediately transfer contents to a sterile 15-ml tube (13). 2. Intermittently add, drop by drop, equal volume of thawing solution (3.5% NaCl) along the sidewall of the tube and mix gently. 3. Centrifuge at 1600 rpm at room temperature for 5 min.

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4. Discard the supernatant and repeat the wash with the thawing solution until a clear supernatant is obtained. 5. Discard the supernatant and wash the packed cells once with incomplete medium. 6. Resuspend the packed cells in 5 ml of complete medium and place the plate in a gas chamber, outgas it, and incubate at 37C. 7. Replace the culture medium daily by transferring the culture plates to a sterile hood and aspirating off the medium through a sterile pipette. 8. Make smears to determine the parasitemia. Add fresh complete medium (prewarmed to 37C in a water bath) using a pipette. 9. Resuspend the settled cells in the fresh medium by gentle mixing and return the plates to gas chamber, outgas, and continue incubation at 37C. 10. At high parasitemia (5%), the parasite is subcultured. The culture is harvested by centrifuging at 1600 rpm for 5 min at room temperature. A 50% cell suspension of packed cells is made and the medium is added to obtain the desired dilution. The culture is maintained as described above. 3.2. Staining of Blood Smears

1. Parasite cultures are monitored by preparing the smears from the culture and staining them with modified Giemsa stain. The stain is diluted with water and used for staining. 2. Make the smears from the culture by spreading the concentrated cells on the glass slide. 3. Air-dry the smears and fix the cells by dipping in absolute methanol for 20 s. 4. Dilute the Giemsa stain 1:20 with de-ionized water and stain the smears by dipping in the diluted stain for 15–30 min. 5. After 30 min, remove the slides and gently wash with water. Dry the stained slides completely and view under microscope using 100 oil immersion lens. 6. Count about 1000 infected as well as total erythrocytes separately per field. Repeat the counting at least twice for a total examination of three different parts of the slide. Single or more than one parasite in an erythrocyte is counted as single infection. Calculate the percent parasitemia as number of infected erythrocytes out of 100 erythrocytes.

3.3. Synchronization of Parasite Culture by Sorbitol Treatment

1. Harvest the culture at about 10% parasitemia with majority at ring stages by centrifuging at 1500 rpm for 5 min at room temperature. 2. Add 5 volumes of 5% sorbitol solution to the cell pellet; mix gently and incubate this mixture for 5 min at 37C.

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3. Centrifuge the mixture at 1600 rpm for 5 min at room temperature and wash the cell pellet twice with complete medium. 4. Make the smear and determine the percent parasitemia by staining. Dilute the culture appropriately and continue the culture in complete medium at 37C. 5. Repeat the sorbitol treatment once again after one cycle of growth (approximately 48 h). 3.4. Isolation of Genomic DNA From the Parasite

1. For the best yield of genomic DNA, a culture with 5–10% parasitemia is used. The volume of each reagent refers to a starting culture of 10 ml. Centrifuge the culture at 1500 rpm for 5 min. Resuspend the cells in 1.5 volume of 1.5% saponin and incubate on ice for 5 min. 2. Centrifuge the suspension at 5000 rpm for 5 min at room temperature and add 9 volumes (450 ml) of the lysis buffer to the pellet. Add 10 ml of 10 mg/ml RNase stock solution and 50 ml of 10% SDS. 3. Mix well and incubate at 37C for 15 min. Add 10 ml of 20 mg/ml proteinase K solution and incubate at 37C for 60–90 min. 4. Extract with equal volume of saturated phenol and centrifuge at 13,000 rpm for 10 min. Extract the supernatant once again with phenol, followed by chloroform. 5. Precipitate the genomic DNA by adding 1/10th volume of sodium acetate and 2.5 volumes of absolute ethanol to the supernatant. Leave the mixture at –20C overnight. 6. Pellet the DNA by centrifugation at 13,000 rpm for 30 min at 4C. Wash the pellet once with 70% ethanol and air-dry. Resuspend the pellet in 25–100 ml of TE. 7. Determine the DNA concentration by measuring optical density at 260 nm and check the quality of DNA by agarose gel electrophoresis (14).

3.5. Isolation of Total RNA from the Parasite and cDNA Synthesis

1. Centrifuge the culture at 1500 rpm for 5 min and wash the cells once with PBS (see Note 1). 2. Add 1 ml of trizol reagent to the pellet and lyse the cells by passing several times through a pipette (see Note 2). Incubate the homogenized sample for 5 min at 15–30C. 3. Add 0.2 ml of chloroform per ml of trizol reagent. Mix by shaking the tube vigorously for 15 s and incubate this mixture at room temperature for 2–3 min. 4. Centrifuge at 12,000 rpm for 15 min at 2–8C. RNA remains in the upper colorless layer.

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5. Transfer the upper layer to a fresh tube and precipitate RNA by adding 0.5 ml of isopropanol per ml of trizol reagent. Incubate the samples at 15–30C for 20 min and pellet the RNA by centrifugation at 12,000 rpm for 20 min at 2–8C. 6. Remove the supernatant and wash the pellet with 75% ethanol. Recover the pellet by centrifuging at 12,000 rpm for 5 min. Briefly dry the pellet and dissolve the RNA in 100 ml of RNase free water (14). 7. For cDNA synthesis, use up to 5 mg of total RNA, 1 ml of dNTP mix, 2 ml of 10X buffer, 1 ml appropriate primer, 4 ml of 25 mM MgCl2, 2 ml of 100 mM DTT, 1 ml of RNase inhibitor and 1 ml of superscript II reverse transcriptase enzyme and make up the volume to 20 ml (see Note 3). Incubate this reaction mix at 42C for 1 h for the cDNA synthesis. 8. Terminate the reaction by incubating at 70C for 15 min and then chill on ice. 9. Centrifuge briefly and add 1 ml of RNase H to the tube and incubate at 37C before proceeding for amplification. 3.6. Amplification of the Target DNA by PCR

1. The helicase gene is amplified using Plasmodium falciparum cDNA as template and oligonucleotide primers PfHF-50 GGGATCCATGAGTACTAAAGAAGA-30 and PfHR-50 CCTCGAGTTATAAATAGTCAGCAA-30 . For the amplification of intronless genes, the Plasmodium falciparum genomic DNA can be used as template. 2. The primers PfHF and PfHR contain BamHI and XhoI sites for cloning. 3. The PCR conditions used for primer pair PfHF and PfHR are 95C for 1 min, 54C for 1 min, and 72C for 2 min. This was repeated for a total of 35 cycles and at the end one elongation was done at 72C for 10 min. 4. The PCR products are analyzed by agarose gel electrophoresis. A single band of 1.2 kb is obtained. 5. The PCR product of 1.2 kb was gel purified and cloned into pGEM-T vector (Promega) to generate PfH45 clones. The DNA was prepared and the clones were checked by using the specific restriction enzymes BamHI and XhoI. The positive clones were further confirmed by sequencing using the dideoxy sequencing reactions.

3.7. DsRNA Preparation of the Amplified Helicase Gene

1. PfH45 cloned in pGEMT easy vector is used as a template to amplify the gene using T7 and SP6 primers (see Note 4). 2. The purified template was used for in vitro transcription to generate sense RNA (sRNA) and antisense RNA(asRNA) using the T7 and SP6 RiboMAX express large-scale RNA production system from Promega.

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3. The reaction components for SP6 or T7 RNA polymerase were added in a 1.5-ml tube as follows: 5X SP6 or T7 transcription buffer, 20 ml; rNTP mix, 20 ml; DNA template 10 mg; SP6 or T7 enzyme mix 10 ml and nuclease free water to make up the volume to 100 ml. After addition of all the components, the contents are mixed gently. The mixture is incubated at 37C for 4 h. 4. The dsRNA is prepared by annealing the sRNA and asRNA as described (11). For the production of dsRNA equal amounts of sRNA and asRNA were mixed and incubated, first at 65C for 30 min and then the incubation was continued at room temperature overnight (see Note 5). The mixture was treated with DNase and precipitated after phenol–chloroform extraction. 5. The pellet of dsRNA was dissolved in diethyl pyrocarbonate (DEPC) water and treated with RNase T1. These samples were checked on 1% (w / v) native agarose gel. 6. This dsRNA was quantitated and used for the following experiments. 7. dsRNA of green fluorescent protein (GFP) used for the control experiment was also synthesized using the same procedure. The dsRNA corresponding to PfH45 and GFP are shown in Fig. 27.1 .

Fig. 27.1 dsRNA of PfH45 (lane 1) and GFP (lane 2). Lane M is the molecular weight marker. The size of the marker is written in kb on the left side.

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1. For analyzing the effect of dsRNA, the parasite culture is adjusted to 4% hematocrit with 1% infected red blood cells (RBCs) (see Note 6). 2. 200 ml of this mixture is centrifuged and the pellet is resuspended in 50 ml of incomplete media and 20 mg per ml of dsRNA. 3. This mixture is incubated at 37C with intermittent mixing to avoid settling of RBCs. 4. After this incubation, serum is added to a final concentration of 20% and the mixture is dispensed in 96-well plate and incubated at 37C for specific times. 5. The smears are made at specific time points (0, 12, 24 and 48 h) and the effect is determined by microscopic examination of parasitemia. 6. The morphology of the cultures is determined by examination of RBCs under oil immersion for the presence of intraerythrocytic Plasmodium falciparum and expressed as percentage parasitemia. An example of the parasite treated with the control GFP dsRNA and the specific dsRNA corresponding to PfH45 is shown in Fig. 27.2. It is clear that all the intraerythrocytic developmental stages are visible in the cultures treated with control GFP dsRNA (Fig. 27.2(a–c)) but the parasite morphology and growth is affected in cultures treated with PfH45-specific dsRNA (Fig. 27.2 (d)).

Fig. 27.2. The parasite morphology in untreated (a–c) or treated (d) parasite cultures. The cytologic examination of blood smears prepared from cultures revealed that the parasite morphology was distorted after treatment with PfH45-specific dsRNA, as evidenced by the presence of abnormal forms of parasites (d) as compared to the parasites prepared from cultures treated with control (GFP) dsRNA (a–c). It is clear that in cultures treated with control (GFP) dsRNA, the parasite undergoes its normal course of intraerythrocytic development and all the developmental stages, i.e., ring, trophozoite, and schizont stages are detectable (a–c). But in cultures treated with PfH45-specific dsRNA, the parasite growth is inhibited and the various developmental stages are not detected (d).

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7. The growth in cultures treated with GFP is considered as 100% and the growth in culture treated with specific dsRNA relative to the control is also determined. The inhibition rate is determined using these two values (11).

4. Notes

1. For RNA extraction, wear gloves for every step and use RNase-free plasticware. 2. Trizol seems to work better if it is prewarmed to 37C. 3. It is better to heat the RNA sample at 65C for 5 min before adding the reagents for cDNA synthesis. 4. For preparation and assay of dsRNA, RNase-free conditions and autoclaved tubes treated with DEPC should be used. 5. It is also advised to check the quality and quantity of the sRNA and asRNA before mixing for the preparation of dsRNA. 6. It is advised to set the experiment with parasite culture in triplicate to obtain statistically significant results.

Acknowledgements The author thanks Arun Pradhan for help in the preparation of figures. The work on helicases in R.T.’s laboratory is partially supported by Department of Science and Technology grant. Infrastructural support from the Department of Biotechnology, Government of India is gratefully acknowledged. References 1. Tuteja, N. and Tuteja, R. (2004) Prokaryotic and eukaryotic DNA helicases. Essential molecular motor proteins for cellular machinery. Eur. J. Biochem. 271, 1835–1848. 2. Tuteja N. and Tuteja R. (2006) Helicases as molecular motors: an insight. Physica A 372, 70–83. 3. Tanner N. K. and Linder P. (2001) DExD/ H box RNA helicases: from generic motors

to specific dissociation functions. Mol. Cell. 8, 251–262. 4. Linder P. (2006) DEAD-box proteins: a family affair—active and passive players in RNP-remodeling. Nucleic Acids Res. 34, 4168–4180. 5. Tuteja R. (2007) Malaria-An overview. FEBS J., 274, 4670–4679. 6. Gardner M. J., Hall N., Fung, E., White O., Berriman M., Hyman R. W., Carlton J. M.,

Inhibit the Growth of Plasmodium falciparum

7.

8.

9.

10.

Pain A., Nelson K. E., Bowman S. et al. (2002) Genome sequence of the human malaria parasite Plasmodium falciparum. Nature 419, 498–511. Tuteja R. and Pradhan A. (2006) Unraveling the ’DEAD-box’ helicases of Plasmodium falciparum. Gene 376, 1–12. Suntornthiticharoen P., Petmitr S., and Chavalitshewinkoon-Petmitr P. (2006) Purification and characterization of a novel 30 –50 DNA helicase from Plasmodium falciparum and its sensitivity to anthracycline antibiotics. Parasitology 133, 389–398. Tuteja R. (2007) Helicases: feasible antimalarial drug target for Plasmodium falciparum FEBS J. 274, 4699–4704. Pradhan A. and Tuteja R. (2006) Plasmodium falciparum DNA helicase 60: dsRNAand antibody-mediated inhibition of the malaria parasite growth and down regulation of its enzyme activities by DNA-

11.

12.

13.

14.

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interacting compounds. FEBS J. 273, 3545–3556. Pradhan A. and Tuteja R. (2007) Bipolar, dual Plasmodium falciparum helicase 45 expressed during intraerythrocytic developmental cycle is required for parasite growth. J. Mol. Biol. 373, 268–281. Seow F., Sato S., Janssen C. S., Riehle M. O., Mukhopadhyay A., Phillips R. S., Wilson R. J., and Barrett M. P. (2005) The plastidic DNA replication enzyme complex of Plasmodium falciparum. Mol. Biochem. Parasitol. 141, 145–153. Trager W. and Jensen J. B. (1976) Human malaria parasite in continuous culture. Science 193, 673–675. Sambrook J., Fritsch E. F., and Maniatis T. (1989) Molecular Cloning: A Laboratory Manual, 2nd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.

SUBJECT INDEX A AAA+ family ........................................................2, 4, 113, 114 fold ............................................................................ 2, 4 Affinity purification ......... 99–110, 198–199, 201–203, 213, 214–215, 365 Agrobacterium .................................. 378, 379, 380, 381–382 Antiviral agents............................................................... 224 Arabidopsis thaliana ......................................................... 195 Archae ........................................................... 4, 29, 127, 128 Assay agarose gel shift................................................. 295, 298 ATPase activity.................16–17, 22–23, 342, 347–349 binding ...37–41, 42, 169, 294–295, 297–298, 299, 328, 334, 335, 336 continuous stopped-flow fluorescence........................ 60 discontinuous gel-based radiometric .......................... 60 DNA strand annealing ..................................... 181–182 DNA unwinding...................................71, 72–74, 175, 177–179, 304 DNA unwinding and protein displacement ............... 88 dsRNA inhibition ............................................. 397–398 electrophoretic mobility shift (EMSA) ........... 306, 307, 315–316, 319–320, 328 exoribonuclease activity............................. 342, 349–351 filter binding ..................................... 294–295, 297–298 fluorescence anisotropy based DNA unwinding ...... 175 fluorometric....................................................... 211–220 helicase dissociation assay ........................................... 41 highresolution assay .................................................. 155 histochemical GUS assay.......................................... 384 leaf disk assay .................................................... 384–385 motility assay for molecular motors.......................... 160 oligonucleotide substrate-based radiometric ............ 360 optical-trapping ........................ 156, 159, 164–167, 170 RNA–protein binding filter.............. 342–343, 351–353 simultaneous helicase/protease ......................... 228, 230 strand displacement ........................ 197, 199–200, 203, 204–206 streptavidin displacement ..............87–88, 90, 138, 139, 142–143, 145–150 telomerase ................................................. 363, 369–370 translocation.............................................. 14–16, 39–40

unwinding assays........40–41, 42, 60, 62, 70, 71, 72–76, 128, 131, 133–134, 175, 177–179, 247, 283, 285–286, 288, 304 video-based tethered particle ............ 159–160, 167–168

B Bacteriophage T7.............................................................. 58 Biosensors ................................................................... 13–25 Branch migration .................................................... 128–130

C Calmodulin affinity chromatography............ 197, 198–199, 202–203 binding peptide ................................................. 197, 200 Cation exchange chromatography .......... 143, 361, 365–366 Chromatin immunoprecipitation (ChIP)...... 113–125, 266, 267, 268, 269, 274–275, 276–277 Computational model for polymerization....................................................... 68 for unwinding ............................................................. 68 Conformational change ............2, 6, 39, 58, 236, 237, 239, 240, 242 Conserved motif ................................................. 1, 2, 3, 390

D Daughter strand .................................................................. 3 Degradosome ...................................................... 7, 339–356 D-loops ........................................................... 185, 187, 197 DNA annealing...................................130–132, 176, 181, 294 binding .........15, 29, 37, 42, 86, 91, 169, 175, 178, 179, 227, 328, 331, 334, 336 polymerase......................3, 4, 30, 57, 58, 60, 61, 62, 64, 71, 76, 80, 114, 121, 269, 276, 306, 308, 321, 329, 337 replication .........................................3–4, 7, 30, 58, 390 secondary structures .................................................. 211 synthesis .....4, 61–63, 64, 71, 72, 76–77, 192, 268, 273, 394–395, 398 DNA-dependent NTPase .............................................. 390 Domain mapping............................................................ 293 Dosage compensation ..................................... 303, 304, 309 Double affinity purification .................... 198–199, 201–203

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

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402 Subject Index Double stranded break repair ............................................................... 113 DNA .........................4, 14, 74, 179, 328, 334, 336, 389 RNA.......................................................... 285, 289–398 RNA binding domain (dsRBD) ............................... 300 Drosophila................................128, 185–194, 303, 308, 323 BLM helicase.................................................... 185–194 Dual NS3 helicase/protease inhibitors ........................... 224 Duplex unwinding ............................ 87, 245–262, 340, 343

E Elastic network model ............................................ 235–243 Elementary rate constants....................................... 257, 258 Exonuclease..................................................... 4, 5, 121, 187

F Flow cell assembly........................................... 159, 164–165 Flow chambers ...................................... 34–35, 40, 167, 170 Fluorescence resonance energy transfer (FRET)............. 14, 29–43, 93, 94, 181, 182, 212 Fluorescent biosensors ................................................ 13–25 Fluorescent dyes.................................................. 50, 60, 212

G Global regression analysis................................................. 69

H Helicase Cyt-19....................................................................... 261 Dda ............................................................................. 86 DDX3 ............................................................... 281–288 Ddx4 ............................................................................. 7 Ddx5 ................................................................. 265–278 DDX9 ....................................................................... 291 DDX28 ......................................................................... 7 DEAD-box...................2, 6, 7, 245–262, 340, 378, 390 DEAH-box................................................................... 2 Ded1p ....................................................................... 254 DExD/H-box ............................................................... 2 DHX30 ......................................................................... 7 DHX32 ......................................................................... 7 DnaB...............................3, 60, 127, 128, 129, 132, 133 eIF4A ............................................................................ 6 helicase-catalyzed reaction.......................................... 58 helicase core ......................2, 3, 176, 227, 245, 246, 328 helicase motif ........2, 196, 197, 200, 203, 226, 245, 291 hepatitis C virus (HCV) non structural protein 3 (NS3) ...................86, 211–220, 223–232, 236 hexameric helicase.....................1, 3, 4, 5, 58, 60, 72–76 mHel61 ......................................................................... 7 mitochondrial................................................ 7, 339–356 monomeric ............................................................ 1, 236 Mrh4 ............................................................................. 7

Mss116p........................................................ 7, 250, 254 Mtr4.............................................................................. 7 NDH II............................................................. 291–301 NPH-II.......................................................3, 86, 95, 96 NS3 ......30, 86, 211–220, 223–232, 236, 239, 240, 241, 242, 243 p68 ................................................................ 5, 265–278 PcrA ..................................22, 30, 86, 88–89, 93, 94–97 PfDH60 .................................................................... 390 PfH45 ....................................................... 390, 395–396 Pif1p.............................................................. 7, 359–375 plant helicase............................. 196, 197–198, 200–201 RecBCD ................................................................... 168 RECQ4......................................................................... 5 RecQ helicases in Drosophila ............................ 185–194 RecQ helicases in plant..................................... 195–208 replication fork helicases................................... 127–134 RHAU .......................................................................... 7 RhlB.............................................................................. 7 Rho................................................5, 139, 142, 148, 150 ring-shaped helicase........................................ 58, 59, 60 RNA helicase .......2, 3, 5, 6, 7, 86, 87, 94–97, 246, 247, 249, 258, 259, 261, 265–278, 281–288, 291–301, 328, 339, 340, 390 RNA helicase A .................................................... 5, 291 Ski2 ....................................................................... 7, 340 Ski2p ..................................................................... 7, 340 Suv3p ............................................7, 340, 343, 347, 349 T7gp4............................................................................ 4 Twinkle ......................................................................... 7 Upf1 .............................................................. 7, 327–337 UvrABC........................................................................ 4 UvrD ................................................................. 4, 30, 85 viral helicase .............................................................. 281 WRN .................................................................... 5, 292 XPD (ERCC2)......................................................... 4, 5 High throughput screening .................................... 212, 230 Holliday junctions........................................... 174, 185, 197 Homologous recombination ........5, 86, 174, 186, 187, 188, 190, 193 Human immunodeficiency virus type 1 (HIV-1)... 281–288

I Inhibitor screening.................................................. 212, 230

K Kinetic model........................................................ 15, 46, 47 Kinetic step-size........................................45, 46, 48, 49, 62

L Ligase ................................................4, 89, 94, 95, 229, 329

HELICASES Subject Index 403 M Malaria parasite............................................................... 390 Maleless (MLE) ............................................. 291, 303–325 Mass spectrometry ............................25, 100, 101, 109, 143 Mathematical model for translocation ....................... 46–51 Mini chromosome maintenance (MCM)....... 2, 29, 38, 114 Mismatch repair (MMR) ................................................... 4 Mitochondrial degradosome complex............................................... 340 DNA repair and recombination ................................... 7 helicase .......................................................... 7, 339–356 RNA editing ................................................................. 7 Model-based methods .......................................... 63, 65, 66 Molecular motor ...................................3, 45, 156, 160, 340 Mutational analysis ......................................... 173–184, 328

Pre-steady state ensemble kinetic approaches....................................... 46 single-turnover measurements .............................. 16, 25 unwinding reactions.................................. 253–258, 259 Primase......................................................3, 4, 7, 30, 58, 80 Primer extension ......................................................... 62, 63 Processivity................2, 13, 30, 43, 48, 57, 59, 81, 258, 375 Progressive external ophthalmoplegia ................................ 7 Protein conformational change................................... 2, 237 Protein displacement by helicases............................... 85–97 Protein–protein interaction ............................ 3, 85, 99–110 PTC premature termination codon............................ 7, 327

Q Quantitative kinetic information.............................. 45, 256 Quantitative PCR, RT-PCR ......................................... 266

N Nonsense-mediated mRNA decay (NMD) ....... 7, 327, 328 Normal mode analysis............................................. 236–237 Northwestern blotting .................................... 293, 299–300 n-step .............................................................. 46, 53, 54, 59 NTP binding....................................................................... 2 NTP hydrolysis ................................................... 2, 6, 57, 85 Nucleic acid binding domain......................................... 291, 292, 300 motor..................................................................... 2, 137 substrates................................... 140–142, 145–150, 292 unwinding ................................................... 57, 105, 292 Nucleotide excision repair (NER) ...................................... 4

O Optical trap ..........................................156, 159, 160, 164–167, 168, 170 tweezers..................................................................... 165 Overexpression.............................................. 101, 143, 157, 160–162, 328–332, 360, 361, 364, 366, 373, 377–386

P PAGE-based duplex unwinding .................................... 247 P element ................................................................ 186, 188 Plant transformation............................................... 378–381 Plasmodium falciparum ............................................ 389–398 Polymerase ....................................................3, 4, 6, 30, 45, 53, 57, 58, 60, 61, 62, 64, 68, 71, 76, 77, 79, 80, 86, 94, 109, 114, 121, 139, 140, 146, 187, 192, 196, 200, 212, 229, 266, 269, 276, 282, 285, 305, 306, 308, 312, 321, 324, 329, 337, 392, 396 Polymerization kinetics............................................... 57–81 Pre-mRNA ..................................................................... 247 Pre-rRNA ........................................................................... 6

R Real-time measurement.............................................. 14, 16 RecA-like domain............................................... 3, 227, 245 Recombination.......2, 5, 7, 85, 86, 100, 101, 104, 106, 113, 114, 117, 174, 186, 187, 188, 190 Repair.........2, 4–5, 7, 85, 86, 113, 117, 155, 174, 185–194, 196, 235, 378, 390 Replication fork............3, 62, 64, 114, 118, 121, 122, 127–134, 197 initiation........................................................................ 2 Replisome.......................................................... 3, 64, 71, 76 RGG-box........................................................................ 291 Ribonucleoprotein complex .................................... 3, 6, 246 Ribosome biogenesis........................................... 6, 247, 390 RNA binding .......94, 143, 236, 238, 241, 242, 243, 246, 291, 300, 309, 327, 328, 330–331, 334, 335–336, 349 degradation ........................................... 7, 151, 339, 343 ribosomal....................................................... 6, 265, 390 secondary structure ...................................5, 6, 211, 246 SnoRNA ....................................................................... 6 SnRNA ......................................................................... 6 splicing .......................................................................... 6 RNPase ............................................................................... 3

S Saccharomyces cerevisiae.................... 100, 113–125, 128, 359 Salinity stress tolerance........................................... 377–386 Single molecule FRET.................................................................... 29–43 studies ............................................................... 155–171 Single-stranded DNA binding protein (SSB)..... 15, 16, 18, 19–21, 22, 24, 25, 62, 69, 74, 75, 91, 157, 159, 160, 162, 168 Software for model-based analysis.............................. 65–66 Spliceosome ........................................................................ 6

HELICASES

404 Subject Index Steady state reactions.............................. 254, 258, 259–261 Step size ....2, 39, 45, 46, 48, 49, 51, 53, 60, 62, 65, 68, 258 Strand annealing reactions...................... 254, 255–256, 257 Strand displacement...............62, 64, 71, 76, 197, 199–200, 203, 204–206 Streptavidin displacement and DNA unwinding by Dda ........................................................... 90–92 Superfamily .........................1, 245, 281, 291, 328, 340, 390 Syndrome Bloom syndrome................................................. 30, 185 Cockayne syndrome...................................................... 5 Rothmund-Thomson syndrome................................... 5 Werner syndrome ................................................. 5, 292

T T7 bacteriophage .......................................................... 4, 58 Tandem affinity purification (TAP)......................... 99–110 Telomere ............................................................. 5, 359, 369 telomerase activity...............7, 361–362, 366–367, 369, 371, 375 Termination ....................5, 7, 137–152, 306, 319, 327, 378 Tethered particle motion ................................................ 167 Thin layer chromatography ............151, 307, 328, 347, 350 Topoisomerase II ............................................ 305, 318, 319 Transcription coactivator ................................................................. 266 regulation .............................................. 5, 113, 265, 309 Transesterification .............................................................. 6 Transgenic plant ..................................... 378, 379, 382–386

Transient pre-steady state kinetic experiments ................ 51 Translation.....6–7, 104, 165, 207, 235, 237, 238, 247, 265, 282, 327, 359, 378, 380, 390 Translocase................2, 4, 46, 47, 48, 49, 50, 51–52, 53, 54 Translocation ...2, 13, 14–16, 17, 29, 30, 32, 37, 39–40, 42, 45–55, 58, 65, 86, 167, 247, 248, 258 directional ............................................................... 2, 45 Transposon.............................................................. 186, 188 Trichothiodystrophy (TTD) .............................................. 5

U Unwinding activity......3, 4, 6, 57, 70, 128, 129, 167, 174, 247, 250, 261, 262, 291, 292, 297, 340, 343 kinetics ............................................................ 59–60, 81

W Walker A motif........................................................... 2, 200 Walker B motif ................................................................... 2

X Xeroderma pigmentosum (XP) ...................................... 4, 5 X-linked dosage compensation............................... 303, 309

Y Yeast.........................6, 7, 86, 100, 102, 109, 114, 115–116, 117–119, 123, 128, 187, 197, 307, 329, 339–356, 359–375, 379