Protein NMR Techniques (Methods in Molecular Biology, v831)

METHODS

IN

MOLECULAR BIOLOGY™

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

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

Protein NMR Techniques Third Edition Edited by

Alexander Shekhtman and David S. Burz Department of Chemistry, University at Albany, State University of New York, Albany, NY, USA

Editors Alexander Shekhtman Department of Chemistry University at Albany State University of New York 1400 Washington Avenue Albany, NY 12222, USA [email protected]

David S. Burz Department of Chemistry University at Albany State University of New York 1400 Washington Avenue Albany, NY 12222, USA [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-61779-479-7 e-ISBN 978-1-61779-480-3 DOI 10.1007/978-1-61779-480-3 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2011943883 © Springer Science+Business Media, LLC 2012 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 Science+Business 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 Humana Press is part of Springer Science+Business Media (www.springer.com)

Preface The field of protein NMR spectroscopy has rapidly expanded into new areas of biochemistry, molecular biology, and cell biology research that were impossible to study as recently as 10 years ago. The potential to study macromolecular systems that were once considered too large or too transient or too complex by using NMR spectroscopy is now being realized with the development of innovative technologies. Standard NMR technologies are also getting a facelift in part due to the pervasive nature of high-throughput approaches in biochemical and biomedical research. These advances warrant a new edition of Protein NMR Techniques that includes an authoritative but down-to-earth description of new methodologies. This edition consists of 24 chapters divided into four major categories: NMR sample preparation, solution NMR methodologies, solid-state NMR methodologies, and data processing. The material presented contains enough detail for use not only in specialized NMR laboratories, but in biochemical, molecular, and cell biology research labs that have access to high-field NMR spectrometers. Preparing proteins for NMR spectroscopy can be a time-consuming process that may take longer than data collection and analysis combined. Expression in bacterial cells still remains one of the most popular ways of preparing NMR samples. Chapter 1 discusses new methods for optimizing and increasing the production of isotope-labeled protein in bacteria. However, some proteins are difficult to express in bacteria, in these cases an alternative approach involves using yeast cells. Chapter 2 describes a methodology for producing proteins in yeast that are usually secreted into the growth medium. This technique is proving to be as robust and economic as bacterial production. One drawback to using proteins secreted by yeast is that they may exhibit altered patterns of glycosylation and phosphorylation. To avoid this problem and achieve proper posttranslational modifications, proteins are best produced in insect or mammalian cells. Advances in the use of these cells for producing NMR samples are detailed in Chapters 3 and 4. Cell-free expression of proteins has become a method of choice for high-throughput protein production especially in cases, where the yields from in vivo overexpression are very low. Cell-free systems allow for the selective incorporation of any isotope-labeled amino acid into a target protein with minimal scrambling. Chapters 5 and 6 describe the cell-free production of proteins for solution and solidstate NMR, respectively. Finally, although well-expressed in bacterial cells, some soluble proteins do not fold properly in sufficient quantity to permit analyses of structure, dynamics, and interactions. Chapter 7 presents a methodology for expressing and purifying such proteins in a cost-efficient manner. The chapters on solution NMR methodologies range from the study of individual proteins, large multidomain proteins, protein–ligand and protein–nucleic acid complexes in vitro, to the study of proteins inside living cells. A strategy for studying supramolecular systems, which has become possible due to advances in isotope labeling and NMR pulse sequences, is described in Chapter 8. Chapter 9 presents basic protocols and the latest improvements for measuring relaxation rates and analyzing protein dynamics. Methods to help overcome difficulties in applying solution NMR to the study of membrane proteins are

v

vi

Preface

detailed in Chapter 10. Structurally characterizing multi-domain proteins can be challenging due to the inherent flexibility present in these systems and requires special approaches outlined in Chapter 11. To regulate biological activity, proteins engage in interactions with other macromolecules present in the cell. A description of methods used to prepare protein–RNA, protein–DNA, and protein–ligand complexes suitable for study by using NMR spectroscopy are presented in Chapters 12, 13, and 14. Lastly, Chapter 15 describes in-cell NMR spectroscopy, a relatively new area of NMR research that affords atomic resolution information about isotope-labeled proteins inside living cells. Solid-state NMR spectroscopy presents a complementary approach to studying proteins, especially since the method is not limited by the molecular size constraints that hamper solution NMR. With the availability of high-field NMR spectrometers, solid-state NMR has become a viable technique for acquiring unique information about protein systems that are difficult to characterize by using solution NMR. Chapter 16 reviews the use of magic angle spinning solid-state NMR to study the structure and dynamics of perdeuterated proteins. The preparation and characterization of protein complexes for solid-state NMR and methodologies to analyze the structures and dynamics of protein complexes are presented in Chapter 17. The area of membrane protein expression has seen extensive advances of late, spurred by intense interest in signaling pathways, but impeded by difficulties in preparing samples in sufficient quantity for NMR spectroscopy. Chapter 18 details methods for producing membrane proteins suitable for study by using solid-state NMR. Processing and analyzing NMR data has historically been an extremely laborious part of NMR research, requiring skillful NMR spectroscopists to assign chemical shifts and to determine atomic resolution structures of proteins. With the advent of high-throughput assignment protocols, this task has become largely manageable by a trained graduate student. Nevertheless, there are difficult cases for which there is no substitution for the experienced spectroscopist. For example, characterization of eukaryotic kinases by NMR spectroscopy is complicated by the extensive dynamics and large size exhibited by these proteins. Chapter 19 describes the procedures used to assign backbone resonances for ERK2. The reactivity of solvent-exposed backbone amides varies by a factor of at least a billion-fold because of electrostatic interactions at the protein surface. The use of electrostatic analysis of hydrogen exchange rates to analyze protein flexibility is reviewed in Chapter 20. Chapter 21 presents a strategy for assigning the backbone resonances of small- to medium-sized globular proteins in a few hours by using a highly automated program, BATCH, to acquire, process, and analyze NMR data. A versatile protocol, UNIO, that provides nearly fully automated structure determination is described in Chapter 22. In UNIO, user-intervention is encouraged and facilitated by graphical tools for preparing, analyzing, validating, and presenting the NMR structure. Chapter 23 details the use of the ARIA software, which incorporates both solution and solid-state NMR structural constraints to perform structure calculations. The final chapter, Chapter 24, introduces the software DYNAMICS for analyzing relaxation rates that characterize the overall tumbling and local dynamics of a protein. This book presents a comprehensive description of the latest innovations in the field of protein NMR. It focuses on the importance of biochemistry, molecular biology, and cell biology to NMR spectroscopy while avoiding excessive repetition of existing material, which is readily available through a number of excellent texts and reviews that cover topics

Preface

vii

relevant to studying proteins by using NMR. Rather than reiterating the fundamental principles behind NMR methodologies, we have emphasized the practical aspects of experimental design combined with practical advice and examples. We hope that this book will provide both experienced NMR spectroscopists and biochemists, who are new to the field of NMR, with enough background to successfully apply these techniques to their research. Albany, NY, USA

Alexander Shekhtman David S. Burz

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

v xi

1 A Novel Bacterial Expression Method with Optimized Parameters for Very High Yield Production of Triple-Labeled Proteins . . . . . . . . . . . . . . . . . . . 1 Victoria Murray, Yuefei Huang, Jianglei Chen, Jianjun Wang, and Qianqian Li 2 Isotopic Labeling of Heterologous Proteins in the Yeast Pichia pastoris and Kluyveromyces lactis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19 Toshihiko Sugiki, Osamu Ichikawa, Mayumi Miyazawa-Onami, Ichio Shimada, and Hideo Takahashi 3 Isotope Labeling in Insect Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 37 Krishna Saxena, Arpana Dutta, Judith Klein-Seetharaman, and Harald Schwalbe 4 Isotope Labeling in Mammalian Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55 Arpana Dutta, Krishna Saxena, Harald Schwalbe, and Judith Klein-Seetharaman 5 Cell-Free Protein Production for NMR Studies. . . . . . . . . . . . . . . . . . . . . . . . . . . . 71 Mitsuhiro Takeda and Masatsune Kainosho 6 Cell-Free Membrane Protein Expression for Solid-State NMR . . . . . . . . . . . . . . . . 85 Alaa Abdine, Kyu-Ho Park, and Dror E. Warschawski 7 Expression and Purification of Src-family Kinases for Solution NMR Studies . . . . . . 111 Andrea Piserchio, David Cowburn, and Ranajeet Ghose 8 NMR Studies of Large Protein Systems. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 133 Shiou-Ru Tzeng, Ming-Tao Pai, and Charalampos G. Kalodimos 9 Protein Dynamics by 15N Nuclear Magnetic Relaxation . . . . . . . . . . . . . . . . . . . . . . 141 Fabien Ferrage 10 Bacterial Production and Solution NMR Studies of a Viral Membrane Ion Channel . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165 Jolyon K. Claridge and Jason R. Schnell 11 Preparation of the Modular Multi-Domain Protein RPA for Study by NMR Spectroscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 181 Chris A. Brosey, Marie-Eve Chagot, and Walter J. Chazin 12 NMR Studies of Protein–RNA Interactions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 197 Carla A. Theimer, Nakesha L. Smith, and May Khanna 13 Preparation and Optimization of Protein–DNA Complexes Suitable for Detailed NMR Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 219 My D. Sam and Robert T. Clubb

ix

x

Contents

14 NMR Studies of Protein–Ligand Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 Michael Goldflam, Teresa Tarragó, Margarida Gairí, and Ernest Giralt 15 In-Cell NMR Spectroscopy in Escherichia coli . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 261 Kirsten E. Robinson, Patrick N. Reardon, and Leonard D. Spicer 16 Deuterated Peptides and Proteins: Structure and Dynamics Studies by MAS Solid-State NMR. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 279 Bernd Reif 17 Solid-State NMR Spectroscopy of Protein Complexes . . . . . . . . . . . . . . . . . . . . . . . 303 Shangjin Sun, Yun Han, Sivakumar Paramasivam, Si Yan, Amanda E. Siglin, John C. Williams, In-Ja L. Byeon, Jinwoo Ahn, Angela M. Gronenborn, and Tatyana Polenova 18 Synthesis, Purification, and Characterization of Single Helix Membrane Peptides and Proteins for NMR Spectroscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 333 Miki Itaya, Ian C. Brett, and Steven O. Smith 19 Assignment of Backbone Resonances in a Eukaryotic Protein Kinase – ERK2 as a Representative Example . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 359 Andrea Piserchio, Kevin N. Dalby, and Ranajeet Ghose 20 Electrostatics of Hydrogen Exchange for Analyzing Protein Flexibility . . . . . . . . . . 369 Griselda Hernández, Janet S. Anderson, and David M. LeMaster 21 Fast Protein Backbone NMR Resonance Assignment Using the BATCH Strategy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 407 Bernhard Brutscher and Ewen Lescop 22 Comprehensive Automation for NMR Structure Determination of Proteins . . . . . . 429 Paul Guerry and Torsten Herrmann 23 ARIA for Solution and Solid-State NMR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 453 Benjamin Bardiaux, Thérèse Malliavin, and Michael Nilges 24 Determining Protein Dynamics from 15N Relaxation Data by Using DYNAMICS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 485 David Fushman Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

513

Contributors ALAA ABDINE • CNRS and Université Paris Diderot, IBPC, Paris, France JINWOO AHN • Department of Structural Biology, Pittsburgh Center for HIV Protein Interactions, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA JANET S. ANDERSON • Department of Chemistry, Union College, NY, USA BENJAMIN BARDIAUX • NMR-supported Structural Biology, Leibnitz-Institut für Molekulare Pharmakologie (FMP), Berlin, Germany IAN C. BRETT • Department of Biochemistry and Cell Biology, Stony Brook University, Stony Brook, NY, USA CHRIS A. BROSEY • Departments of Biochemistry and Chemistry, Center for Structural Biology, Vanderbilt University, Nashville, TN, USA BERNHARD BRUTSCHER • Institut de Biologie Structurale – Jean-Pierre Ebel, CNRS, CEA, UJF, UMR5075, Grenoble Cedex, France IN-JA L. BYEON • Department of Structural Biology, Pittsburgh Center for HIV Protein Interactions, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA MARIE-EVE CHAGOT • Departments of Biochemistry and Chemistry, Center for Structural Biology, Vanderbilt University, Nashville, TN, USA WALTER J. CHAZIN • Departments of Biochemistry and Chemistry, Center for Structural Biology, Vanderbilt University, Nashville, TN, USA JIANGLEI CHEN • Department of Biochemistry and Molecular Biology, School of Medicine, Wayne State University, Detroit, MI, USA JOLYON K. CLARIDGE • Department of Biochemistry, University of Oxford, Oxford, UK ROBERT T. CLUBB • Department of Chemistry and Biochemistry, University of California, Los Angeles, CA, USA DAVID COWBURN • Departments of Biochemistry and Physiology & Biophysics, Albert Einstein College of Medicine of Yeshiva University, Bronx, NY, USA KEVIN N. DALBY • Division of Medicinal Chemistry, University of Texas, Austin, TX, USA; Graduate Programs, Cellular and Molecular Biology, Pharmacy, Biomedical Engineering and Biochemistry, University of Texas, Austin, TX, USA ARPANA DUTTA • Department of Structural Biology, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA FABIEN FERRAGE • Département de chimie, Ecole normale supérieure et Laboratoire des Biomolécules, CNRS UMR 7203, Paris, Cedex, France DAVID FUSHMAN • Department of Chemistry and Biochemistry and Center for Biomolecular Structure and Organization, University of Maryland, MD, USA MARGARIDA GAIRÍ • Servicios Cientifico Tecnicos, Universitat de Barcelona, Barcelona, Spain RANAJEET GHOSE • Department of Chemistry, The City College of New York, New York, NY, USA; Graduate Center of the City University of New York, New York, NY, USA

xi

xii

Contributors

ERNEST GIRALT • Institute for Research in Biomedicine (IRB Barcelona), Parc Científic de Barcelona, Barcelona, Spain; Departament de Química Orgànica, Universitat de Barcelona, Barcelona, Spain MICHAEL GOLDFLAM • Institute for Research in Biomedicine (IRB Barcelona), Parc Científic de Barcelona, Barcelona, Spain ANGELA M. GRONENBORN • Department of Structural Biology, Pittsburgh Center for HIV Protein Interactions, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA PAUL GUERRY • Centre Européen de RMN à très Hauts Champs, Université de Lyon, Ecole Normale Supérieure de Lyon, CNRS, Université Claude, Villeurbanne, France YUN HAN • Department of Chemistry and Biochemistry, University of Delaware, Newark, DE, USA GRISELDA HERNÁNDEZ • Department of Health and Department of Biomedical Sciences, Wadsworth Center, School of Public Health, University at Albany – SUNY, Albany, NY, USA TORSTEN HERRMANN • Centre Européen de RMN à très Hauts Champs, Université de Lyon, CNRS, Ecole Normale Supérieure de Lyon, Université Claude, Villeurbanne, France YUEFEI HUANG • Department of Biochemistry and Molecular Biology, School of Medicine, Wayne State University, Detroit, MI, USA OSAMU ICHIKAWA • Graduate School of Pharmaceutical Sciences, The University of Tokyo, Tokyo, Japan MIKI ITAYA • Department of Biochemistry and Cell Biology, Stony Brook University, Stony Brook, NY, USA MASATSUNE KAINOSHO • Graduate School of Science, Nagoya University, Nagoya, Japan; Center for Priority Areas, Tokyo Metropolitan University, Hachioji, Japan CHARALAMPOS G. KALODIMOS • Department of Chemistry and Chemical Biology, Rutgers University, Piscataway, NJ, USA MAY KHANNA • Department of Biochemistry and Molecular Biology, Indiana University School of Medicine, Indianapolis, IN, USA JUDITH KLEIN-SEETHARAMAN • Department of Structural Biology, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA DAVID M. LEMASTER • Department of Health and Department of Biomedical Sciences, Wadsworth Center, School of Public Health, University at Albany – SUNY, Albany, NY, USA EWEN LESCOP • Laboratoire de Chimie et Biologie Structurales, Institut de Chimie des Substances Naturelles, Centre de Recherche de Gif, CNRS, Gif-sur-Yvette, France QIANQIAN LI • Department of Biochemistry and Molecular Biology, School of Medicine, Wayne State University, Detroit, MI, USA THÉRÈSE MALLIAVIN • Unité de Bioinformatique Structurale, CNRS URA 2185, Institut Pasteur, Paris, France MAYUMI MIYAZAWA-ONAMI • Japan Biological Informatics Consortium (JBiC), Tokyo, Japan; Biomedicinal Information Research Center (BIRC), National Institute of Advanced Industrial Science and Technology (AIST), Tokyo, Japan

Contributors

xiii

VICTORIA MURRAY • Department of Biochemistry and Molecular Biology, School of Medicine, Wayne State University, Detroit, MI, USA MICHAEL NILGES • Unité de Bioinformatique Structurale, CNRS URA 2185, Institut Pasteur, Paris, France MING-TAO PAI • Department of Chemistry and Chemical Biology, Rutgers University, Piscataway, NJ, USA SIVAKUMAR PARAMASIVAM • Department of Chemistry and Biochemistry, University of Delaware, Newark, DE, USA KYU-HO PARK • CNRS and Université Paris Diderot, IBPC, Paris, France ANDREA PISERCHIO • Department of Chemistry, The City College of New York, New York, NY, USA TATYANA POLENOVA • Department of Chemistry and Biochemistry, University of Delaware, Newark, DE, USA PATRICK N. REARDON • Department of Biochemistry, Duke University NMR Center, Durham, NC, USA BERND REIF • Munich Center for Integrated Protein Science (CIPSM) at Department Chemie, Technische Universität München, Garching, Germany; Leibniz-Institut für Molekulare Pharmakologie (FMP), Berlin, Germany; Helmholtz-Zentrum München (HMGU), German Research Center for Environmental Health, Neuherberg, Germany KIRSTEN E. ROBINSON • Department of Biochemistry, Duke University NMR Center, Durham, NC, USA MY D. SAM • Department of Biological Chemistry and Molecular Pharmacology, Harvard Medical School, Boston, MA, USA KRISHNA SAXENA • Institute for Organic Chemistry and Chemical Biology, Center for Biomolecular Magnetic Resonance, Johann Wolfgang Goethe-University Frankfurt, Frankfurt am Main, Germany JASON R. SCHNELL • Department of Biochemistry, University of Oxford, Oxford, UK HARALD SCHWALBE • Institute for Organic Chemistry and Chemical Biology, Center for Biomolecular Magnetic Resonance, Johann Wolfgang Goethe-University Frankfurt, Frankfurt am Main, Germany ICHIO SHIMADA • Biomedicinal Information Research Center (BIRC), National Institute of Advanced Industrial Science and Technology (AIST), Tokyo, Japan; Graduate School of Pharmaceutical Sciences, The University of Tokyo, Tokyo, Japan AMANDA E. SIGLIN • Department of Molecular Medicine, Beckman Research Institute of City of Hope, Duarte, CA, USA NAKESHA L. SMITH • Department of Chemistry, University at Albany SUNY, Albany, NY, USA STEVEN O. SMITH • Department of Biochemistry and Cell Biology, Stony Brook University, Stony Brook, NY, USA LEONARD D. SPICER • Department of Biochemistry, Duke University NMR Center, Durham, NC, USA; Department of Radiology, Duke University NMR Center, Durham, NC, USA TOSHIHIKO SUGIKI • Japan Biological Informatics Consortium (JBiC), Tokyo, Japan; Biomedicinal Information Research Center (BIRC), National Institute of Advanced Industrial Science and Technology (AIST), Tokyo, Japan

xiv

Contributors

SHANGJIN SUN • Department of Chemistry and Biochemistry, University of Delaware, Newark, DE, USA HIDEO TAKAHASHI • Department of Supramolecular Biology, Graduate School of Nanobioscience, Yokohama City University, Yokohama, Japan; Biomedicinal Information Research Center (BIRC), National Institute of Advanced Industrial Science and Technology (AIST), Tokyo, Japan MITSUHIRO TAKEDA • Graduate School of Science, Nagoya University, Nagoya, Japan TERESA TARRAGÓ • Institute for Research in Biomedicine (IRB Barcelona), Parc Científic de Barcelona, Barcelona, Spain CARLA A. THEIMER • Department of Chemistry, University at Albany SUNY, Albany, NY, USA SHIOU-RU TZENG • Department of Chemistry and Chemical Biology, Rutgers University, Piscataway, NJ, USA JIANJUN WANG • Department of Biochemistry and Molecular Biology, School of Medicine, Wayne State University, Detroit, MI, USA DROR E. WARSCHAWSKI • CNRS and Université Paris Diderot, IBPC, Paris, France JOHN C. WILLIAMS • Department of Molecular Medicine, Beckman Research Institute of City of Hope, Duarte, CA, USA SI YAN • Department of Chemistry and Biochemistry, University of Delaware, Newark, DE, USA

Chapter 1 A Novel Bacterial Expression Method with Optimized Parameters for Very High Yield Production of Triple-Labeled Proteins Victoria Murray, Yuefei Huang, Jianglei Chen, Jianjun Wang, and Qianqian Li Abstract The Gram-negative bacterium Escherichia coli offer a means for rapid, high-yield, and economical production of recombinant proteins. However, when preparing protein samples for NMR, high-level production of functional isotopically labeled proteins can be quite challenging. This is especially true for the preparation of triple-labeled protein samples in D2O (2H/13C/15N). The large expense and time-consuming nature of triple-labeled protein production for NMR led us to revisit the current bacterial protein expression protocols. Our goal was to develop an efficient bacterial expression method for very high-level production of triple-labeled proteins that could be routinely utilized in every NMR lab without changing expression vectors or requiring fermentation. We developed a novel high cell-density IPTG-induction bacterial expression method that combines tightly controlled traditional IPTG-induction expression with the high cell-density of auto-induction expression. In addition, we optimize several key experimental protocols and parameters to ensure that our new high cell-density bacterial expression method routinely produces 14–25 mg of triple-labeled proteins and 15–35 mg of unlabeled proteins from 50-mL bacterial cell cultures. Key words: High yield protein production, Bacterial expression, Isotopic labeling, NMR

1. Introduction To perform NMR structural studies of proteins, we have to produce proteins that are isotopically labeled with 13C and 15N for small proteins (<20 kDa) and with 13C, 15N, and 2H for larger proteins (>20 kDa). Among the many systems available for heterologous protein production, the Gram-negative bacterium Escherichia coli remains one of the most attractive hosts (1, 2). This is especially true for isotopically labeling proteins since bacterial expression Alexander Shekhtman and David S. Burz (eds.), Protein NMR Techniques, Methods in Molecular Biology, vol. 831, DOI 10.1007/978-1-61779-480-3_1, © Springer Science+Business Media, LLC 2012

1

2

V. Murray et al.

provides the cheapest way to prepare these proteins for NMR studies (3). Protein expression and purification is a routine practice in many NMR labs, but it is not uncommon to see a drastic reduction in protein yield when isotopically labeling the proteins, especially when D2O must be used. To overcome these difficulties, we have developed a novel bacterial expression method that combines the tightly controlled traditional IPTG-induction bacterial expression with the high celldensity of the auto-induction method (4). To summarize our procedure, we first determine how to make a proper starting culture, followed by double colony selection and finally high cell-density expression. With these optimized protocols and parameters, our new bacterial expression method offers near gram quantity production of triple-labeled proteins from one-liter bacterial cell cultures, without changing expression vectors and without fermentation (5). Thus, every NMR laboratory can easily apply this novel bacterial expression method on a routine basis for the production of a very high yield of triple-labeled proteins for their NMR structural studies of proteins.

2. Materials 2.1. Sample Preparation and SDS-PAGE

1. 4× SDS loading buffer: 200 mM Tris–HCl, pH 6.8, 8% (w/v) sodium dodecyl sulfate (SDS), 0.4% (w/v) bromophenol blue, 40% glycerol. Store at room temperature. 2. Dithiothreitol (DTT): Prepare 1 M solution using sterile water. Filter through a 0.22-μm pore size membrane (syringe filter) and store 1-mL aliquots at −20°C. 3. 30% Acrylamide/bis solution (29:1). 4. 1.5 M Tris–HCl, pH 8.8. 5. 10% SDS. 6. 10% ammonium persulfate. 7. TEMED. 8. M Tris–HCl, pH 6.8. 9. Protein molecular weight markers. 10. 5× SDS running buffer: 0.5% SDS, 125 mM Tris base, 1.25 M glycine. Store at room temperature. Do not adjust pH. 11. Coomassie blue-staining solution: 0.25% (w/v) Coomassie brilliant blue, 45% methanol, 10% acetic acid. Add brilliant blue to methanol and stir for 60 min. Add water and acetic acid and stir for another 30 min. Store at room temperature (see Note 1). 12. Destaining solution: 30% methanol, 10% acetic acid. Store at room temperature.

1

2.2. Protein Expression

A Novel Bacterial Expression Method with Optimized Parameters…

3

1. LB medium (Miller): Dissolve 25 g of powdered LB medium in 1 L of distilled water or D2O. Adjust the pH to 7.4 using NaOH. Autoclave and store at room temperature (see Note 2). Add antibiotics (KAN or AMP) prior to use. 2. Kanamycin monosulfate stock solution (KAN): Dissolve KAN monosulfate to a concentration of 30 mg/mL in distilled water. Syringe filter and store 1 mL aliquots at −20°C. 3. Ampicillin sodium sulfate stock solution (AMP): Dissolve AMP sodium sulfate to a concentration of 50 mg/mL in distilled water. Syringe filter and store 1 mL aliquots at −20°C. 4. LB agar plates (Miller): Dissolve 40 g of LB agar (Miller) in 1 L of distilled water and/or D2O in a 2-L flask. Cover with foil and autoclave. Monitor the temperature as it cools. When the temperature reaches ~50°C, add 1 mL of the KAN or AMP stock solution. Pour ~10 mL into 100 × 10-mm Petri dishes and swirl to coat the plate. Let the LB agar solidify at room temperature. Place plates back into a plastic bag, seal with tape, and store at 4°C (see Note 2). 5. Isopropyl-β-D-thiogalactopyranoside (IPTG): Prepare a 1 M solution using distilled water. Syringe filter and store 1 mL aliquots at −20°C. 6. 100% Glycerol: Autoclave to sterilize. Store at room temperature. 7. 5× M9 Salts (1 L): Dissolve 64 g of Na2HPO4, 15 g of KH2PO4, 5 g of NH4Cl, and 2.5 g of NaCl in distilled water or D2O, adjust the volume to 1 L. Autoclave to sterilize and store at room temperature (see Note 2). Omit NH4Cl from this recipe when 5× M9 salts are used for isotope-labeling. Do not adjust pH. 8. 20% Glucose: Dissolve 20 g glucose in distilled water, adjust volume to 100 mL. Sterilize by filtration and store at 4°C. 9. MgSO4: Prepare a 1-M solution, autoclave, and store at room temperature. 10. CaCl2: Prepare 1-M solution, autoclave, and store at room temperature. 11. M9 minimal medium for traditional IPTG method and double colony selection (100 mL): 78 mL of distilled, sterilized water, 20 mL of 5× M9 salts, 2 mL of 20% glucose, 200 μL of 1 M MgSO4, 10 μL of 1 M CaCl2, and 100 μL of antibiotic. Add the CaCl2 last and immediately swirl the flask to dissolve the cloudy precipitate. Adjust the pH to 7.4 using NaOH. 12. M9 minimal medium for high cell-density IPTG-induction method (100 mL): 75 mL of distilled, sterilized water, 20 mL of 5× M9 salts, 5 mL of 20% glucose, 200 μL of 1 M MgSO4, 10 μL of 1 M CaCl2, and 100 μL of antibiotic. Add the CaCl2 last and immediately swirl the flask to dissolve the cloudy precipitate (see Note 3). Adjust the pH to 7.4 using NaOH (see Note 4).

4

V. Murray et al.

13. M9 for double-labeling (100 mL): 80 mL of distilled, sterilized water, 20 mL of 5× M9 salts without NH4Cl, 100 mg of 15 NH4Cl and 0.2 g of 13C-glucose (for traditional IPTG method) or 1 g of 13C-glucose (for high cell-density IPTGinduction method), 200 μL of 1 M MgSO4, 10 μL of 1 M CaCl2, and 100 μL of antibiotic. Add the CaCl2 last and immediately swirl the flask to dissolve the cloudy precipitate (see Note 3). Adjust the pH to 7.4 using NaOH (see Note 4). Use a filtration unit with a 0.22-μm pore size to sterilize the medium (see Note 5). 14. M9 for triple-labeling (100 mL): 80 mL of 99% D2O, 20 mL of 5× M9 salts without NH4Cl in D2O, 100 mg of 15NH4Cl, and 0.2 g of 13C/2H-glucose (for traditional IPTG-induction method), or 1 g of 13C/2H-glucose (for high cell-density IPTG-induction method), 200 μL of 1 M MgSO4, 10 μL of 1 M CaCl2, and 100 μL of antibiotic. You can also use 13 C-glucose; however, this usually generates ~90% deuterated triple-labeled protein samples. Add the CaCl2 last and immediately swirl the flask to dissolve the cloudy precipitate (see Note 3). Adjust the pH to 7.4 using NaOH (see Note 4). Use a filtration unit with a 0.22-μm pore size to sterilize the medium (see Note 5). 15. 1000× Trace metals (100 mL): Dissolve 811 mg of FeCl3 (50 mM), 222 mg of CaCl2 (20 mM), 125.8 mg of MnCl2 (10 mM), 161.5 mg of ZnSO4 (10 mM), 26 mg of CoCl2 (2 mM), 26.9 mg of CuCl2 (2 mM), 25.9 mg of NiCl2 (2 mM), 41.2 mg of Na2MoO4 (2 mM), 34.6 mg of Na2SeO3 (2 mM), and 12.4 mg of H3BO3 (2 mM) in 60 mM HCl. Autoclave to sterilize. Store at room temperature. 16. BME vitamins (see Note 3). 2.3. Protein Purification

1. Affinity resin: His-Bind Resin. 2. 8× Charge buffer: 400 mM NiSO4. Store at 4°C. 3. 8× Binding buffer: 160 mM Tris–HCl, pH 7.9, 2 M NaCl, 20 mM imidazole. Store in an amber bottle at room temperature. 4. 8× Wash buffer: 160 mM Tris–HCl, pH 7.9, 2 M NaCl, 240 mM imidazole. Store in an amber bottle at room temperature. 5. 4× Elute buffer: 80 mM Tris–HCl, pH 7.9, 1 M NaCl, 4 M imidazole. Store in an amber bottle at room temperature. 6. (NH4)2CO3: Dissolve 474.4 g of (NH4)2CO3 in 3.5 L of water. Once dissolved, adjust the volume to 4 L (final concentration 1.23 M) and store at room temperature. Do not adjust pH. 7. Urea.

1

A Novel Bacterial Expression Method with Optimized Parameters…

5

3. Methods When triple-labeling proteins, bacteria have to be grown in D2O, usually causing a significant reduction in protein yields. We sought to overcome this obstacle. Our strategy mainly focuses on increasing the cell density of bacterial expression without manipulation of the expression vector or use of a fermenter. Unfortunately, bacterial expression at a high cell-density in D2O usually causes several major problems, including (1) plasmid loss, (2) significant reduction in the pH of the growth medium due to cell metabolites, and (3) limited availability of dissolved oxygen. These problems often result in a low or even no protein production with high cell-density bacterial expression. We developed several practical protocols that solved these problems, including (1) preparation of a proper starting culture, (2) double colony selection in D2O, (3) optimization of bacterial expression conditions, and (4) better control of the pH of the medium. We further developed a high cell-density IPTG-induction bacterial expression method that combines the tightly controlled traditional IPTG-induction expression with high cell-density autoinduction expression. Our optimized protocols ensure plasmid stability inside bacterial cells, resulting in routine production of 14–25 mg triple-labeled proteins from a 50-mL bacterial cell culture. Importantly, this novel bacterial expression method uses the same expression vectors as the traditional IPTG-induction method and does not require a fermenter. Thus, every NMR laboratory can easily adopt this novel bacterial expression method to produce large quantities of triple-labeled proteins. 3.1. PAGE Sample Preparation

PAGE samples come either from the cell lysate isolated immediately after bacterial expression or from the column flow through during protein purification. 1. For samples collected directly from bacterial cell culture: Collect 500 μL of cells and place in a 1.7-mL microcentrifuge tube. Spin down at 12,000 × g for 5 min at room temperature using a microcentrifuge, discard the supernatant, and tap out the excess on a paper towel. 2. Add 25 μL of 4× SDS loading buffer and 25 μL of water and resuspend the pellet. Store the samples in a freezer until ready to run a gel. 3. Before running the culture samples on a gel, place the samples on a 90°C heat block for 30 min. Remove from the heat block; add 50 μL of water and vortex for 30 s (see Note 6). 4. For samples collected from flow through during protein purification, mix 60 μL of column flow through with 20 μL of

6

V. Murray et al.

4× SDS loading buffer and mix thoroughly by repeated pipetting up and down. Store the samples in a freezer until ready to run a gel. 5. Before running the flow through samples on a gel, place the samples on a 90°C heat block for 5 min (see Note 6). 6. Before loading samples on a gel, centrifuge at 12,000 × g for 10 min at room temperature using a microcentrifuge to pellet cellular debris (see Note 7). 3.2. SDS-PAGE

1. Depending on the protein size, choose an appropriate acrylamide percentage for the resolving gel (see Note 8). 2. To prepare a 10% SDS-PAGE mini-resolving gel (5 mL), using a mini-gel apparatus: Mix 1.9 mL of water, 1.7 mL of 30% acrylamide/bis solution, 1.3 mL of 1.5 M Tris–HCl, pH 8.8, 50 μL of 10% SDS, 50 μL of 10% ammonium persulfate, and 2 μL of TEMED. Mix well and pour between glass plates set in a loading cassette. Leave about 1.5-cm space on top for the stacking gel. Gently pipet water on top and let the gel set (about 20 min). 3. Pour a stacking gel (2 mL) once the resolving gel has set. Mix 1.4 mL of water, 330 μL of 30% acrylamide/bis solution, 250 μL of 1 M Tris–HCl, pH 6.8, 20 μL of 10% SDS, 20 μL of ammonium persulfate, and 2 μL of TEMED. Pour the water off the top of the resolving gel, remove excess water with filter paper, and pour the stacking gel. Insert a comb containing the appropriate number of lanes and let the stacking gel set (about 20 min). 4. Prepare 1× running buffer by diluting 100 mL of 5× running buffer solution with 400 mL of distilled water. Make sure to mix the solution well. Pour the buffer into the inner and outer chambers of the gel apparatus. For cell culture samples, load 7.5 μL of each sample into the lanes. For column flow through samples, load 20 μL into each lane. Make sure to load 5 μL of molecular weight markers in one lane. 5. Secure the lid on the gel box and plug into a power supply. Run the gel at 88 V for ~2 h. Turn off the power supply when the blue dye front reaches the bottom of the gel. 6. Remove the gel from the glass plates and place it in a small box with 20 mL of Coomassie blue-staining solution. Allow the gel to stain for at least 30 min (see Note 1). Pour out the stain, rinse the gel with water to remove excess stain and then add 30 mL of destaining solution. Place a small piece of paper towel in the box to accelerate the destaining process. This process may take a few hours; however, you can start to detect bands within 30–60 min.

1

3.3. A Proper Starting Culture

A Novel Bacterial Expression Method with Optimized Parameters…

7

A critical consideration for high-level bacterial expression is the preparation of a proper starting culture in a rich medium for scaling up in minimal medium. The general practice in NMR laboratories is to grow an overnight culture using a rich medium, such as LB, at 37°C. We observe that an overnight culture usually reaches saturation by the next morning, and may result in plasmid instability and loss due to several factors including basal leakage of the T7 expression system that expresses the toxic target proteins to the host cells under this overgrowth condition (4, 6, 7). This usually results in a poor yield of target protein. Figure 1 shows a growth curve for E. coli BL21 (DE3) cells carrying the LCAT/pET30a vector, in H2O-based LB, suggesting that the bacteria are in the exponential or log phase of growth between 6 and 7.5 h at 37°C (see Note 9). We placed particular emphasis on double colony selection (Subheading 3.5) to ensure that a high percentage of bacterial cells within this colony contain the DNA expression plasmid. 1. Perform a time course of bacterial growth of a new protein expression vector in rich (LB) medium by measuring the OD600 every 30 min for ~10–12 h in water-based rich medium and ~16–18 h in D2O-based rich medium (see Note 10). The OD600

Fig. 1. Plot of E. coli growth in 5 mL of LB medium over a 10-h period at 37°C starting with a glycerol stock. The bacterial strain, BL-21(DE3), contains a pET30a vector expressing the gene for lecithin:cholesterol acyltransferase (LCAT). Based on this plot, the log phase of the culture is between an OD600 of 1 and 3.5. Note: The growth curve is vector, protein, and bacterial strain dependent. Reproduced from Murray 2010 with permission from Cold Spring Harbor.

8

V. Murray et al.

of the log phase and the time to reach saturation are vector, protein, and bacterial strain-dependent; therefore, we suggest performing this experiment before actually expressing protein for purification and labeling (see Note 11). 2. Once the optimal OD600 of the log phase of the starting culture is determined, this will be the OD600 for all future starting cultures using both traditional IPTG expression and high celldensity expression methods. 3.4. Traditional IPTG Method

This method can be scaled up or down to suit your needs. This following protocol is used to make a 100-mL expression culture. 1. Prepare a 5-mL proper starting culture as described in Subheading 3.3 in a 50-mL conical tube with holes poked in the lid. 2. Prepare 100 mL of M9 minimal medium in a 250-mL flask. Add 1.5 mL of the starting culture and measure the OD600. (To check the OD600 at this point, aliquot 1 mL of the cell culture into a cuvette and measure the OD600.) We suggest a starting OD600 between 0.05 and 0.10 for healthy bacterial cell growth. Place the flask in a 37°C incubator with a shaking speed of 200 rpm. 3. Start to monitor the OD600 after 3 h (to check the OD600 at this point, dilute 100 μL of cell culture into a cuvette containing 900 μL of distilled water and measure the OD600. The final OD600 value is ten times the spectrophotometer reading). Once the OD600 reaches between 0.8 and 1.2 (see Note 12), remove 2 mL of cell culture and place it in a 15-mL culture tube (for a noninduced reference). Induce the remaining culture with 0.5 mM IPTG. Place both the flask and the 15-mL culture tube in a 20°C incubator overnight with a shaking speed of 200 rpm. 4. The following morning, measure the final OD600 of the cell culture. If the protein expression is induced at an OD600 of 1, the OD600 of the bacterial cell culture will be around 2–3, indicating healthy bacterial cell growth. Harvest the cells by spinning down at 10,000 × g for 10 min at 4°C using a benchtop centrifuge. Remove the supernatant and either store the cell pellet at −80°C or use immediately for protein purification. 5. To check protein expression levels, take 500 μL samples of each culture (non-induced and IPTG-induced), follow the sample preparation protocol (Subheading 3.1) and run an SDS-PAGE (Subheading 3.2) to compare non-induced and IPTG-induced samples.

3.5. Double Colony Selection

Since plasmid loss is encountered during bacterial expression in D2O much more frequently than in H2O, we describe a double colony selection procedure for triple-labeling protein in D2O. Based on our

1

A Novel Bacterial Expression Method with Optimized Parameters…

9

Fig. 2. SDS-PAGE of protein expression of apoE(1–215)/pTYB1 in D2O before (a), during (b) and after (c) double colony selection. Arrows indicate the expected protein band (~80 kDa). (a) Shows four different colonies before colony selection. (b) Shows the results of three different colonies selected from the single colony selection (lanes 1–3) and another three colonies selected from the double colony selection (lanes 4–6). The second colony selection was based on Colony 3 (lane 3, b). (c) Shows the results of six colonies from the double colony selection, indicating a high level of protein expression for all six colonies. Reproduced from Sivashanmugam 2009 with permission from Wiley Interscience.

experience, this is a critical protocol that significantly increases the yield of triple-labeled proteins. Figure 2 shows the result of a typical double colony selection of apoE(1–215)/pTYB1 in D2O, demonstrating that high protein expression levels are achieved after double colony selection. This high-level expression of apoE(1–215)/pTYB1 in D2O has been stable for more than 2 years. Usually, we perform double colony selection before we optimize the expression conditions. Thus, the traditional IPTGinduction method is the default method with which to start double colony selection. The procedure, described in Subheading 3.4, can be applied here for double colony selection, except for culture volume and the use of D2O. 1. Prepare LB agar plates with 50% D2O. 2. Perform a bacterial transformation using LB agar plates prepared with D2O. 3. Next afternoon, choose nine colonies to make starting cultures of 5 mL of LB in 50% D2O and 5 μL of antibiotic in 50-mL conical tubes, and prepare a master plate (see Note 13). Punch holes in the lids of the tubes. Eight of the nine colonies will be induced with IPTG and the ninth colony will be used as a negative, non-induced control. 4. When the starting culture is ready, add 50–100 μL to 5 mL of M9 minimal medium in 70% D2O and 5 μL of antibiotic in 50-mL conical tubes. Ensure that the starting OD600 is between 0.05 and 0.10. Place the tube in a 37°C incubator with a shaking speed of 200 rpm.

10

V. Murray et al.

5. When the OD600 of the culture reaches between 0.8 and 1 (see Note 14), add 0.5 mM IPTG to eight of the nine cultures to induce protein expression, clearly marking which culture is serving as a negative control. Place the tubes in a 20°C incubator overnight with a shaking speed of 200 rpm. 6. The following morning, remove 500 μL of cell culture from each tube and place in a microcentrifuge tube. Centrifuge at 12,000 × g for 10 min at room temperature and discard the supernatant. Prepare SDS-PAGE samples and run a gel. 7. Choose the colony expressing the biggest protein band (using the negative control as reference) and prepare 5 mL of LB culture in either 70 or 99% D2O and 5 μL of antibiotic containing the colony from the master plate. Grow at 37°C until the OD600 reaches 0.7–0.9 and spread 150 μL on an LB plate prepared in 50% D2O. Invert the plate and incubate it at 37°C overnight. 8. The following day, repeat steps 3–6 using 70 or 99% D2O for the second round of colony selection (see Note 14). When completed, choose the colony expressing the biggest protein band to make a 5-mL LB culture in 100% D2O and 5 μL of antibiotic. Grow the culture halfway through its log phase. Add 800 μL of culture to 200 μL of 100% sterile glycerol in a 2-mL cryogenic tube with a screw top cap. Pipet up and down to mix thoroughly and flash freeze the tubes by dipping them in liquid nitrogen, store at −80°C. If the protein expression level is extremely high, we suggest that you make at least 5–10 glycerol stocks of this double selected colony for future use. 3.6. High Cell-Density IPTG-Induction Method

This is a hybrid method combining traditional IPTG-induction and auto-induction bacterial expression methods. It takes advantage of tightly controlled IPTG-induction and the high cell-density of the auto-induction bacterial expression. We use rich media, such as LB and 2× YT, to reach a high cell-density before IPTGinduction and then switch the culture medium by gently spinningdown the cells and resuspending them into an equal volume of minimal medium. However, many problems may occur during a high cell-density bacterial expression that can cause a significant reduction in protein yield, such as reduced pH of the expression medium, poor aeration, and/or plasmid loss during expression. We describe the following procedure to avoid these problems, ensuring a high yield of triple-labeled protein. 1. Using the glycerol stock prepared after double colony selection (Subheading 3.5), make a starting culture in 50 mL of LB medium containing 50 μL antibiotic in a 250-mL flask (Dip the pipet tip in the glycerol stock and scratch the surface. Place the tip into the culture medium, pipet up and down a few

1

A Novel Bacterial Expression Method with Optimized Parameters…

11

times and remove the tip. Immediately return the glycerol stock to the −80°C freezer). Place the flask in a 37°C incubator with a shaking speed of 200 rpm and incubate until the OD600 is halfway through the log phase (Subheading 3.3, step 1). DO NOT let the starting culture grow overnight since the saturation of cell growth may cause plasmid loss. 2. Transfer the cells into sterile tubes and spin down the culture at 5,000 × g for 7 min at room temperature. Remove the supernatant and tap the tubes on paper towels to remove as much LB as possible. 3. Gently resuspend the pellets in 50 mL of M9 minimal medium (see Note 4) and transfer the resuspended cells to a 250-mL sterile flask. Place the flask in an incubator that is set at the optimal induction temperature (see Subheading 3.7). Maintain the shaking speed at 200 rpm for efficient aeration and keep the culture at the optimal temperature for 1–1.5 h to allow the cells to adapt to the new medium. An enhancement of approximate 0.5–1 U of OD600 should be observed at the end of this time. For example, if the OD600 is 3 right after medium exchange, you can expect the OD600 to reach 3.5–4 at the end of the 1–1.5 h incubation, indicating healthy cell growth (this increase in OD600 will be slightly less when expressing in D2O). 4. Add the optimal concentration of IPTG (Subheading 3.7) to induce protein expression and keep the culture in the incubator for an optimal period of time (Subheading 3.7) at a shaking speed of 200 rpm. 5. Measure the final OD600 before harvesting cells. You can expect to see a two- to fourfold enhancement at the end of the expression. For example, if your OD600 is 4 after the 1–1.5 h incubation, the final OD600 will be between 8 and 16, indicating healthy bacterial growth. 6. Harvest the cells by spinning down the culture at 10,000 × g for 10 min at 4°C. Remove the supernatant and either store the cell pellet at −80°C, or immediately use for protein purification. 3.7. Optimization of Various Conditions

Another important step for high-level protein production using high cell-density bacterial expression is to optimize the expression conditions such as culture temperature and the induction time. These steps are critical for the initial expression of a protein using the high cell-density expression method. We usually use the traditional IPTG-induction method first to check if a new protein can be expressed by using bacteria. Once the protein expression is confirmed by the traditional IPTG-induction method, we can optimize the high cell-density expression method to produce high-yield isotopically labeled proteins.

12

V. Murray et al.

We usually carry out time courses at different temperatures, such as 15, 20, room temperature, 30 and 37°C. We normally prepare a 5-mL starting culture either in D2O or in water for the time course and closely monitor the following parameters during expression: OD600, pH, and target protein production. The detailed procedure of optimization follows: 1. Prepare 10 mL starting cultures in 50-mL flasks: 10 mL of LB medium in 99% D2O, 10 μL of antibiotic and bacterial cells from a glycerol stock after double colony selection. Incubate at 37°C with a shaking speed of 200 rpm. 2. Once the optimal OD600 has been reached, gently centrifuge the cells at 5,000 × g for 7 min at room temperature and discard the supernatant. Resuspend the cell pellets with 10 mL of M9 minimal medium in 99% D2O in 50-mL flasks and incubate at various temperatures, such as 15, 20, room-temperature, 30° and 37°C, for 1 h. Check the OD600 of the cultures before and after this 1-h cell incubation. 3. If the OD600 of each culture after the 1-h incubation increases by 0.5–1 OD600 units, this indicates that the cells are healthy and growing after the medium exchange. Induce protein expression by adding 0.5 mM IPTG. Be sure to choose one culture without IPTG-induction to serve as the negative control. Return the tubes to appropriate shakers at different temperatures. We found that 0.5 mM IPTG usually gives a reasonable protein production, thus we always used this IPTG concentration as our starting point. However, an independent optimization of IPTG concentration can be carried out and is discussed in step 6 of this section. 4. For cultures growing below 25°C, let them grow overnight (~14–16 h). The following morning, collect 500 μL of cell culture samples every 2 h (typically collect between 16 and 28 h and one more sample the next morning). For the cultures growing above 25°C, start to collect samples every 2 h after induction for at least 8 h and one the next morning. At each time point, check the OD600 and the pH of the cell culture (see Note 4). Be sure to keep the collected cell pellets at −20°C. 5. When all of the samples have been collected, prepare samples for SDS-PAGE analysis. A comparison with the negative control (noninduced culture) allows you to determine which temperature and induction time give you the best protein yield (see Note 15). 6. Repeat the above procedure to optimize the IPTG concentration using the optimized temperature and incubation time. We usually test IPTG concentrations of 0.1, 0.25, 0.5, 0.75, and 1 mM. 7. Once the optimal conditions have been determined, you can now perform high cell-density bacterial expression on a larger

1

A Novel Bacterial Expression Method with Optimized Parameters…

13

scale. Be sure to use a large flask for better aeration. We usually use a 250-mL flask for a 50-mL cell culture and a 500-mL flask for a 100-mL cell culture. If you want to grow a 200-mL cell culture, we suggest dividing the culture into 2 × 100 mL cultures in two 500-mL flasks. Figure 3 shows a typical time course experiment performed during optimization of experimental conditions. Table 1 shows the expression parameters for the time course during our optimization of the expression of human apolipoprotein A-I (apoAI). It is clear that at the maximum OD600 the bacterial expression produces the highest yield of triple-labeled apoAI, as confirmed by a Western blot (Fig. 3).

Fig. 3. Left panel: An SDS-PAGE showing auto-induction time course of triple-labeled human apoAI expression in D2O at room temperature. Lanes 1–7 = 24, 28, 32, 36, 40, 44, and 54 h, respectively. Right panel is a Western blot of the same time course using anti-apoAI monoclonal antibody. Reproduced from Sivashanmugam 2009 with permission from Wiley Interscience.

Table 1 Parameters of the time course of human apolipoprotein A–I expression Time

24 h

28 h

32 h

36 h

40 h

44 h

54 h

OD600

2.5

3.9

7.2

9.1

8.4

8

8.1

pH

6.6

6.5

6.3

6

6

6

6.1

Protein yield

−

+

++

+++

++

++

−

14

V. Murray et al.

3.8. Protein Purification

Protein purification depends on the fusion tag that is used, as different tags serve different purposes. In our laboratory, we generally use histidine tags. In this section, we describe a typical protein purification procedure using a His-Bind Resin column. 1. Prepare 1× dilutions of all buffers. Recheck the pH to ensure that they are still 7.9. 2. Centrifuge the cell culture at 10,000 × g for 10 min at 4°C. Remove the supernatant and resuspend the pellets in 20 mL of 1× binding buffer. If the protein is in inclusion bodies and can be refolded readily during dialysis, you can resuspend the pellet in 20 mL of 1× binding buffer containing 6 M urea. 3. Lyse the cells by using either sonication or a French press. Centrifuge the lysate at 16,000 × g for 20 min at 4°C. Collect the supernatant and store it on ice. 4. Add 10 mL of 1× binding buffer and repeat step 2 at least twice, and then combine all the supernatants. Depending on the protein, you may need to add additional binding buffer and repeat step 2 3–5 times to completely extract the protein from the cells. 5. Equilibrate the affinity column with 50 mL of 1× charge buffer (see Note 16). Remove the charge buffer and equilibrate the column with 50 mL of 1× binding buffer. The column should be a light blue color after equilibration. 6. Load the column with the clear lysate from step 3. The flow rate should be ~1 mL/minute (see Note 17). Collect the flow through and remove a 60-μL sample for SDS-PAGE. 7. Wash the column with 200 mL of 1× binding buffer, followed by an additional 100 mL of 1× wash buffer. The flow rate of wash buffer should also be ~1 mL/min. Elute the column with 100 mL of 1× elution buffer. Collect the last drop of elution for SDS-PAGE to make sure that all of the protein has been eluted from the column. 8. Perform SDS-PAGE analysis with all of the collected samples to assess the purification. 9. Place the eluted protein into a dialysis bag and dialyze extensively against water containing 20 mM (NH4)2CO3 to remove imidazole, salts and possibly urea. After dialysis, freeze the protein sample with liquid nitrogen and lyophilize to obtain pure triple-labeled protein powder. Run a gel to assess the purity of the protein powder.

3.9. Conclusion

With the high cell-density IPTG-induction bacterial expression method and the practical protocols described above, we routinely produce 14–25 mg of triple-labeled proteins and 15–35 mg of unlabeled proteins from a 50-mL cell culture for all the proteins we tested. Table 2 lists the final yields of unlabeled and triplelabeled proteins obtained using high cell-density bacterial expression

1

A Novel Bacterial Expression Method with Optimized Parameters…

15

Table 2 Final yields of unlabeled and triple labeled proteins: high cell-density vs. traditional IPTG method Protein

High cell densityb (mg)

IPTGb (mg)

M.W. (Cal) (Da)

M.W. (MS) (Da)

%Dc

Triple-labeled RAP(1–210) RAP(91–323) ApoE(1–183)a Mouse apoAI(1–216) Human apoAI

20 ± 3 25 ± 3 18 ± 4 15 ± 2 14 ± 1

0.5 0.8 2 0.8 0.6

33,801 36,633 22,866 28,014 32,814

33,525 ± 195 36,376 ± 200 22,686 ± 116 27,732 ± 125 32,401 ± 150

~92 ~93 ~89 ~90 ~88

Unlabeled Human apoAI Human apoE

34 ± 1 17 ± 2

1 0.2

M.W. molecular weight Reproduced from Sivashanmugam 2009 with permission from Wiley Interscience a ApoE(1–183) was expressed in 40% D2O, the rest are expressed in 99.7% D2O b High cell density (50-mL culture volume): high cell-density expression methods, including auto-induction and high cell-density IPTG-induction; IPTG: the optimized traditional IPTG-induced expression. We repeated the expressions at least three times for all proteins, the yield shown is the average ± standard deviation c Estimated percentage of deuteration, assuming 100% 13C and 15N-labeling. For apoE(1–183), the %D is the estimated percentage of deuteration based on 40% D2O. For the other four proteins, the %D is the estimated percentage of deuteration based on 99.7% D2O

compared with the yields obtained by using the traditional IPTGinduction method in a 50-mL cell culture, the results suggest a 5–100-fold enhancement in protein yield. In addition, the protocols described produce a consistent high-level of triple-labeled protein, which is always reproducible. Table 2 also gives mass spectroscopic data for the triple-labeled protein, indicating the efficiency of deuteration for triple-labeled protein using auto-induction expressions. Overall, the deuteration efficiency is around 90% if we assume the 13C and 15N-labeling are 100%. This is because we used 99.7% D2O and nondeuterated 13C-glycerol or 13C-glucose in high celldensity expressions. This result is comparable to the deuteration efficiency of the traditional IPTG-induction expression method using single labeled 13C-glucose.

4. Notes 1. The stain can be reused multiple times. When using fresh stain, you only need to stain gels for 15–30 min. Pour the used stain into a separate container. When reusing stain, you may need to stain gels longer.

16

V. Murray et al.

2. When using D2O to replace water in LB agar, broth or M9 salts, solutions CANNOT be autoclaved to sterilize. When making LB broth, follow the directions as stated, but instead of autoclaving, use a filtration unit with a 0.22-μm pore size to sterilize. When making LB agar plates, use a microwave to bring the solution just to a boil (to dissolve agar), add antibiotics once the agar temperature cools to ~50°C, and pour into plates. When making 5× M9 salts, follow the directions as stated, however omit NH4Cl for isotopic-labeling. Once again, do not autoclave; sterilize using a filtration unit with a 0.22-μm pore size. 3. For the auto-induction method, Studier suggests using vitamins and trace metals (4). We found vitamins and trace metals help promote healthy bacterial growth when using our high cell-density IPTG-induction bacterial expression. We purchased the trace metals and BME vitamins stock solution from Sigma. The trace metals used in our laboratory is based on Studier’s recipe provided in the supplement materials of his elegant paper on the auto-induction bacterial expression method (4). For our optimized high cell-density IPTGinduction minimal medium, we added 0.25× vitamins and 0.25× trace metals (see Table 1 in ref. 5). 4. While monitoring OD600 during the optimization time course (Subheading 3.7), the pH should be monitored as well. As the cell density increases, the pH of the culture lowers due to the release of cell metabolites. If the pH becomes too low (pH < 6), it will affect bacterial cell health and protein production. If the pH drops below 6 during the time course, we increase the pH of the M9 minimal medium to 8 using NaOH to allow for a larger buffering capacity of the culture medium. 5. M9 minimal medium containing isotopes (2H, 13C, or 15N) cannot be autoclaved. It must be sterilized using a filtration unit containing a 0.22-μm pore size. 6. If the protein contains cysteine residue(s), add 10–20 mM DTT after adding water. Vortex the sample well and let it sit for 30–60 min at room temperature. 7. The release of DNA can cause the sample to become quite viscous, making it hard to load on the gel. If you notice this, you can simply sonicate your sample at a low wattage (4–6 W) for 5–10 s. Afterward, spin down the sample at 12,000 × g for 2 min at room temperature. Also, when loading the gel, remove the sample from the top portion of the supernatant to avoid the pelleted cellular debris at the bottom. 8. To determine which percentage gel to use, follow the guidelines found in Table A8-8 in Sambrook and Russell (8). Also, if you are working with proteins that weigh less than 15 kDa,

1

A Novel Bacterial Expression Method with Optimized Parameters…

17

12% Tricine gels or gradient gels are highly recommended for better resolution in the molecular weight range of 5–15 kDa. 9. Bacteria display a four-phase pattern of cell growth in liquid media. First, there is an initial lag phase when bacteria are adapting to the growth conditions; at this point, an increase in OD600 will not be seen. Second, bacteria enter their exponential or log phase at which point the bacterial cells start dividing (doubling in number). The OD600 during this log phase climbs steadily. Third, bacteria enter the stationary phase during which the rate of cell growth significantly slows due to a decrease in available nutrients and an accumulation of toxins. The OD600 will level off during this phase. Finally, if fresh medium is not made available and toxins are not removed, bacteria will enter the death phase and a noticeable drop in OD600 will be observed. The key is to utilize a starting culture during the exponential or log phase of their growth curve (see Fig. 1). 10. When performing a time course using rich media, such as LB or 2× YT in D2O, bacteria grow much more slowly than in water. 11. Based on our experience, we suggest the appropriate OD600 range for the starting culture is between 3 and 5 in LB medium and between 5 and 7 in 2× YT medium. However, the OD600 of the log phase is vector, protein, and bacterial strain-dependent. Thus, the best way to determine the middle point of the log phase is to perform a time course of bacterial growth for each new protein expression vector. 12. Based on our experience, for healthy bacterial cell growth, the culture is expected to reach an OD600 of 0.8–1.2 within 4–6 h for bacterial expressions in water and within 6–9 h for bacterial expressions in D2O if the starting OD600 is between 0.05 and 0.10. 13. When performing colony selection, it is helpful to make a “master plate.” Take a KAN or AMP plate and make a 9-square grid under the agar plate. Label boxes 1–9. When you are ready to inoculate the medium, take a sterilized tip, gently touch the selected colony, and then gently touch the agar of the master plate in the corresponding box. Go back to the original plate, retouch the same colony, and drop it into a tube. Repeat this procedure for all selected colonies. Once finished, put the lid back on the plate, invert and incubate at 37°C for about 8 h. Colonies should be about 1–2-mm in diameter. Cut a long strip of parafilm, wrap the edges of the plate, and store inverted at 4°C. You can use this plate to regrow the colonies for future cell cultures. However, the plate is only good for about 2 weeks, so be sure to also make glycerol stocks. 14. If the bacteria do not grow well in 70% D2O, then they must be trained to adapt to D2O medium. For this purpose, pick a colony off a D2O plate and start a 5-mL bacterial culture of LB

18

V. Murray et al.

medium in 25% D2O. Once the OD600 of the culture reaches 1 at 37°C, transfer 100 μL of the cell culture into 5 mL of LB medium in 50% D2O. The starting OD600 of this new culture is about 0.1. Let the cell culture grow at 37°C until the OD600 reaches 1 and transfer 100 μL of the cell culture into 5 mL of LB medium in 75% D2O and let the culture to grow at 37°C until the OD600 reaches 2–3. Use this cell culture as your starting culture. 15. A Western blot of this time course will further allow an unambiguous determination of the time point that produces the best protein yield. 16. Typically, we prepare columns containing 5 mL of affinity resin for a 50-mL cell culture since 2.5 mL of resin can bind about 20 mg of protein. This can be scaled up or down to suit your needs based on the expected protein yield. 17. If the flow rate is too slow, we have found that using a 1.5-μm syringe filter to remove cellular debris from the clear lysate prevents columns from becoming clogged and running slow. References 1. Swartz, J.R. (2001) Advances in Escherichia coli production of therapeutic proteins. Curr. Opin. Biotechnol. 12, 195–201. 2. Hewitt, L., and McDonnell, J.M. (2004) Screening and optimizing protein production in E. coli methods. Methods Mol. Biol. 278, 1–16. 3. McIntosh, L.P. and Dahlquist, F.W. (1990) Biosynthetic incorporation of 15N and 13C for assignment and interpretation of nuclear magnetic resonance spectra of proteins. Q Rev. Biophys. 23, 1–38. 4. Studier, F.W. (2005) Protein production by auto-induction in high density shaking cultures. Protein Expr. Purif. 41, 207–234. 5. Sivashanmugam, A., Murray, V., Cui, C., Zhang, Y., Wang, J., and Li, Q. (2009) Practical

protocols for production of very high yields of recombinant proteins using Escherichia coli. Protein Sci. 18, 936–948. 6. Chen, H.C., Hwang, C.F., and Mou, D.G. (1992) High-density Escherichia coli cultivation process for hyperexpression of recombinant porcine growth hormone. Enzyme Microb. Technol. 14, 321–326. 7. Baneyx, F. (1999) Recombinant protein expression in Escherichia coli. Curr. Opin. Biotechnol. 10, 411–421. 8. Sambrook, J. and Russell, D. (2001) Molecular Cloning: A Laboratory Manual (3rd ed.). Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York (ISBN 978087969577-4).

Chapter 2 Isotopic Labeling of Heterologous Proteins in the Yeast Pichia pastoris and Kluyveromyces lactis Toshihiko Sugiki, Osamu Ichikawa, Mayumi Miyazawa-Onami, Ichio Shimada, and Hideo Takahashi Abstract Several protein expression systems are available for the preparation of stable isotope-labeled recombinant proteins for NMR studies. Yeast expression systems have several advantages over prokaryotic systems, such as the widely used Escherichia coli expression system. Protein expression using the methylotrophic yeast Pichia pastoris is commonly employed for the preparation of isotope-labeled proteins. Recently, the hemiascomycete yeast Kluyveromyces lactis expression system was reported as being useful for preparing proteins for NMR studies. Since each yeast expression system has different features, their applications have increased in number. In this chapter, we describe procedures for the efficient production of uniformly isotope-labeled proteins using the P. pastoris and the K. lactis yeast expression systems. Key words: Yeast expression systems, Pichia pastoris, Kluyveromyces lactis, Stable isotope labeling, Fed-batch fermentation, NMR

1. Introduction The most widely used method for the expression of isotopically labeled heterologous recombinant proteins is the Escherichia coli expression system because of easy handling, rapid and high-density cell growth, high levels of protein production, and relatively low costs for isotope-labeling. However, in many cases, expression of structurally and functionally intact eukaryotic proteins by E. coli and other prokaryotic cells is fundamentally difficult due to (1) the lack of intracellular organelles, (2) a limited number of molecular chaperones, and (3) the absence of posttranslational modification mechanisms (1–6). Yeast, however, combines several advantages of

Alexander Shekhtman and David S. Burz (eds.), Protein NMR Techniques, Methods in Molecular Biology, vol. 831, DOI 10.1007/978-1-61779-480-3_2, © Springer Science+Business Media, LLC 2012

19

20

T. Sugiki et al.

both eukaryotic and prokaryotic expression systems (1) processing, folding, complex disulfide-bond network formation, and posttranslational modification (e.g., glycosylation) of proteins are possible, (2) yeast exhibit rapid growth rates and grow to high-density, and (3) methods for molecular genetic manipulation are well established and are simple to perform (1–6). Here, we introduce the Pichia pastoris and Kluyveromyces lactis yeast expression systems for the stable isotopic labeling of heterologous proteins. 1.1. The P. pastoris Expression System

Host cells and expression vectors are available from Invitrogen. Based upon Invitrogen instruction manuals (7), previous reports (8–15), and our experience, we describe here optimized cell culture procedures for the production of isotope-labeled heterologous proteins by P. pastoris. P. pastoris is capable of utilizing methanol as both a carbon source and to induce protein expression. Methanol is oxidized by alcohol oxidases, AOX1 and AOX2, in yeast cell peroxisomes (though the majority of alcohol oxidase activity is attributed to AOX1). The AOX1 promoter tightly regulates expression of the AOX1 gene, and methanol induces AOX1 promoter activity. Thus, the AOX1 promoter is used to drive the expression of a target protein by replacing the AOX1 gene with a cDNA encoding the desired heterologous protein (1–6). Invitrogen supplies several vector series that are commonly used for protein expression in P. pastoris, including pPIC9K, pPIC3.5K, and pPICZ. Using these vectors, a cDNA cassette containing the target gene under the control of the AOX1 promoter is inserted into the genome of P. pastoris by using homologous recombination along with a gene coding for resistance to a drug, such as Zeocin™ or G418, for subsequent selection of transformed cells (7). Target proteins are expressed either intracellularly or secreted into the medium by P. pastoris. For secretion of target proteins, vectors pPICZα and pPIC9K are recommended (7). The pPICZα and pPIC9K series contain a gene that encodes a prepro signal sequence [such as the Saccharomyces cerevisiae α-mating factor (α-MF)] between the AOX1 promoter and the target gene (7). Although target proteins may contain native secretion signals, in many cases, the α-MF sequence is generally used as the sole secretion signal. Since P. pastoris secretes a low amount of its native proteins, a secreted target protein comprises the vast majority of the total proteins in the culture medium, simplifying purification of the target protein (1–7). A different set of vectors lacking secretion signal sequences are required if the target protein is to be expressed intracellularly. For cytosolic and nonglycosylated proteins, vectors pPICZ or pPIC3.5K are recommended (7). In this chapter, we describe methods and provide technical advice for producing uniformly labeled [U-13C, 15N] and [U-2H, 15N] target proteins in P. pastoris (8–14). The procedures described are

2

Isotopic Labeling of Heterologous Proteins in the Yeast…

21

optimized for secretory expression of isotope-labeled DDR2 from P. pastoris strain X-33 (Mut+ phenotype transformants) using the pPICZα vector system (15). 1.2. The K. lactis Expression System

Host cells and expression vectors are available from New England Biolabs. Based upon instruction manuals (16), previous reports, and our experience, we describe here optimized cell culture procedures for the production of isotope-labeled heterologous proteins by K. lactis. Heterologous recombinant proteins expressed by K. lactis can be sequestered intracellularly or secreted into the medium. As with P. pastoris, K. lactis secretes very low levels of native proteins, therefore secreted target proteins comprise the vast majority of the total protein present in the culture medium, which simplifies purification of the target protein (16–18). In K. lactis, the LAC4 promoter drives expression of the target protein gene. Using the pKLAC vector, a cDNA cassette containing the target gene under the control of the LAC4 promoter is inserted into the genome of K. lactis by using homologous recombination (16–18). The transcriptional activity of the LAC4 promoter is induced by galactose, which K. lactis also utilizes as a carbon source for cell growth. Therefore, expression of the target protein is constitutively induced by cultivation in a medium containing galactose (16–18). The pKLAC vectors impart a fungal acetamidase gene (amdS) to the transformants. Since only transformants that express amdS can utilize acetamide as a sole nitrogen source, amdS is used for selection of the desired transformants, thereby obviating the need for expensive antibiotics, such as Zeocin™ and rendering this auxotrophic selection method more cost-effective (16–18). In addition, auxotrophic acetamide selection enriches populations of transformants in which multiple tandem copies of target cDNA are integrated into the yeast genome (16–18). Thus, the K. lactis protein expression system is simple, cost-effective, easily scaled-up, and highly reproducible. We recently established a cost-effective isotope-labeling method utilizing the hemiascomycete yeast K. lactis (6, 19). In the most commonly employed K. lactis expression system, 20 g/L galactose is required as a carbon source for cell growth and as an inducer of target protein expression (16). However, while a 20 g/L carbon source is acceptable for uniform 15N-labeling, it is economically infeasible for uniform 13C-labeling. Sugiki et al. reported that by combining K. lactis strain GG799, which is characterized by weak glucose suppression for the LAC4 promoter (16), with a fed-batch culture method, larger amounts of protein can be expressed using a smaller amount of glucose, thus reducing costs to a level comparable to isotope-labeling by E. coli systems (6, 19). In this chapter, we describe methods and provide technical advice for producing uniformly labeled [U-13C, 15N] and [U-2H, 15N] target proteins in K. lactis.

22

T. Sugiki et al.

2. Materials The recipes for the culture media described here were derived mainly from the manuals for the EasySelect™ Pichia Expression Kit (Invitrogen), the Pichia Fermentation Process Guidelines (Invitrogen), and the K. lactis Protein Expression Kit (New England Biolabs). Autoclave sterilization is performed at 121°C for 15 min. 2.1. Uniform 13C, 15 N-Labeling Using P. pastoris

1. 40% unlabeled D-glucose stock solution: Dissolve 200 g of D-glucose in 1 L of water. Sterilize by aseptic filtration and store at room temperature. The shelf life of this solution is approximately 1 year. 2. 100 mg/mL Stock solution of Zeocin™ (Invitrogen). 3. Yeast extract peptone dextrose sorbitol (YPDS) agar plates: Dissolve 20 g of bacto peptone, 10 g of yeast extract, 182.2 g of sorbitol, and 20 g of bacto agar in 950 mL of water. Sterilize by autoclaving and cool to 50–60°C. Aseptically add 1 mL of 100 mg/mL Zeocin™ and 50 mL of 40% glucose stock solution, gently mix well, and dispense into sterile disporsable Petri dishes. Store at 4°C in the dark. The shelf life of these agar plates is 1–2 weeks. 4. 10× Yeast nitrogen base (YNB): Dissolve 34 g of YNB without amino acids and ammonium sulfate in 1 L of water (see Note 1). Warm it to 40–50°C to dissolve completely if needed. Sterilize by aseptic filtration and store at 4°C. The shelf life of this solution is approximately 1 year. 5. 1 M Potassium phosphate: Mix 132 mL of 1 M K2HPO4 and 868 mL of 1 M KH2PO4. Adjust the pH to 6.0 ± 0.1 using KOH or phosphoric acid. Sterilize by autoclaving and store at room temperature. The shelf life of this solution is greater than 1 year. 6. 500× Biotin: Dissolve 20 mg of biotin in 100 mL of water and warm it to 40–50°C to dissolve completely. Sterilize by aseptic filtration and store at 4°C. The shelf life of this solution is approximately 1 year. 7. 10% (w/v) Glycerol: Dissolve 50 mL of glycerol in 450 mL of water. Sterilize by aseptic filtration and store at room temperature. The shelf life of this solution is over 1 year. 8. 200× PTM1 trace salts: Mix together the following ingredients and dissolve to a final volume of 1 L in water: 6.0 g of CuSO 4⋅5H2O, 0.08 g of NaI, 3.0 g of MgSO4⋅H2O, 0.2 g of Na2MoO4⋅2H2O, 0.02 g of boric acid, 0.5 g of CoCl2, 20.0 g of ZnCl2, 65.0 g of FeSO4⋅7H2O, 0.2 g of biotin, 5.0 mL of H2SO4. Warm to 40–50°C to dissolve completely if needed. Sterilize by aseptic filtration and store at 4°C or room temperature. The shelf life of this solution is approximately 3 months at 4°C.

2

Isotopic Labeling of Heterologous Proteins in the Yeast…

23

9. Buffered glycerol-complex (BMGY) medium: Dissolve 20 g bacto peptone and 10 g yeast extract in 700 mL of water. Sterilize by autoclaving and cool it to room temperature. Aseptically add 100 mL of sterile 1 M potassium phosphate, 100 mL of sterile 10× YNB, 2 mL of sterile 500× biotin, and 100 mL of sterile 10% (w/v) glycerol solutions. If needed, 5 mL of 200× PTM1 trace salts solution can be added to this medium. Store at 4°C. The shelf life of this medium is approximately 3 months. 10. 10% 15N-ammonium chloride: Dissolve 6 g of 15N-ammonium chloride (98 atom% 15N) in 60 mL of water (see Note 2). Sterilize by aseptic filtration. Prepare this solution just before use. 11.

15

N-BM medium: To prepare 1.2 L of uniformly 15N-labeled buffered minimal (15N-BM) medium, autoclave 900 mL of water in a 2-L media bottle and allow the water to cool to room temperature. Aseptically add 120 mL of 10× YNB, 2.4 mL of 500× biotin, 120 mL of 1 M potassium phosphate, and 60 mL of 10% 15N-ammonium chloride to the autoclaved water. If needed, 6 mL of 200× PTM1 trace salts solution can be added to 15N-BM medium.

12. 5% D-[13C6]glucose: Dissolve 1 g of D-[13C6]glucose (99 atom% 13 C) in 20 mL of 15N-BM medium and sterilize by aseptic filtration. Prepare this solution just before use. 13. Antifoaming agent: (see Note 3). 14.

13

C-methanol (99 atom% 13C).

15. 50% (w/v) ethanol. Ethanol is just necessary for immersing air outlet port of the fermentation system. 2.2. Uniform 2H, 15 N-Labeling Using P. pastoris

1. YPDS agar plates (Subheading 2.1). 2. BMGY medium (Subheading 2.1). 3. 10× YNB(2H2O): Prepare this solution in a manner similar to 10× YNB solution (Subheading 2.1), but use 2H2O (99.8 atom % 2H) instead of water (H2O). Sterilize by aseptic filtration and store at room temperature. The shelf life of this solution is approximately 2–3 months. 4. 1 M potassium phosphate(2H2O): Prepare 150 mL of this solution in a manner similar to 1 M potassium phosphate solution (Subheading 2.1), but use 2H2O instead of water. Sterilize by aseptic filtration. Prepare this solution just before use. 5. 500× biotin(2H2O): Prepare this solution in a manner similar to 500× biotin solution (Subheading 2.1), but use 2H2O instead of water. Sterilize by aseptic filtration and store at 4°C. The shelf life of this solution is approximately 2–3 months. 6. 200× PTM1 trace salts(2H2O): Prepare this solution in a manner similar to 200× PTM1 trace salts solution (Subheading 2.1), but use 2H2O instead of water. Sterilize by aseptic filtration and store at 4°C.

24

T. Sugiki et al.

7. 10% 15N-ammonium chloride(2H2O): Dissolve 7 g of 15 N-ammonium chloride in 70 mL of 2H2O (see Note 2) and sterilize by aseptic filtration. Prepare this solution just before use. 8. 10% unlabeled D-glucose(2H2O): Dissolve 1 g of D-glucose in 10 mL of 2H2O and sterilize by aseptic filtration. Prepare this solution just before use. 9. 15N-BMD (90% 2H2O) medium: To prepare 10 mL of uniformly 15 N-labeled buffered minimal medium containing unlabeled glucose prepared in 90% 2H2O, mix 6 mL of 2H2O and 1 mL of H2O, and sterilize by aseptic filtration. Aseptically add 1 mL of 10× YNB(2H2O), 0.02 mL of 500× biotin(2H2O), 1 mL of 1 M potassium phosphate(2H2O), 0.5 mL of 10% unlabeled D-glucose(2H2O) (0.5% final concentration), 0.5 mL of 10% 15 N-ammonium chloride(2H2O), and 5–10 μL of antifoaming agents. If needed, 0.05 mL of 200× PTM1 trace salts(2H2O) can be added to this medium. Prepare this medium just before use. 10.

15

N-BM(2H2O) medium: To prepare 1.3 L of uniformly N-labeled buffered minimal medium in 100% 2H2O, sterilize 956 mL of fresh 2H2O by aseptic filtration. Aseptically add 130 mL of 10× YNB(2H2O), 3 mL of 500× biotin(2H2O), 130 mL of 1 M potassium phosphate(2H2O), and 66 mL of 10% 15N-ammonium chloride(2H2O) into the sterile 2H2O. If needed, 7 mL of 200× PTM1 trace salts(2H2O) can be added to this medium. Prepare this medium just before use. 15

11.

15

N-BMD(2H2O) medium: To prepare 5 mL of uniformly N-labeled buffered minimal medium containing unlabeled glucose in 100% 2H2O, aseptically add 0.25 mL of 10% unlabeled D-glucose(2H2O) to 4.75 mL of 15N-BM(2H2O) in a 50-mL Corning tube. Prepare this medium just before use. 15

12. 10% D-[2H7]glucose(2H2O): Dissolve 1.5 g of D-[2H7]glucose (98 atom% 2H) in 15 mL of 2H2O and sterilize by aseptic filtration. Prepare this solution just before use. 13. 2H, 15N-BMD(2H2O) medium: To prepare 300 mL of uniformly 15 N-labeled buffered minimal medium containing D-[2H7]glucose in 100% 2H2O, transfer 285 mL of 15N-BM(2H2O) into a sterilize, well-dried 1-L baffled flask. Aseptically add 15 mL of 10% D-[2H7]glucose(2H2O) and 0.15–0.30 mL of antifoaming agents. Prepare this medium just before use. 14. Antifoaming agent (Subheading 2.1). 15. [2H4]methanol (99.5 atom% 2H). 16. 50% (w/v) ethanol. Ethanol is just necessary for immersing air outlet port of the fermentation system. 2.3. Uniform 13C, 15 N-Labeling Using K. lactis

1. Yeast peptone (YP): Dissolve 20 g of bacto peptone and 10 g of yeast extract in 950 mL of water. Sterilize by autoclaving and store at room temperature or 4°C. The shelf life of this solution is approximately 1 year.

2

Isotopic Labeling of Heterologous Proteins in the Yeast…

25

2. 40% unlabeled D-glucose stock solution (Subheading 2.1). 3. YPD medium: Aseptically add 50 mL of sterile 40% D-glucose stock solution to the autoclaved 950 mL YP solution. The shelf life of this solution is approximately 1 month. 4. 1 M potassium phosphate (Subheading 2.1). 5. YCB-acetamide agar plates: Dissolve 5.85 g of yeast carbon base (YCB) medium powder and 10 g of bacto agar in 470 mL of water. Add 25 mL of 1 M potassium phosphate solution. Autoclave and cool it to 50–60°C. Aseptically add 5 mL of 100× acetamide stock solution (supplied by New England Biolabs), gently mix well, and dispense into sterile disporsable Petri dishes. Store at 4°C. The shelf life of these plates is approximately 3 months. 6. 10× YNB (Subheading 2.1). 7. 500× biotin (Subheading 2.1). 8. 200× PTM1 trace salts (Subheading 2.1). 9. 10% 15N-ammonium chloride: Dissolve 7.5 g of 15N-ammonium chloride in 75 mL of water (see Note 2) and sterilize by aseptic filtration. Prepare this solution just before use. 10. 4% D-[13C6]glucose: Dissolve 2 g of D-[13C6]glucose in 50 mL of water and sterilize it by aseptic filtration. Prepare this solution just before use. 11.

13

C, 15N-BMD medium: To prepare 0.5 L of 15N-BM medium containing 0.4% D-[13C6]glucose, autoclave 325 mL of water containing 0.25–0.50 mL of antifoaming agents in a 2-L fermentation vessel and allow it to cool to room temperature. Aseptically add 50 mL of 10× YNB, 1 mL of 500× biotin, 50 mL of 1 M potassium phosphate, 25 mL of 10% 15N-ammonium chloride, and 50 mL of 4% D-[13C6]glucose to the autoclaved water for a final volume of 500 mL. If needed, 2.5 mL of 200× PTM1 trace salts solution can be added to this medium.

12. 6% D-[13C6]glucose: Dissolve 6 g of D-[13C6]glucose in 100 mL of water and sterilize by aseptic filtration. Prepare this solution just before use. 13. 13C, 15N-BMD feeding medium: To prepare 1 L of 15N-BM medium containing 0.6% D-[13C6]glucose feeding medium, autoclave 650 mL of water containing 0.5–1.0 mL of antifoaming agents in a 1-L media bottle and allow it to cool to room temperature. Aseptically add 100 mL of 10× YNB, 2 mL of 500× biotin, 100 mL of 1 M potassium phosphate, 50 mL of 10% 15 N-ammonium chloride and 100 mL of 6% D-[13C6]glucose to the autoclaved water for a total volume of 1 L. If needed, 5 mL of 200× PTM1 trace salts solution can be added to this medium. 14. Antifoaming agent (Subheading 2.1). 15. 50% (w/v) ethanol. Ethanol is just necessary for immersing air outlet port of the fermentation system.

26

T. Sugiki et al.

2.4. Uniform 2H, 15 N-Labeling Using K. lactis

1. YPD medium (Subheading 2.3). 2. YCB-acetamide agar plates (Subheading 2.3): If needed, prepare the plates with 2H2O (see Note 4). 3. 10× YNB(2H2O) (Subheading 2.2). 4. 500× biotin(2H2O) (Subheading 2.2). 5. 1 M potassium phosphate(2H2O) (Subheading 2.2). 6. 200× PTM1 trace salts(2H2O) (Subheading 2.2). 7. 10% 15N-ammonium chloride(2H2O): Dissolve 7.5 g of 15 N-ammonium chloride in 75 mL of 2H2O (see Note 2) and sterilize by aseptic filtration. Prepare this solution just before use. 8. 4% D-[2H7]glucose(2H2O): Dissolve 2 g of D-[2H7]glucose in 50 mL of 2H2O and sterilize by aseptic filtration. Prepare this solution just before use. 9. 2H, 15N-BMD(2H2O): To prepare 0.5 L of uniformly 15 N-labeled buffered minimal medium containing D-[2H7]glucose in 2H2O, sterilize 325 mL of 2H2O by aseptic filtration. Aseptically add 50 mL of 10× YNB(2H2O), 1 mL of 500× biotin(2H2O), 50 mL of 1 M potassium phosphate(2H2O), 25 mL of 10% 15N-ammonium chloride(2H2O), 50 mL of 4% D-[2H7]glucose(2H2O), and 0.25–0.50 mL of antifoaming agent to the sterilized 2H2O for a final volume of 500 mL. If needed, 2.5 mL of 200× PTM1 trace salts(2H2O) can be added to this medium. 10. 6% D-[2H7]glucose(2H2O): Dissolve 6 g of D-[2H7]glucose in 100 mL of 2H2O and sterilize by aseptic filtration. Prepare this solution just before use. 11. 2H, 15N-BMD(2H2O) feeding medium: Sterilize 650 mL of 2 H2O by aseptic filtration. Aseptically add 100 mL of 10× YNB(2H2O), 2 mL of 500× biotin(2H2O), 100 mL of 1 M potassium phosphate(2H2O), 50 mL of 10% 15N-ammonium chloride(2H2O), 100 mL of 6% D-[2H7]glucose(2H2O), and 0.5–1.0 mL of antifoaming agent to the sterilized 2H2O for a final volume of 1 L. If needed, 5 mL of 200× PTM1 trace salts(2H2O) can be added to this medium. 12. Antifoaming agent (Subheading 2.1). 13. 50% (w/v) ethanol. 13. 50% (w/v) ethanol. Ethanol is just necessary for immersing air outlet port of the fermentation system.

3. Methods 3.1. Uniform 13C, 15 N-Labeling Using P. pastoris

1. Prepare necessary media. (a) Using aluminum foil, loosely cover the open lid of an empty 1-L baffled flask and autoclave it.

2

Isotopic Labeling of Heterologous Proteins in the Yeast…

27

(b) Prepare 1.2 L of fresh, sterile 15N-BM medium in a 2-L medium bottle and 20 mL of 5% D-[13C6]glucose. (c) Aseptically combine 180 mL of fresh 15N-BM medium, 20 mL of 5% D-[13C6]glucose and 0.1–0.2 mL of antifoaming agents in the sterile 1-L baffled flask to make 13 C, 15N-BMD medium. (d) Using aluminum foil, loosely mask the open lid of the empty 2-L fermentation vessel. Clamp all of the lines that could come into contact with the growth medium and autoclave the vessel. (e) Aseptically combine 1 L of fresh 15N-BM medium, 0.5–1.0 mL of antifoaming agent and 5 mL of 13C-methanol into the sterile empty 2-L fermentation vessel, to make 13 C, 15N-BMM medium. 2. Using a sterile loop, scoop a small aliquot of P. pastoris transformants from a frozen glycerol stock, and streak it onto a YPDS agar plate containing Zeocin™. Incubate the plate for 24–48 h at 30°C. 3. To produce the primary culture, inoculate a fresh, single colony of the P. pastoris transformants into 5 mL of BMGY medium (in a 50-mL Corning tube) and shake at 200–250 rpm at 30°C for 18–24 h. 4. Pellet the primary culture cells by centrifugation at 3,000 × g for 5 min at 20°C and discard the supernatant. Gently resuspend the pellet with 30 mL of fresh, sterile 13C, 15N-BMD medium, and pour the resuspended cells into the remaining 13 C, 15N-BMD medium in the baffled flask for a final volume of 200 mL (see Note 5). Shake the flask at > 200 rpm at 30°C until the cell density reaches an OD600 of 4–6. 5. Pellet the cells in sterile tubes by centrifuging at 2,000 × g for 10 min at 20°C and discard the supernatant. Gently resuspend the pellet with 30 mL of fresh, sterile 13C, 15N-BMM medium, and pour the resuspended cells into the remaining fresh, sterile 13 C, 15N-BMM medium in the 2-L fermentation vessel to reach an OD600 of approximately 1 (see Notes 6–8). 6. Assemble the fermentation system. Aseptically attach the pH and dissolved O2 sensors to the fermentation vessel and connect the probes to their respective controllers. Connect the outlet port of the air-feeding tube to the air flow inlet port of the vessel. Aseptically attach a Liebig condenser to the outlet port of the vessel, and immerse the air-exhaust port of the Liebig condenser under 1 L of 50 % (w/v) ethanol solution. Insert the temperature probe into the vessel and connect it to a temperature controller, such as a circulating water bath. 7. Agitate the culture medium at 300–800 rpm with 0.1–0.3 L/ min of feeding air at 30°C (see Notes 9 and 10). Add

28

T. Sugiki et al. 13

C-methanol (0.5 % (w/v) of the total volume) to the culture medium every 24 h during this induction phase (20) (see Note 11). The total amount of 13C-methanol required for sufficient protein expression in a cost-effective manner is approximately 10–20 mL per 1 L culture medium (see Note 12). 8. Pellet the cells by centrifuging at 6,000 × g for 20 min at 4°C. If the target protein is secreted into the culture medium, filter the supernatant by passing it through a < 0.45-μm pore size membrane and collect the filtrate. If needed, add appropriate protease inhibitors to the filtrate to prevent proteolytic degradation of secreted target proteins. If the target protein is expressed in the cytoplasm of the host cells, discard the supernatant after centrifugation and retain the cell pellet. 3.2. Uniform 2H, 15 N-Labeling Using P. pastoris

Deuterium isotope-labeling of target proteins is one of the most important techniques for protein NMR studies, especially for analyses of large molecular weight (MW > 25 K) proteins and in crosssaturation experiments to identify intermolecular-binding sites (6, 21–24). The P. pastoris expression system can be used to overexpress deuterium-labeled heterologous proteins. For the efficient production of deuterated target proteins, cells should be adapted to grow in deuterated broth medium. Adaptation is achieved by multistage subculturing in which the deuterium concentration is raised in a stepwise fashion. For instance, subculturing of cells is performed using semideuterated (25–95% 2H2O) medium, with subsequent culturing in fully deuterated medium (14) (see Note 4). In the case of uniform 2H-labeling, using a 2 H-labeled carbon source (such as [2H4]methanol) only during the induction phase is insufficient for producing fully deuterated target proteins (a considerable amount of protons remain on methyl and Lys δ/ε groups) (14). To achieve the nearly-complete deuteration level required for cross-saturation experiments, a deuterium-labeled carbon source, such as D-[2H7]glucose should also be used during the cell growth phase (prior to the induction phase) (7, 14). In this section, we describe a cultivation procedure for overexpressing perdeuterated heterologous proteins in P. pastoris. Using this procedure, Ichikawa and coworkers successfully prepared a discoidin domain of DDR2 that was more than 95% deuterated, and clearly identified the collagen-binding site of DDR2 through transferred cross-saturation experiments (7). 1. Prepare necessary media: (a) Autoclave a 1-L baffled flask and a 2-L fermentation vessel, and dry them in an oven at 60–80°C (for at least 48 h) to completely remove residual H2O. (b) Prepare 1.3 L of 15N-BM(2H2O) medium in a 2-L media bottle, 10 mL of 10% unlabeled D-glucose(2H2O), and 15 mL of 10% D-[2H7]glucose(2H2O) solutions.

2

Isotopic Labeling of Heterologous Proteins in the Yeast…

29

(c) Prepare 5 mL each of 15N-BMD(90% 2H2O), 15N-BMD (2H2O), and 2H, 15N-BMD(2H2O) media in 50-mL Corning tubes. (d) Prepare 300 mL of 2H, 15N-BMD(2H2O) medium in a sterilize, well-dried 1-L baffled flask. (e) Prepare 2H, 15N-BMM(2H2O) medium by mixing 1 L of 15 N-BM(2H2O) medium, 0.5–1.0 mL of antifoaming agent and 5 mL of [2H4]methanol in the sterilize, well-dried 2-L fermentation vessel. 2. Using a sterile loop, scoop a small aliquot of P. pastoris transformants from a frozen glycerol stock and streak it onto a YPDS agar plate containing Zeocin™. Incubate the plate for 24–48 h at 30°C. 3. To produce the primary culture, inoculate 5 mL of BMGY medium in a sterile 50-mL Corning tube with a fresh single colony of the P. pastoris transformant from the YPDS agar plate and shake at 200–250 rpm at 30°C for 18–24 h. 4. Add 0.1 mL of the primary culture to 5 mL of fresh, sterile 15 N-BMD(90% 2H2O) medium in a sterile 50-mL Corning tube. Shake the culture tube at 200–300 rpm at 30°C until the cell density reaches an OD600 of approximately 3–5. 5. Add 0.1 mL of the step 4 culture to 5 mL of fresh, sterile 15 N-BMD(2H2O) medium (in a sterile 50-mL Corning tube). Shake the culture tube at 200–300 rpm at 30°C until the cell density reaches an OD600 of 3–5. 6. Add 0.1 mL of the step 5 culture to 5 mL of fresh, sterile 2H, 15 N-BMD(2H2O) medium in a sterile 50-mL Corning tube. Shake the culture tube at 200–300 rpm at 30°C until the cell density reaches an OD600 of 3–5. 7. Pellet the cells by centrifuging at 3,000 × g for 5 min at 20°C and discard the supernatant. Gently resuspend the pellet with 30 mL of fresh, sterile 2H, 15N-BMD(2H2O) medium and pour the resuspended cells into the remaining 2H, 15N-BMD(2H2O) medium in the 1-L baffled flask to a final volume of 300 mL. Shake the flask at > 200 rpm at 30°C until the cell density reaches an OD600 of 3–4. 8. Pellet the cells in sterile tubes by centrifuging at 2,000 × g for 10 min at 20°C and discard the supernatant. Gently resuspend the pellet with 30 mL of fresh, sterile 2H, 15N-BMM(2H2O) medium and pour the resuspended cells into the remaining sterile 2H, 15N-BMM(2H2O) medium in the 2-L fermentation vessel. The starting OD600 of the culture in the fermenter vessel should be approximately 1 (see Notes 6–8). 9. Assemble the fermentation system Subheading 3.1, step 6 (see Note 13).

as

described

in

30

T. Sugiki et al.

10. Agitate the culture medium at 300–800 rpm with 0.1–0.3 L/min of feeding air at 30°C (see Notes 9 and 10). Add [2H4] methanol (0.5 % (w/v) of the total volume) to the culture medium every 24 h during this induction phase (see Notes 11 and 12). 11. Collect the target proteins as described in Subheading 3.1, step 8. 3.3. Uniform 13C, 15 N-Labeling Using K. lactis

1. Prepare 0.5 L of sterile 13C, 15N-BMD and 1.0 L of 13C, 15N-BMD feeding medium in a 2-L fermentation vessel and 1-L media bottle, respectively. 2. Using a sterile loop, scoop a small aliquot of K. lactis transformants from a frozen glycerol stock and streak it onto a YCBacetamide agar plate. Incubate the plate for 24–48 h at 30°C. 3. To produce the primary culture, inoculate a fresh, single colony of K. lactis transformants into 5 mL of YPD medium (in a 50-mL Corning tube) and shake at 200–250 rpm at 30°C for 48 h to obtain a saturated biomass (OD600 > 20–30). 4. Assemble the fermentation system as described in Subheading 3.1, step 6. Connect the media bottle containing 1 L of fresh 13C, 15N-BMD feeding medium to the inlet port of the vessel with an appropriate length of feeding tube, and attach a peristaltic pump at the midpoint of the feeding tube (Fig. 1). 5. Pellet the primary culture cells by centrifuging at 3,000 × g for 5 min at 20°C and discard the supernatant. Gently resuspend the pellet with 30 mL of fresh, sterile 13C, 15N-BMD medium from the 2-L fermentation vessel, and pour the resuspended cells into the remaining 13C, 15N-BMD medium in the fermentation vessel. Agitate the culture medium at 600–800 rpm with 0.1–0.3 L/min feeding air at 30°C (see Notes 9, 10, and 12). 6. During the fermentation, continuously feed fresh, sterile 13C, 15 N-BMD feeding medium into the fermentation vessel at a constant flow rate of 8.3 mL/h using the peristaltic pump (see Note 14). 7. Pellet the cells by centrifuging at 10,000 × g for 15 min at 4°C. If the target protein is secreted into the culture medium, filter the supernatant by passing it through a < 0.45-μm pore size membrane and collect the filtrate. If needed, add appropriate protease inhibitors to the collected filtrate to prevent proteolytic degradation of the target protein. If the target protein is expressed in the cytoplasm of the host cells, discard the supernatant after centrifugation, and retain the cell pellet.

3.4. Uniform 2H, 15 N-Labeling Using K. lactis

In our experience, perdeuteration (approximately 90% deuteration estimated by MS analysis) of maltose-binding protein (MBP) is successfully achieved using K. lactis with fed-batch fermentation.

2

Isotopic Labeling of Heterologous Proteins in the Yeast…

31

Fig. 1. Overview of the fed-batch fermentation system. (a) Schematic diagram of fed-batch fermentation. (b) Photograph of the assembled fed-batch fermentation device. a: fresh media which is continuously fed into the fermentation vessel, b: peristaltic pump, c: fermentation vessel, d: air flow inlet, and e: air exhaust port.

Furthermore, insertion of one or two subcultivation steps following the primary culture improves cell growth and expression of MBP with a highly efficient (> 92%) level of deuteration (Miyazawa-Onami and coworkers, unpublished data). 1. Prepare necessary media: (a) Autoclave an empty 1-L media bottle and a 2-L fermentation vessel, and dry them in an oven at 60–80°C (for at least 48 h) to completely remove residual H2O. (b) Prepare 0.5 L of fresh, sterile 2H, 15N-BMD(2H2O) and 1.0 L of 2H, 15N-BMD(2H2O) feeding medium in the sterile 2-L fermentation vessel and 1-L media bottle, respectively. 2. Using a sterile loop, scoop a small aliquot of K. lactis transformants from a frozen glycerol stock and streak it onto a YCBacetamide agar plate. Incubate the plate for 24–48 h at 30°C.

32

T. Sugiki et al.

3. To produce the primary culture, inoculate 5 mL of YPD medium in a 50-mL Corning tube using a fresh single colony of the K. lactis transformants from the YCB agar plate and shake at 200–250 rpm at 30°C for 48 h to obtain a saturated biomass (OD600 > 20–30). 4. Assemble the fermentation system as described in Subheading 3.1, step 6 (see Note 13). Connect the bottle containing 1 L of fresh 2H, 15N-BMD(2H2O) feeding medium to the inlet port of the vessel with an appropriate length of feeding tube, and attach a peristaltic pump at the midpoint of the feeding tube (Fig. 1). 5. Pellet the cells by centrifuging at 3,000 × g for 5 min at 20°C and discard the supernatant. Gently resuspend the pellet with 30 mL of fresh, sterile 2H, 15N-BMD(2H2O) medium from the 2-L fermentation vessel, and pour the resuspended cells into the remaining 2H, 15N-BMD(2H2O) medium in the fermentation vessel. Agitate the culture medium at 600–800 rpm with 0.1–0.3 L/min feeding air at 30°C (see Notes 9, 10, and 12). 6. During the fermentation, continuously feed fresh, 2H, 15 N-BMD(2H2O) feeding medium from the 1-L media bottle into the fermentation vessel at a constant flow rate of 8.3 mL/h using the peristaltic pump (see Note 14). 7. Collect the target proteins as described in Subheading 3.3, step 7.

4. Notes 1. Detailed information about the composition of YNB without amino acids and ammonium sulfate can be found in the Becton Dickinson Difco Yeast Media Recipes, available at http://www. bd.com/ds/technicalCenter/inserts/Yeast_Media.pdf. 2. Ammonium sulfate or ammonium chloride is used as a nitrogen source by the yeast. In many cases, 5–10 g/L of ammonium sulfate or ammonium chloride are used in the culture of yeast. In our experience, a sufficient amount of isotope-labeled protein could be expressed while keeping costs to a minimum by using 2 g/L of 15N-ammonium chloride (7). 3. Foaming of the culture medium caused by agitation and aeration will severely affect the yield of target protein due to a reduction in the level of protein expression and partial denaturation of the expressed proteins (25). Foaming should be controlled

2

Isotopic Labeling of Heterologous Proteins in the Yeast…

33

by adding antifoaming agents to the culture medium. We use 0.05–0.10% (w/v) of “Antifoam 204” (supplied from Sigma). 4. For many organisms, 2H2O-containing medium negatively affect physiological processes. In many cases, stagnation of cell growth occurs in 75–100% 2H2O, necessitating adaptation to the deuterated medium by subcultivating the cells several times prior to the induction of protein expression (2). Furthermore, the selection of transformants using deuterated agar plates may be effective in generating special transformants that are more adaptable to deuterated medium. 5. Although glycerol is widely used as a carbon source of P. pastoris during growth phase, in the case of 13C-labeling, 13C-glycerol can be replaced by 13C-glucose to reduce labeling costs (9, 26, 27). 6. An OD600 = 1 is equivalent to 5 × 107 P. pastoris cells (2). 7. Nonmethanolic carbon sources, especially glucose, should be completely eliminated (or consumed) prior to the addition of methanol to achieve full induction efficiency of the AOX1 promoter (2, 7, 20). However, the use of mixtures of carbon sources during the induction phase to improve cell growth and the production of target proteins has been reported (28–32). 8. The optimal starting point of the induction phase should be determined by performing a time course study of cell growth in a “cold-run” prior to beginning a “hot-run,” since the optimal point for starting induction varies according to the type of target protein and culture conditions. 9. The level of oxygen supplied to the fermentation medium should be kept at 0.1–0.3 vvm (volume of oxygen (liters) per volume of fermentation culture (liters) per minute) during the cultivation. This level can be easily achieved using any glass fermenter and mixing the broth medium via an impeller in an air feeding condition. Maintaining an adequate concentration of dissolved O2 during cultivation is crucially important for sufficient cell growth and protein expression (2, 20, 33). 10. Proteolytic degradation of heterologous proteins is one of the drawbacks of the secretory expression strategy. In our experience, this often occurs in a culture grown in a poor-nutrient medium such as the one that may be found under isotope labeling conditions. To prevent extensive proteolytic degradation of target proteins, the following measures can be taken (1) optimizing the pH range of the culture medium (34); (2) cultivating at low-temperature (35–38); (3) adding extracts of algae, such as BioExpress. (26); (4) adding protease inhibitors (2, 27, 39); however, some types of protease inhibitors are toxic; and (5) utilizing protease-deficient strains (34, 40). Several

34

T. Sugiki et al.

protease-deficient strains of P. pastoris and K. lactis are commercially available. 11. In the P. pastoris expression system, the methanol concentration during induction directly affects cell growth and protein production since an excess of methanol (> 1–2% (w/v)) and an excessive rate of feeding is toxic to cells (5). Monitoring the level of dissolved O2 in the culture medium is a convenient method for assessing the timing for feeding and the appropriate dose of methanol because sharp “spike” signals are detected by the O2 electrode when the carbon sources are completely consumed and/or when the concentration of methanol reaches a toxic level (20, 33). 12. The typical induction time is between 48 and 96 h. However, in the case of a deuterium-labeling culture, an induction period longer than 96 h is typically required. The optimal induction period and the amount of methanol should be determined by following the time course of target protein expression level using SDS-PAGE. 13. To achieve a higher level of deuteration, a glass tube filled with dried granular calcium chloride should be connected in front of the air inlet of the fermentation vessel to remove H2O from the supplied air. 14. The optimal culture conditions (composition of media, pH, temperature, etc.) vary according to the type of host and target proteins. In our experience, the most critical factors for obtaining sufficient protein expression using K. lactis are the concentration of the carbon source and the concentration of dissolved oxygen. Especially, in cases involving the use of minimal media for isotope labeling, we recommend fermentation cultivation rather than flask culture. Furthermore, addition of isotopically labeled extracts of algae, such as Celtone, BioExpress (both supplied by Cambridge Isotope Laboratories, Inc.) or C.H.L. (Chlorella Industry Co., Ltd.) significantly improves cell growth and protein expression (41).

Acknowledgments This work was financially supported in part by the Ministry of Economy, Trade and Industry (METI) and the New Energy and Industrial Technology Development Organization (NEDO).

2

Isotopic Labeling of Heterologous Proteins in the Yeast…

35

References 1. Cregg, J. M., Vedvick, T. S., and Raschke, W. C. (1993) Recent advances in the expression of foreign genes in Pichia pastoris. Biotechnology 11, 905–910. 2. Pickford, A. R., and O’Leary, J. M. (2008) Isotopic labeling of recombinant proteins from the methylotrophic yeast Pichia pastoris, in Methods in Molecular Biology (Downing, A. K., Ed.), vol. 278, pp. 17–33, Humana Press, Totowa, NJ. 3. Lin Cereghino, G. P., Cereghino, J. L., Ilgen, C., and Cregg, J. M. (2002) Production of recombinant proteins in fermenter cultures of the yeast Pichia pastoris. Curr. Opin. Biotechnol. 13, 329–332. 4. Daley, R., and Hearn, M. T. (2005) Expression of heterologous proteins in Pichia pastoris : a useful experimental tool in protein engineering and production. J. Mol. Recognit. 18, 119–138. 5. Cos, O., Ramόn, R., Montesinos, J. L., and Valero F. (2006) Operational strategies, monitoring and control of heterologous protein production in the methylotrophic yeast Pichia pastoris under different promoters: A review. Microbial Cell Factories 5, 17–36. 6. Takahashi, H., and Shimada, I. (2010) Production of isotopically labeled heterologous proteins in non-E. coli prokaryotic cells. J. Biomol. NMR 46, 3–10. 7. Invitrogen Corporation. EasySelect™ Pichia Expression Kit: A manual of methods for expression of recombinant proteins using pPICZ and pPICZα in Pichia pastoris, Version I. Available at http://tools.invitrogen.com/content/sfs/manuals/easyselect_man.pdf. 8. Laroche, Y., Strome, V., De Meutter, J., Messens, J., and Lauwereys, M. (1994) Highlevel secretion and very efficient isotopic labeling of tick anticoagulant peptide (TAP) expressed in the methylotrophic yeast, Pichia pastoris. Bio/Technol 12, 1119–1124. 9. Denton, H., Smith, M., Husi, H., Uhrin, D., Barlow, P. N., Batt, C. A. , and Sawyer, L. (1998) Isotopically labeled bovine β-lactoglobulin for NMR studies expressed in Pichia pastoris. Protein Expr. Purif. 14, 97–103. 10. Wood, M. J., and Komives, E. A. (1999) Production of large quantities of isotopically labeled protein in Pichia pastoris by fermentation.. J. Biomol. NMR 13, 149–159. 11. Mine, S., Ueda, T., Hashimoto, Y., Tanaka, Y., and Imoto, T. (1999) High-level expression of uniformly 15N-labeled hen lysozyme in Pichia pastoris and identification of the site in hen

lysozyme where phosphate ion binds using NMR measurements. FEBS Lett. 448, 33–37. 12. van den Burg, H. A., de Wit, P. J., and Vervoort, J. (2001) Efficient 13C/15N double labeling of the avirulence protein AVR4 in a methanolutilizing strain (Mut+) of Pichia pastoris. J. Biomol. NMR 20, 251–261. 13. Rodriguez, E., and Krishna, N. R. (2001) An economical method for 13C/15N isotopic labeling of proteins expressed in Pichia pastoris. J. Biochem. 130, 19–22. 14. Morgan, W. D., Kragt, A., and Feeney, J. (2000) Expression of deuterium-isotope-labelled protein in the yeast Pichia pastoris for NMR study. J. Biomol. NMR 17, 337–347. 15. Ichikawa, M., Osawa, M., Nishida, N., Goshima, N., Nomura, N., and Shimada, I. (2007) Structural basis of the collagen-binding mode of discoidin domain receptor 2, EMBO J. 26, 4168–4176. 16. New England Biolabs, Inc. K. lactis Protein Expression Kit: Instruction manual. Available at http:/ /www.neb.com/nebecomm/ManualFiles/ manualE1000.pdf. 17. Colussi, P. A., and Taron, C. H. (2005) Kluyveromyces lactis LAC4 promoter variants that lack function in bacteria but retain full function in K. lactis, Appl. Environ. Microbiol. 71, 7092–7098. 18. Read, J. D., Colussi, P. A., Ganatra, M. B., and Taron, C. H. (2007) Acetamide selection of Kluyveromyces lactis cells transformed with an integrative vector leads to high-frequency formation of multicopy strains. Appl. Environ. Microbiol. 73, 5088–5096. 19. Sugiki, T., Shimada, I., and Takahashi, H. (2008) Stable isotope labeling of protein by Kluyveromyces lactis for NMR study. J. Biomol. NMR 42, 159–162. 20. Invitrogen Corporation. Pichia Fermentation Process Guidelines. Available at http://toolsja. invitrogen.com/content/sfs/manuals/pichiaferm_prot.pdf. 21. Takahashi, H., Nakanishi, T., Kami, K., Arata, Y., and Shimada, I. (2000) A novel NMR method for determining the interface of large protein-protein complexes. Nat. Struct. Biol. 7, 220–223. 22. Nakanishi, T., Miyazawa, M., Sakakura, M., Terasawa, H., Takahashi, H., and Shimada, I. (2002) Determination of the interface of a large protein complex by transferred cross-saturation measurements. J. Mol. Biol. 318, 245–249. 23. Takahashi, H., Miyazawa, M., Ina, Y., Fukunishi, Y., Mizukoshi, Y., Nakamura, H., and Shimada,

36

T. Sugiki et al.

I. (2006) Utilization of methyl proton resonances in cross-saturation measurement for determining the interfaces of large protein-protein complexes. J. Biomol. NMR 34, 167–177. 24. Shimada, I., Ueda, T., Matsumoto, M., Sakakura, M., Osawa, M., Takeuchi, K., Nishida, N., and Takahashi, H. (2008) Cross-saturation and transferred cross-saturation experiments. Prog. Nucl. Magn. Reson. Spectrosc. 54, 123–140. 25. Koch, V., Rüffer, H.-M., Schügerl, K., Innertsberger, E., Menzel, H., and Weis, J. (1995) Effect of antifoam agents on the medium and microbial cell properties and process performance in small and large reactors. Process Biochem. 30, 435–446. 26. Macauley-Patrick, S., Fazenda, M. L., McNeil, B., and Harvey, L. M. (2005) Heterologous protein production using the Pichia pastoris expression system. Yeast 22, 249–270. 27. Shapiro, R. I., Wen, D., Levesque, M., Hronowski, X., Gill, A., Garber, E. A., Galdes, A., Strauch, K. L., and Taylor, F. R. (2003) Expression of sonic hedgehog-Fc fusion protein in Pichia pastoris. Identification and control of post-translational, chemical, and proteolytic modifications. Protein Expr. Purif. 29, 272–283. 28. Files, D., Ogawa, M., Scaman, C. H., and Baldwin, S. A. (2001) A Pichia pastoris fermentation process for producing high-levels of recombinant human cystatin-C. Enzyme Microbial Technol. 29, 335–340. 29. Zhang, W., Hywood Potter K. J., Plantz, B. A., Schlegel, V. L., Smith, L. A., and Meagher, M. M. (2003) Pichia pastoris fermentation with mixed-feeds of glycerol and methanol: growth kinetics and production improvement. J. Ind. Microbiol. Biotechnol. 30, 210–215. 30. Xie, J., Zhang, L., Ye, Q., Zhou, Q., Xin, L., Du, P., and Gan, R. (2003) Angiostatin production in cultivation of recombinant Pichia pastoris fed with mixed carbon sources. Biotechnol. Lett. 25, 173–177. 31. McGrew, J. T., Leiske, D., Dell, B., Klinke, R., Krasts, D., Wee, S. F., Abbott, N., Armitage, R., and Harrington, K. (1997) Expression of trimeric CD40 ligand in Pichia pastoris: use of a rapid method to detect high-level expressing transformants. Gene 187, 193–200. 32. d’Anjou, M. C., and Daugulis, A. J. (2001) A rational approach to improving productivity in

Pichia pastoris fermentation. Biotechnol. Bioeng 72, 1–11. 33. Cai, M., Huang, Y., Sakaguchi, K., Clore, G. M., Gronenborn, A. M., and Craigie, R. (1998) An efficient and cost-effective isotope labeling protocol for proteins expressed in Escherichia coli. J. Biomol. NMR 11, 97–102. 34. Zhang, Y., Liu, R., and Wu, X. (2007) The proteolytic systems and heterologous proteins degradation in the methylotrophic yeast Pichia pastoris. Annal. Microbiol. 57, 553–560. 35. Li, Z., Xiong, F., Lin, Q., d’Anjou, M., Daugulis, A. J., Yang, D. S., and Hew, C. L. (2001) Lowtemperature increases the yield of biological active herring antifreeze protein in Pichia pastoris. Protein Expr. Purif. 21, 438–445. 36. Jahic, M., Gustavsson, M., Jansen, A.-K., Martinelle, M., and Enfors, S.-O. (2003) Analysis and control of proteolysis of a fusion protein in Pichia pastoris fed-batch processes. J. Biotechnol. 102, 45–53. 37. Jahic, M., Wallberg, F., Bollok, M., Garcia, P., and Enfors, S.-O. (2003) Temperature limited fed-batch technique for control of proteolysis in Pichia pastoris bioreactor cultures. Microbial Cell Factories 2, 6–17. 38. Surribas, A., Stahn, R., Montesinos, J. L., Enfors, S.-O., Valero, F., and Jahic, M. (2007) Production of Rhizopus oryzae lipase from Pichia pastoris using alternative operational strategies. J. Biotechnol. 130, 291–299. 39. Shi, X., Karkut T., Chamankhah, M., AltingMees, M., Hemmingsen, S. M., and Hegedus, D. (2003) Optimal conditions for the expression of a single-chain antibody (scFv) gene in Pichia pastoris. Protein Expr. Purif. 28, 321–330. 40. Yao, X. Q., Zhao, H. L., Xue, C., Zhang, W., Xiong, X. H., Wang, Z. W., Li, X. Y., and Liu, Z. M. (2009) Degradation of HSAAX15(R13K) when expressed in Pichia pastoris can be reduced via the disruption of YPS1 gene in this yeast. J. Biotechnol. 139, 131–136. 41. Madduri, K., Badger, M., Li, Z.-S., Xu, X., Thornburgh, S., Evans, S., and Dhadialla, T. S. (2009) Development of stable isotope and selenomethionine labeling methods for proteins expressed in Pseudomonas fluorescens. Protein Expr. Purif. 65, 57–65.

Chapter 3 Isotope Labeling in Insect Cells Krishna Saxena, Arpana Dutta, Judith Klein-Seetharaman, and Harald Schwalbe Abstract Recent years have seen remarkable progress in applying nuclear magnetic resonance (NMR) spectroscopy to proteins that have traditionally been difficult to study due to issues with folding, posttranslational modification, and expression levels or combinations thereof. In particular, insect cells have proved useful in allowing large quantities of isotope-labeled, functional proteins to be obtained and purified to homogeneity, allowing study of their structures and dynamics by using NMR. Here, we provide protocols that have proven successful in such endeavors. Key words: Isotope labeling, Baculovirus, Nuclear magnetic resonance, Recombinant protein expression, Insect cells

1. Introduction: Baculovirus-Insect Cell Expression System

Isotope labeling of proteins represents an important and often required tool for the application of nuclear magnetic resonance (NMR) spectroscopy to investigate the structure and dynamics of proteins. So far, the great majority of isotope-labeled proteins have been expressed in Escherichia coli (1) because of the ease of cloning and expressing proteins at low cost. When possible, protein production should be performed in a prokaryotic system (i.e., E. coli), since this strategy is the most cost-effective and allows the most flexibility. However, human or complex proteins often cannot be expressed in E. coli in an active, correctly folded, posttranslationally modified form (glycosylated, phosphorylated, etc.). The capacity of E. coli for protein folding and forming disulfide bonds is not

Alexander Shekhtman and David S. Burz (eds.), Protein NMR Techniques, Methods in Molecular Biology, vol. 831, DOI 10.1007/978-1-61779-480-3_3, © Springer Science+Business Media, LLC 2012

37

38

K. Saxena et al.

sufficient for many recombinant proteins, although there are a number of new developments to overcome these limitations: 1. Decreasing the temperature of the cell culture (2, 3) 2. Coexpressing molecular chaperones (4) 3. Fusing highly soluble tags (gst, mbp, trxa, nusa, sumo, etc.) to the target proteins (5) 4. Overexpressing (6, 7) the target protein or using an engineered E. coli strain capable of forming disulfide bonds in the cytoplasm (e.g., Shuffle, New England Biolabs) 5. Refolding in vitro However, low-expression yield and solubility problems of the target protein in E. coli often force the change from a bacterial recombinant protein expression system to a eukaryotic host. Numerous eukaryotic-based expression systems are currently available in protein expression laboratories. For NMR use, only three expression hosts are generally considered: yeast (Pichia pastoris (8), Hansenula polymorpha or Kluyveromyces lactis), baculovirusmediated insect cells, and mammalian cells. More recently, the generation of baculoviruses harboring mammalian promoters (BacMams) have extended the use of baculovirus-mediated expression systems (BvE) to the development of gene delivery (9) and expression vectors in mammalian cells (10). BacMams cannot replicate in mammalian cells, which renders them a much safer alternative to conventional virus vectors (11). Yeast offers a powerful, simple system for expressing recombinant proteins. Besides the capability of performing posttranslational modification to the recombinant protein, the main advantage of these expression hosts is the feasibility of isotope labeling in simple minimal-defined media. Therefore, the costs of 15N, 13C, or 2 H uniform isotope incorporation are negligible in comparison to the costs of other eukaryotic cell media. Higher eukaryotes need well-defined expression media supplemented with expensive, labeled amino acids. But there are also disadvantages to recombinant protein expression in yeast. Since they provide N- and O-linked high-mannose-type glycans that could be immunogenic in humans (12, 13), the production of glycosylated proteins in yeast is not optimal, especially if glycosylation is required for the biological functionality of the target protein. Additionally, yeast cannot perform tyrosine O-sulfation (14), and proteins whose native forms are nonglycosylated may be hyperglycosylated when expressed in yeast (15). A recent study reported the successful reengineering of the glycosylation pathways in P. pastoris to allow the expression of recombinant proteins with human-type glycans (16). This may allow future improvements in the expression of glycosylated proteins in yeast. However, glycosylation is not the only challenge for successful expression of proteins in yeast. Certain proteins are

3

Isotope Labeling in Insect Cells

39

expressed at low levels even when glycosylation is not necessarily an issue (17–19). While the reasons are not fully clear, low-expression yields have been attributed to defects in the ER folding machinery (20–22). Even for proteins in which high-expression yields can be obtained, much of the protein may be misfolded (23). In order to detect which part of the folding machinery may be responsible, protein disulfide isomerase (PDI), an enzyme that catalyzes disulfide exchange in the ER, was overexpressed and found to enhance protein yields (24, 25). On the other hand, human adenosine A2A receptor levels were not found to increase with an increase in PDI expression (26). Thus, the process of disulfide bond formation in yeast remains uncertain. Finally, the case is most uncertain for membrane proteins, where there is a lack of knowledge on folding mechanisms and also on the way chaperones and the translocon participate in the folding pathway. Therefore, it is currently recommended to use higher eukaryotic hosts with advanced cell machinery systems for the production of recombinant proteins that have to be glycosylated, disulfide bonded, and/or membrane inserted for functional activity. Advanced expression systems with higher cell machineries for posttranslational modifications are offered by the baculovirusmediated expression system in insect cells (BvE) and mammalian expression methods. The BvE is one of the most efficient and popular systems among the eukaryotic hosts to use for expressing recombinant proteins. Therefore, its application is widespread in industrial as well as in academic environments for structural and functional studies of diverse proteins. However, for the production of therapeutic recombinant proteins, mammalian expression systems (mainly Chinese hamster ovary cells (CHO) and Human embryonic kidney (HEK) 293 cells) are required. Since CHO and HEK293 cells have been extensively characterized and have the ability for human-like glycosylation, the production of recombinant therapeutic monoclonal antibodies and Fc fusion proteins in this host is safe. The rate of production of therapeutic proteins, the largest class of new products being developed by the biopharmaceutical industry, has increased significantly in recent years (27). Mammalian expression systems have conventionally been considered to be too weak and inefficient for protein expression. However, recent advances have significantly improved the expression levels of these systems. This chapter and the following one, therefore, attempt to provide an overview of some of the recent developments in expression strategies for baculovirus-mediated insect cell and mammalian expression systems in view of NMR investigations. Since NMR requires isotope-labeled protein, the focus of this article is directed toward strategies to incorporate stable isotopes (15N, 13C) into the target protein in insect and mammalian cells (see Chapter 4). A major bottleneck of uniform labeling in higher eukaryotes is the high cost of complex medium

40

K. Saxena et al.

with labeled amino acids. Another limitation of these hosts is that they cannot survive in deuterium oxide (D2O)-containing media, so cost-effective generation of perdeuterated proteins is not available, either for insect or for mammalian cell systems. It is beyond the scope of both chapters to discuss features of the BvE or mammalian expression systems in detail. Comprehensive guides and detailed methodologies for the construction and analysis of recombinant baculovirus for insect cell expression, maintenance of insect cells in culture, and analysis of recombinant protein expression can be found elsewhere (28, 29), including in Baculovirus Manuals from Invitrogen, Pharmingen, Novagen, and others. However, given the number of different BvE strategies, we begin this chapter by explaining the principle of BvE and subsequently give a short survey of BvE options. 1.1. Principle of Baculovirus-Mediated Recombinant Protein Expression

The insect cell baculovirus-mediated expression system (BvE) is a powerful platform to rapidly produce high levels of recombinant proteins (see ref. 28 for an excellent review). Unlike bacterial expression hosts, the baculovirus system relies on a eukaryotic expression system and thus offers protein modification, sorting, and transportation machineries similar to those found in higher eukaryotic organisms. Baculoviruses are insect viruses that predominantly infect insect larvae of the order Lepidoptera (butterflies and moths) (30). A baculovirus expression vector is a recombinant baculovirus that has been genetically modified to contain a foreign gene of interest, which can be expressed in insect cells under the control of a baculovirus gene promoter. The BvE uses a helper-independent virus that can be propagated to high titers in insect cells adapted for growth in suspension cultures, enabling the production of large amounts of protein with relative ease (31). Finally, baculoviruses are noninfectious to vertebrates and their promoters have been shown to be inactive in mammalian cells (32). The most commonly used baculovirus for recombinant protein expression is Autographa californica, a multicapsid nucleopolyhedrovirus (AcMNPV) (33). AcMNPV is a large (130 kb), lytic, double-stranded DNA virus and can accommodate large segments of foreign DNA for the expression of recombinant protein (34). The BvE is based on the infection of cultured insect cells by a recombinant virus vector in which the target DNA (or multiple genes) is integrated under the control of the strong viral polyhedron promoter (28). The polyhedrin gene (polh) is necessary for the formation of polyhedral or occlusion bodies in the cell nucleus, but is nonessential for viral replication in insect cells. Polyhedra are large particles that appear in the nuclei of AcMNPV-infected insect cells. The first recombinant baculoviruses were generated by replacing the viral polyhedrin gene with a foreign gene of interest through homologous recombination (33). Homologous exchange between the flanking sequences common to both DNA molecules facilitates

3

Isotope Labeling in Insect Cells

41

the insertion of the gene of interest into the viral genome at the polh locus, resulting in the production of a recombinant virus genome and allowing the powerful polyhedron promoter to drive protein expression of the foreign gene. Since the efficiency of homologous recombination is quite low, identification, isolation, and selection of recombinant virus were traditionally achieved by labor-intensive, technically demanding plaque assays. Due to the deletion of the polyhedrin gene, the recombinant plaque has a more clearly distinct morphology than the parental virus containing the polh gene. Subsequently, additional rounds of plaque screening are required to separate the desired recombinant virus from the parental wild-type virus. However, discriminating between polyhedron-positive and -negative plaques and isolating recombinant virus turned out to be a serious problem for many investigators who used the BvE for the production of recombinant proteins. Nowadays, these technical issues and the time-consuming plaque purification processes are eliminated. In the next section, some of the key developments in the BvE are presented and discussed. 1.2. Commercially Available Baculovirus Expression Systems

Generally, the baculovirus genome is considered too large to insert a foreign gene directly. In most applications of the BvE, the gene of interest is therefore cloned into a transfer vector, which contains sequences that flank the polyhedron gene in the baculovirus genome. The virus genome and the transfer vector are cotransfected into the insect cells and the gene of interest inserts into the virus genome via homologous recombination (see Fig. 1) under the control of the strong late viral polyhedrin promoter (35). Since a mixture of recombinant and original parental virus is produced after the initial replication, time-consuming plaque purification and isolation are required before protein expression can proceed. BacVector (Merck Biosciences), Baculo-Gold and pBacPAK (BD Biosciences), and Bac-N-Blue (Invitrogen) are commercially available BvEs that use homologous recombination to integrate foreign genes into the virus genome. New developments in generating recombinant virus by using site-specific transpositions (Bac-to-Bac or BaculoDirect, Invitrogen) or progress in recombination methodology with an engineered baculovirus containing a lethal mutation in an essential gene (open reading frame (ORF1629), flashBAC from Oxford Expression Technologies, or BacMagic from EMD Chemicals, Novagen) have facilitated the use of BvE for a larger user community (28). Generally, these improvement strategies can be classified into transfer plasmid modifications and parental baculovirus genome modification (28). The flashBAC and BacMagic are the most promising BvEs so far, since the efficiency of recombination in both systems is 100%. Therefore, these BvEs overcome the requirement of timeconsuming plaque assays and protein expression can be started directly after one or two rounds of virus amplification. This technology

42

K. Saxena et al.

Fig. 1. Construction of baculovirus recombinants with Novagen® BacMagic™ system. This expression system is based on a modified baculovirus genome containing a bacterial artificial chromosome (BAC) at the polyhedrin locus and a partial deletion of the essential ORF1629 viral gene. The BacMagic DNA is mixed with a transfer vector, containing a foreign gene at the polh locus and the complete ORF1629, to generate the recombinant virus via homologous recombination in insect cells. Picture modified from the Novagen manual. © EMD Chemicals Inc., an Affiliate of Merck KGaA. Darmstadt, Germany. BacMagic™ and Novagen® are trademarks of Merck KGaA.

reduces the production of recombinant virus to a one-step procedure, fully amenable to high-throughput and automated production systems. Moreover, this approach is compatible with all baculovirus transfer vectors based on homologous recombination in insect cells at the polyhedrin locus, including several multigene coexpression plasmids. The technology of the flashBAC and BacMagic is driven by a modified bacmid, in which the baculovirus genome AcMNPV with a portion of the essential viral gene (ORF1629) deleted. In addition, a bacterial artificial chromosome (BAC) replaces the polyhedrin-coding region. This combination prevents nonrecombinant virus from replicating in insect cells, yet allows the viral DNA to be propagated as circular DNA in bacteria. This circular viral DNA is then isolated and purified from bacterial cells (flashBAC or BacMagic DNA provided in the kits). Homologous recombination with a compatible expression plasmid (containing the gene of interest

3

Isotope Labeling in Insect Cells

43

flanked by the lef2 and ORF1629 recombination sites) restores the function of the viral ORF1629 allowing the virus DNA to replicate and replaces the BAC sequence with the target coding sequence under the control of the polyhedrin promoter (Fig. 1). Since only recombinant viruses with a restored ORF1629 can replicate, this results in a unique recombinant virus population. This population can then be used directly to infect a larger insect cell culture (50–200 mL) to produce a high-titer working stock. 1.3. Insect Cell Lines

The main insect cell lines used for cotransfections and baculovirus amplification are Spodoptera frugiperda Sf9 or Sf21 (derivatives of the fall armyworm). Trichoplusia ni BTI 5B1-4 (36) (High Five™) cells are generally used for the production of secreted recombinant proteins and not for virus production because of the increased possibility of generating virus mutants (37). Due to the high-mannose and paucimannose types of glycosylation that are obtained in insect cells, no therapeutic protein is currently produced using this system as this would compromise in vivo bioactivity and potentially induce allergenic reactions. Engineering insect cells with glycosyltransferases allows the production of proteins with mammaliantype sugars (38).

1.4. Expression of Labeled Recombinant Protein in the Baculovirus Expression System

Due to the high costs of incorporating stable isotopes into insect cells, it is recommended that the recombinant protein expression is optimized using the chosen BvE before starting with labeled fermentation. There are only a few studies reporting the incorporation of stable isotopes into proteins expressed by baculovirusmediated insect cell expression (39–47). In contrast to the initial trials of amino acid-type selective labeling of proteins in user-defined insect cell media, nowadays there are commercial media (BioExpress-2000, Cambridge Isotope Laboratories, CIL) available for the different labeling applications. Expression of uniformly 13C- 15N-labeled Abelson Kinase domain (13C–15N BioExpress-2000) was the first example of backbone NMR resonance assignments of a recombinant protein expressed using the BvE (48). So far, most uniform labeling protein work in insect cells is only performed by industrial research groups due to the extraordinary costs of the required media. It is not possible to cultivate insect cells in minimal medium, since this host requires essential amino acids for its growth. Reports of selective amino acid isotope labeling in BvE are more frequently cited in the literature, since this approach is easy and fast and does not require expensive medium for labeling. Even in the absence of a backbone assignment of the target protein, structural information can be deduced from the selective labeling approach based on an existing X-ray structure of the protein. From a practical aspect, it should be considered that there are essential and nonessential amino acids (alanine, cysteine, glutamic acid, glutamine, aspartic acid, and asparagine) in insect cells whose content in the medium

44

K. Saxena et al. NH4

Glutamin

Lysine

Aspartate

Asparagine

Glutamate

Tyrosine

Alanine

Phenylalanine

Valine

Leucine

Isoleucine

Glycine

Serine

Fig. 2. A schematic presentation of amino acid metabolism in E. coli and Sf9 with respect to 15N: The black arrows symbolize pathways present in both expression hosts. Pathways that only exist in E. coli are shown in grey. The strength of the arrows reflects the intensity of the conversion. Picture modified from Bruggert et al. (42).

depends on the specific provider of the medium. Before starting with site-specific amino acid labeling, the unlabeled quantity of the desired amino acid in the medium should be checked to calculate the required amount of the amino acid to be labeled. In Fig. 2, a schematic presentation of amino acid metabolism is shown for E. coli and insect cells (42). Interestingly, selective labeling of amino acids in insect cells is more effective than in bacteria, since the amino acid pathways in insect cells do not harbor as many aminotransferases as in prokaryotes, which leads to cross-labeling problems.

2. Materials 2.1. Cotransfection of Insect Cells

1. 35-mm2 tissue culture dishes. 2. Sf9 or Sf21 insect cells. 3. SF900 II insect cell culture medium (serum-free, antibioticfree; Invitrogen). 4. Insect GeneJuice® transfection reagent (Novagen) (see Note 1). 5. BacMagic DNA: 100 ng (5 μL) per cotransfection (20 ng/μL). 6. Sterile baculovirus transfer vector DNA containing the gene under investigation (500 ng per cotransfection). 7. Plastic box to house dishes in the incubator. 8. Sterile pipettes, bijoux (sterile tubes).

3

2.2. Amplification of Recombinant Virus

Isotope Labeling in Insect Cells

45

1. Recombinant virus seed stock (Subheading 3.1). 2. Sf 9 insect cells. 3. SF900 II insect cell culture medium (Subheading 2.1). 4. Inverted phase-contrast microscope.

2.3. Analysis of Recombinant Protein Expression

1. 35-mm2 tissue culture dishes. 2. Sf 9 or High Five insect cells. 3. SF900 II insect cell culture medium (Subheading 2.1). 4. Recombinant virus stock (Subheading 3.2). 5. Phosphate-buffered saline (PBS; Invitrogen): pH 6.2, sterilize by autoclaving.

2.4. Production of Isotopically Labeled Protein

1. Sf 9 insect cells. 2. SF900 II insect cell culture medium (Subheading 2.1). 3. BioExpress-2000-U (CIL): Unlabeled insect cell culture medium. 4. BioExpress-2000-CN (CIL): 15N-, 13C-labeled insect cell culture medium. 5. Recombinant virus stock (Subheading 3.2). 6. Protease inhibitor (Complete™, Roche). 7. PBS (Subheading 2.3).

3. Methods For a general overview of the BvEs and cloning, expression, analysis, and purification of recombinant proteins in insect cells, please refer to ref. 29. The following procedures describe the cloning of a recombinant baculovirus expression vector, production (Subheading 3.1) and amplification of recombinant baculovirus (Subheading 3.2), analysis of recombinant protein expression (Subheading 3.3), and finally production of isotope-labeled protein (Subheading 3.4) expressed in insect cells. All of the procedures are based on the flashBAC or BacMagic systems (no plaque purification required) and must be carried out using sterile technique. Since the flashBAC system is compatible with all transfer vectors designed for homologous recombination in insect cells at the polh l (BacPAK technology), the target gene can be cloned into many suitable transfer vectors (see Note 2). Moreover, this BvE is compatible with traditional (T4 DNA ligase) and elegant recombinatorial cloning techniques, such as Creator or In-Fusion (BD Biosciences) (see Note 3). While Gateway (Invitrogen) is one of the most popular recombinatorial cloning systems, it is limited to

46

K. Saxena et al.

specifically engineered expression vectors with specifically engineered expression vectors with λ recombination sites. These lead to incorporation of additional amino acids into the protein of interest. A new development to avoid multiple cloning of target genes into host-specific expression vectors is triple host transfer vector, pTriEx, from Novagen. Due to the parallel existence of three promoters in this vector series, recombinant protein expression is enabled in vertebrates, insect cells, and bacteria. 3.1. Cotransfection of Insect Cells

For efficient transfection, high-quality DNA of the transfer plasmid is prepared using commercially available plasmid DNA purification kits (Qiagen, Novagen) (see Note 4). Additionally, it is recommended to use fresh, rapidly proliferating cells (see Note 5) for transfection experiments and to have positive and negative transfection controls (see Note 6). 1. For each cotransfection, prepare one 35-mm2 plate. Seed the dishes with insect cells at least 1 h before use. Use 1 × 106 cells/ dish for Sf9 cells and 1.5 × 106 cells/dish for Sf21 cells in 2 mL of SF900 II insect cell culture medium. Allow the cells to attach by incubating at 28°C for 20 min. 2. During the 1-h incubation period, prepare a DNA–liposome complex cotransfection mix of BacMagic DNA and Insect GeneJuice® transfection reagent for each transfection. Assemble the following components, in the order listed, in a sterile tube (bijoux) (see Note 7): 1 mL serum-free, antibiotic-free SF900 II insect cell culture medium 5 μL Insect GeneJuice 5 μL BacMagic DNA (100 ng total) 5 μL Transfer vector DNA (500 ng total) 1.015 mL Total volume 3. Incubate at room temperature for 15–30 min to allow the DNA–liposome complexes to form. 4. Remove the culture medium from the 35-mm2 dishes of cells using a sterile pipette, ensuring that the cell monolayer is not disrupted (see Note 8). 5. Immediately after the medium has been removed from the cells, add the 1 mL of the DNA–liposome complex dropwise to the center of each dish (see Note 9) and incubate in a plastic sandwich box at 28°C for a minimum of 5 h or overnight. 6. After the incubation period, add another 1 mL of SF900 II insect cell culture medium to each dish and continue the incubation for 5 days in total.

3

Isotope Labeling in Insect Cells

47

7. Following the 5-day incubation period (see Note 10), harvest the medium containing the recombinant virus into a sterile bijoux and store in the dark at 4°C. This is the seed stock of recombinant baculovirus (see Note 11). Due to the limited size of the stock, the next step is to amplify the virus recombination sites. 3.2. Amplification of Recombinant Virus

Amplification of the recombinant virus (produced in Subheading 3.1) is necessary before proceeding with recombinant protein expression. The following provides a protocol (adapted from the Novagen and flashBAC Manual) for amplification and preparation of high-titer recombinant virus (passage 1 stock) in cells grown in suspension culture. 1. Observe the health and viability of cells under an inverted phase-contrast microscope (see Note 12). 2. Prepare a 100–200 mL culture of Sf9 cells at an appropriate cell density in serum-free SF900 II insect cell culture medium (e.g., 2 × 106 Sf9 cells/mL in log-phase growth; high aeration is recommended (see Note 13)). Cells should be infected at a multiplicity of infection (MOI) of <1 pfu/cell. 3. Add 0.5 mL of recombinant virus seed stock to the cell culture. Incubate with shaking until the cells are well-infected (usually, 4–5 days) (see Note 14). 4. When the cells appear to be well-infected with virus, harvest the cell culture medium by centrifugation at 1,500 × g for 15 min at 4°C. Remove the supernatant aseptically. Store the supernatant (recombinant virus stock) in the dark at 4°C (see Note 15).

3.3. Analysis of Recombinant Protein Expression

Before proceeding with use of the virus in subsequent protein expression experiments, a plaque assay to determine the accurate titer is strongly recommended by the manufacturer of the flashBAC and BacMagic technology (see Note 16). This allows the calculation of MOI and ensures cross-referencing and reproducibility in the following experiments. However, for analysis of protein expression, it is not absolutely required to know the titer of the recombinant virus (see Note 17). Pilot expression analysis can also be performed by infecting cells with different amounts of P1 virus and monitoring the expression of the recombinant protein. According to the protein expression literature, it is recommended that exponentially growing cells should be infected at a high MOI to ensure that all cells are infected simultaneously and that the culture is synchronous. However, in our lab, various P1 viruses are directly added (0.5–10%) to the insect cells for determining the best ratio for optimal recombinant protein expression without knowing the exact titer of the different viruses. The best time to

48

K. Saxena et al.

harvest the recombinant protein can be examined by taking samples at different time points after infection (see Note 18). It is recommended to also prepare noninfected or mock-infected cells as a control for host cell proteins and a positive control of a successfully tested recombinant virus. Recombinant protein expression can then be evaluated by SDS-PAGE or Western blot analysis. Additionally, protein expression should be compared between Sf9 and High Five cell lines, especially if the protein of interest is secreted (see Note 19). The following protocol is adapted from the Novagen and flashBAC Manual. 1. Seed 35-mm2 dishes (one per virus to be tested plus a positive and negative control) with 1 × 106 of S9 or High Five cells per dish and allow the cells to attach by incubating at 28°C for 1 h. 2. Remove the medium, add 200 μL of the recombinant virus stock to be tested, and incubate at room temperature for an additional hour. 3. Remove the virus inoculum, add 1.5 mL of SF900 II insect cell culture medium, and incubate at 28°C for 48 h. 4. Use a sterile pipette tip to scrape the cells from the dishes into suspension and to transfer them into sterile Eppendorf tubes. 5. Centrifuge the cells at 1,500 × g for 10 min at room temperature. Remove the supernatant and discard (see Note 20). 6. Resuspend the pellet (protein of interest in the cytosol) in 80 μL of PBS. 7. Analyze the extent of protein overexpression by using SDSPAGE and Western blotting if a suitable antibody is available for the recombinant protein (see Note 21 and Fig. 3).

Fig. 3. Protein overexpression and purification using insect cells. SDS PAGE (left ), Western Blot (right ): Protein samples loaded are BC (before Ni-NTA chromatography), FT (Flowthrough), M (Molecular weight marker), W (Wash), E (Elution).

3

3.4. Production of Isotopically Labeled Protein

Isotope Labeling in Insect Cells

49

While fermenters and bioreactors are now routinely used to produce recombinant proteins in insect cells, good yields can be achieved in shake cultures at a fraction of the cost. Large, disposable shake flasks can be obtained that hold up to 1.5 L of insect cell culture on a shaking platform in a warm room or incubator. The most suitable culture protocol for isotope labeling of a recombinant protein in insect cells involves an initial growth phase for the insect cells in an unlabeled growth/expression medium and a subsequent expression phase, where cells are centrifuged and resuspended into the labeling medium prior to infection with the recombinant baculovirus (see Note 22). This centrifugation step allows much higher expression levels of the labeled protein compared to one with growth and expression in labeling medium without the change in medium. Recently, isotope-labeled expression media for insect cells have become commercially available (BioExpress-2000, CIL), but the source and the concentration of the amino acids in these media are not published. However, for site-specific labeling of amino acids in proteins expressed in insect cells, adding the isotopelabeled amino acid of interest to a standard insect cell medium before infection (see Note 23) also works well and the cost for this labeling strategy is comparable to that when using E. coli. The following protocol is adapted from the Application Note: Efficient uniform isotope labeling of proteins expressed in Baculovirus-infected insect cells using BioExpress® 2000 (Insect cell) medium by André Strauss, Gabriele Fendrich, and Wolfgang Jahnke. Prior to performing isotope labeling of a protein, optimize the culture and BV-infection conditions in unlabeled medium (e.g., BioExpress 2000-U or SF900 II for expression of the protein). 1. Cultivate several 100 mL cultures of Sf9 cells, adapted to growth in serum-free SF900 II medium, in 500-mL Erlenmeyer flasks for 3 days at 27°C with shaking at 90 rpm. 2. Prepare the uniform isotope-labeling medium (see Note 24). 3. When the final cell density of the culture has reached 1.5 × 106 cells/mL (~3 days), sterile centrifuge the cells at 400 × g for 20 min at 20°C. 4. Resuspend the pelleted cells in 100-mL portions in labeled BioExpress® 2000-CN medium and transfer them to fresh 500-mL Erlenmeyer flasks. 5. Add the recombinant virus stock at a titer of 0.5–2 × 108 pfu/mL to an MOI = 1–2, according to optimized conditions. 6. Grow the 100 mL cultures of baculovirus-infected Sf9 cells for 3 days post infection in labeled BioExpress® 2000-CN medium at 27°C, with shaking at 90 rpm (see Note 25). 7. Harvest the cells expressing the labeled recombinant protein by centrifuging at 400 × g for 20 min at 20°C (see Note 26); resuspend the pelleted cells in 20 mL of PBS containing

50

K. Saxena et al.

protease inhibitor mix (see Note 27), followed by a second centrifugation in 50-mL plastic tubes at 400 × g for 20 min at 20°C. Store the pelleted cells at −80°C. 8. Isolate and purify the recombinant protein according to protocols generated for the unlabeled protein.

4. Notes 1. Lipofectin® (Invitrogen), FuGENE 6 (Roche), Tfx-20™ (Promega), and CELLFECTIN® (Invitrogen) have also been successfully tested. 2. Several suitable vectors from different providers are available and are summarized: http://www.expressiontechnologies. com/flashBAC/vectors.asp. 3. Ligation protocols are described in the manuals of the specific cloning technology providers. 4. Plasmid DNA purification protocols are provided by the manufacturers of the purification kits. The DNA must be sterile and must be of a quality suitable for transfection into cells. 5. Healthy cells look bright, round, and refractile, and many should be in the process of dividing into daughter cells. 6. A control transfer vector is supplied with the BacMagic kit and can be used to make recombinant virus; the lacZ-positive infected cells can be stained using X-gal. Add 1 mL of appropriate insect cell culture medium containing 15 μL of 2% (w/v) X-gal in N,N-dimethylformamide and incubate at 28°C. After 5 h, the cells and culture medium appear blue in color, confirming the production of recombinant virus. 7. Plasticware used to prepare the transfection mixture has to be made from polystyrene and not from polypropylene, since the complexes bind to polypropylene. 8. When removing liquid from a dish of cells, tip the dish at a 30–60° angle so that the liquid pools toward one side of the dish. It is important not to allow the cell monolayer to dry out at this point. 9. Adding the mixture dropwise should not disturb the cell monolayer if it is done slowly and gently. 10. Cell monolayers in which recombinant virus has been produced appear very different from mock-transfected control cells under the inverted microscope. Control cells form a confluent monolayer while virus-infected cells do not form a confluent monolayer and appear grainy with enlarged nuclei.

3

Isotope Labeling in Insect Cells

51

11. The expected titer of this initial viral seed stock is generally about 1 × 107 pfu/mL. In contrast to virus-free cells, virusinfected cells appear grainy with enlarged nuclei and do not form a confluent monolayer. 12. It is important that the cells are healthy and in the log phase of growth to ensure that virus replication occurs efficiently to generate high-titer stocks of virus for subsequent use in expression studies suitable for transfection into cells. 13. Virus-infected cells have an increased need for oxygen, and therefore the contents of the flasks should be shaken at quite high speeds to maximize aeration. The surface area-to-volume ratio should also be as large as possible for maximum gas exchange. Do not overfill the flasks. 14. Under a phase-contrast inverted microscope, cells infected with virus appear grainy when compared to healthy cells. The infected cells become uniformly rounded and enlarged, with distinct enlarged nuclei. 15. The virus stock can be stored in the dark at 4°C for 6–12 months, although the titer begins to drop after 3–4 months. Titer the virus before use and reamplify if necessary. The addition of 2–5% serum when using serum-free medium can be helpful in avoiding a drop in titer. Virus may be frozen at −80°C for longer periods of time. Avoid multiple freeze–thaw cycles. 16. Protocols for plaque assays can be found in the manuals of Novagen and Oxford Expression Technologies. 17. After cotransfecting and harvesting the seed stock of virus for further amplification, it is possible to harvest the remaining cells from the dish and prepare these for SDS-PAGE/Western blotting. This gives a quick check for gene expression. 18. The most commonly used times are at 24, 48, 72, and 96 h post infection (hpi). Some proteins may be very stable and accumulate to high levels by 96 hpi, and others may start to degrade and thus need to be harvested much earlier. 19. High Five cell lines (Invitrogen) often increase the yield of secreted recombinant proteins. 20. For secreted protein expression, the supernatant has to be analyzed. 21. Standard molecular biology procedures like SDS-PAGE and Western-blotting are precisely described in Molecular Cloning: A Laboratory Manual) (49). 22. A similar cost-reducing labeling strategy in E. coli was also described for the production of isotope-labeled protein by Marley and coworkers (50).

52

K. Saxena et al.

23. 300–600 mg of 15N-Phe or 15N-Leu (per liter culture medium) was immediately added to the insect cells after being infected with virus. The medium used for the amino acid-selective labeling was 1LSF-900 II serum-free medium from Invitrogen. 24. Medium can be stored, and filter-sterilized for several months at 4°C in the dark without loss of capacity for protein expression. Warm to 28°C before using. 25. The harvesting time of the infected cells depends on the stability of the target protein. 26. For secreted protein expression, the supernatant has to be further purified. 27. Protease inhibitor mix: Roche complete (one tablet dissolved in 200 mL of PBS) References 1. Goto, N. K., and Kay, L. E. (2000) New developments in isotope labeling strategies for protein solution NMR spectroscopy. Curr. Opin. Struct. Biol. 10, 585–592. 2. Schein, C. H. (1991) Optimizing protein folding to the native state in bacteria. Curr Opin. Biotechnol. 2, 746–750. 3. Qing, G., Ma, L. C., Khorchid, A., Swapna, G. V., Mal, T. K., Takayama, M. M., Xia, B., Phadtare, S., Ke, H., Acton, T., Montelione, G. T., Ikura, M., and Inouye, M. (2004) Coldshock induced high-yield protein production in Escherichia coli. Nat. Biotechnol. 22, 877–882. 4. Young, J. C., Agashe, V. R., Siegers, K., and Hartl, F. U. (2004) Pathways of chaperonemediated protein folding in the cytosol. Nat. Rev. Mol. Cell. Biol. 5, 781–791. 5. Esposito, D., and Chatterjee, D. K. (2006) Enhancement of soluble protein expression through the use of fusion tags. Curr. Opin. Biotechnol. 17, 353–358. 6. Andersen, C. L., Matthey-Dupraz, A., Missiakas, D., and Raina, S. (1997) A new Escherichia coli gene, dsbG, encodes a periplasmic protein involved in disulphide bond formation, required for recycling DsbA/DsbB and DsbC redox proteins. Mol. Microbiol. 26, 121–132. 7. Bardwell, J. C. (1994) Building bridges: disulphide bond formation in the cell. Mol. Microbiol. 14, 199–205. 8. Pickford, A. R., and O’Leary, J. M. (2004) Isotopic labeling of recombinant proteins from the methylotrophic yeast Pichia pastoris. Methods Mol. Biol. 278, 17–33.

9. Hu, Y. C. (2008) Baculoviral vectors for gene delivery: a review. Curr. Gene Ther. 8, 54–65. 10. Kost, T. A., Condreay, J. P., Ames, R. S., Rees, S., and Romanos, M. A. (2007) Implementation of BacMam virus gene delivery technology in a drug discovery setting. Drug Discov. Today 12, 396–403. 11. Hitchman, R. B., Possee, R. D., and King, L. A. (2009) Baculovirus expression systems for recombinant protein production in insect cells. Recent Pat. Biotechnol. 3, 46–54. 12. Lam, J. S., Huang, H., and Levitz, S. M. (2007) Effect of differential N-linked and O-linked mannosylation on recognition of fungal antigens by dendritic cells. PLoS One 2, e1009. 13. Dasgupta, S., Navarrete, A. M., Bayry, J., Delignat, S., Wootla, B., Andre, S., Christophe, O., Nascimbeni, M., Jacquemin, M., MartinezPomares, L., Geijtenbeek, T. B., Moris, A., Saint-Remy, J. M., Kazatchkine, M. D., Kaveri, S. V., and Lacroix-Desmazes, S. (2007) A role for exposed mannosylations in presentation of human therapeutic self-proteins to CD4+ T lymphocytes. Proc. Natl. Acad. Sci. U.S.A 104, 8965–8970. 14. Moore, K. L. (2003) The biology and enzymology of protein tyrosine O-sulfation. J. Biol. Chem. 278, 24243–24246. 15. Daly, R., and Hearn, M. T. (2005) Expression of heterologous proteins in Pichia pastoris: a useful experimental tool in protein engineering and production. J. Mol. Recognit. 18, 119–138. 16. Hamilton, S. R., and Gerngross, T. U. (2007) Glycosylation engineering in yeast: the advent

3

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

27.

28.

of fully humanized yeast. Curr. Opin. Biotechnol. 18, 387–392. Chisholm, V., Chen, C. Y., Simpson, N. J., and Hitzeman, R. A. (1990) Molecular and genetic approach to enhancing protein secretion. Methods Enzymol. 185, 471–482. Dorner, A. J., and Kaufman, R. J. (1990) Analysis of synthesis, processing, and secretion of proteins expressed in mammalian cells. Methods Enzymol. 185, 577–596. Moir, D. T. (1989) Yeast mutants with increased secretion efficiency. Biotechnology 13, 215–231. Biemans, R., Thines, D., Rutgers, T., De Wilde, M., and Cabezon, T. (1991) The large surface protein of hepatitis B virus is retained in the yeast endoplasmic reticulum and provokes its unique enlargement. DNA Cell. Biol. 10, 191–200. Gennaro, D. E., Hoffstein, S. T., Marks, G., Ramos, L., Oka, M. S., Reff, M. E., Hart, T. K., and Bugelski, P. J. (1991) Quantitative immunocytochemical staining for recombinant tissue-type plasminogen activator in transfected Chinese hamster ovary cells. Proc. Soc. Exp. Biol. Med. 198, 591–598. Shuster, J. R. (1991) Gene expression in yeast: protein secretion. Curr. Opin. Biotechnol. 2, 685–690. Mollaaghababa, R., Davidson, F. F., Kaiser, C., and Khorana, H. G. (1996) Structure and function in rhodopsin: expression of functional mammalian opsin in Saccharomyces cerevisiae. Proc. Natl. Acad. Sci. U.S.A 93, 11482–11486. Robinson, A. S., Hines, V., and Wittrup, K. D. (1994) Protein disulfide isomerase overexpression increases secretion of foreign proteins in Saccharomyces cerevisiae. Biotechnology (N Y) 12, 381–384. Shusta, E. V., Raines, R. T., Pluckthun, A., and Wittrup, K. D. (1998) Increasing the secretory capacity of Saccharomyces cerevisiae for production of single-chain antibody fragments. Nat. Biotechnol. 16, 773–777. Butz, J. A., Niebauer, R. T., and Robinson, A. S. (2003) Co-expression of molecular chaperones does not improve the heterologous expression of mammalian G-protein coupled receptor expression in yeast. Biotechnol. Bioeng. 84, 292–304. Durocher, Y., and Butler, M. (2009) Expression systems for therapeutic glycoprotein production. Curr. Opin. Biotechnol. 20, 700–707. Jarvis, D. L. (2009) Baculovirus-insect cell expression systems. Methods Enzymol. 463, 191–222.

Isotope Labeling in Insect Cells

53

29. O´Reilly, D. R., Miller, L., Luckow, V.A. (1992) Baculovirus Expression Vectors - a Laboratory Manual. WH Freeman, New York. 30. van Regenmortel, M. H., Mayo, M. A., Fauquet, C. M., and Maniloff, J. (2000) Virus nomenclature: consensus versus chaos. Arch. Virol. 145, 2227–2232. 31. Hunt, I. (2005) From gene to protein: a review of new and enabling technologies for multiparallel protein expression. Protein Expr. Purif. 40, 1–22. 32. Carbonell, L. F., Klowden, M. J., and Miller, L. K. (1985) Baculovirus-mediated expression of bacterial genes in dipteran and mammalian cells. J. Virol. 56, 153–160. 33. Smith, G. E., Summers, M. D., and Fraser, M. J. (1983) Production of human beta interferon in insect cells infected with a baculovirus expression vector. Molecular Cell. Biol. 3, 2156–2165. 34. Ayres, M. D., Howard, S. C., Kuzio, J., LopezFerber, M., and Possee, R. D. (1994) The complete DNA sequence of Autographa californica nuclear polyhedrosis virus. Virology 202, 586–605. 35. Kitts, P. A., and Possee, R. D. (1993) A method for producing recombinant baculovirus expression vectors at high frequency. Biotechniques 14, 810–817. 36. Kost, T. A., Condreay, J. P., and Jarvis, D. L. (2005) Baculovirus as versatile vectors for protein expression in insect and mammalian cells. Nat. Biotechnol. 23, 567–575. 37. Friesen, P. D., and Nissen, M. S. (1990) Gene organization and transcription of TED, a lepidopteran retrotransposon integrated within the baculovirus genome. Mol. Cell. Biol. 10, 3067–3077. 38. Harrison, R. L., and Jarvis, D. L. (2006) Protein N-glycosylation in the baculovirusinsect cell expression system and engineering of insect cells to produce “mammalianized” recombinant glycoproteins. Adv. Virus Res. 68, 159–191. 39. DeLange, F., Klaassen, C. H., Wallace-Williams, S. E., Bovee-Geurts, P. H., Liu, X. M., DeGrip, W. J., and Rothschild, K. J. (1998) Tyrosine structural changes detected during the photoactivation of rhodopsin. J. Biol. Chem. 273, 23735–23739. 40. Creemers, A. F., Klaassen, C. H., Bovee-Geurts, P. H., Kelle, R., Kragl, U., Raap, J., de Grip, W. J., Lugtenburg, J., and de Groot, H. J. (1999) Solid state 15N NMR evidence for a complex Schiff base counterion in the visual G-proteincoupled receptor rhodopsin. Biochemistry 38, 7195–7199.

54

K. Saxena et al.

41. Bellizzi, J. J., Widom, J., Kemp, C. W., and Clardy, J. (1999) Producing selenomethioninelabeled proteins with a baculovirus expression vector system. Structure 7, R263-267. 42. Bruggert, M., Rehm, T., Shanker, S., Georgescu, J., and Holak, T. A. (2003) A novel medium for expression of proteins selectively labeled with 15N-amino acids in Spodoptera frugiperda (Sf9) insect cells. J. Biomol. NMR 25, 335–348. 43. Strauss, A., Bitsch, F., Cutting, B., Fendrich, G., Graff, P., Liebetanz, J., Zurini, M., and Jahnke, W. (2003) Amino-acid-type selective isotope labeling of proteins expressed in Baculovirus-infected insect cells useful for NMR studies. J. Biomol. NMR 26, 367–372. 44. Strauss, A., Bitsch, F., Fendrich, G., Graff, P., Knecht, R., Meyhack, B., and Jahnke, W. (2005) Efficient uniform isotope labeling of Abl kinase expressed in Baculovirus-infected insect cells. J. Biomol. NMR 31, 343–349. 45. Betz, M., Vogtherr, M., Schieborr, U., Elshorst, B., Grimme, S., Pescatore, B., Langer, T., Saxena, K., and Schwalbe, H. (2008) Chemical Biology of Kinases Studied by NMR Spectroscopy, In Chemical Biology (Prof. Dr. Stuart L. Schreiber, P. D. T. M. K. P. D. G., Ed.), pp 852–890.

46. Stockman, B. J., Kothe, M., Kohls, D., Weibley, L., Connolly, B. J., Sheils, A. L., Cao, Q., Cheng, A. C., Yang, L., Kamath, A. V., Ding, Y. H., and Charlton, M. E. (2009) Identification of allosteric PIF-pocket ligands for PDK1 using NMR-based fragment screening and 1H-15N TROSY experiments. Chem. Biol. Drug Des. 73, 179–188. 47. Jahnke, W., Grotzfeld, R. M., Pelle, X., Strauss, A., Fendrich, G., Cowan-Jacob, S. W., Cotesta, S., Fabbro, D., Furet, P., Mestan, J., and Marzinzik, A. L. (2010) Binding or bending: distinction of allosteric Abl kinase agonists from antagonists by an NMR-based conformational assay. J. Am. Chem. Soc. 132, 7043–7048. 48. Vajpai, N., Strauss, A., Fendrich, G., CowanJacob, S. W., Manley, P. W., Jahnke, W., and Grzesiek, S. (2008) Backbone NMR resonance assignment of the Abelson kinase domain in complex with imatinib. Biomol. NMR Assign. 2, 41–42. 49. Maniatis, T. (1982) Molecular cloning : a laboratory manual / T. Maniatis, E.F. Fritsch, J. Sambrook, Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y. 50. Marley, J., Lu, M., and Bracken, C. (2001) A method for efficient isotopic labeling of recombinant proteins. J. Biomol. NMR 20, 71–75.

Chapter 4 Isotope Labeling in Mammalian Cells Arpana Dutta, Krishna Saxena, Harald Schwalbe, and Judith Klein-Seetharaman Abstract Isotope labeling of proteins represents an important and often required tool for the application of nuclear magnetic resonance (NMR) spectroscopy to investigate the structure and dynamics of proteins. Mammalian expression systems have conventionally been considered to be too weak and inefficient for protein expression. However, recent advances have significantly improved the expression levels of these systems. Here, we provide an overview of some of the recent developments in expression strategies for mammalian expression systems in view of NMR investigations. Key words: Isotope labeling, Nuclear magnetic resonance, Recombinant protein expression, Human embryonic kidney cells

1. Introduction: Mammalian Cell Expression System 1.1. Principle of Mammalian Cell-Mediated Protein Expression

Mammalian cell expression systems are being increasingly used to express proteins for structural biology. This is evident from the increase in the number of structures available in the Protein Data Bank (PDB) (1) of proteins purified from such sources. The main reason for switching to these higher eukaryotic expression systems is the ability to produce biologically active cell surface receptors and secreted glycoproteins. The functionality of these proteins is linked to the requirement for posttranslational modifications, such as glycosylation and disulfide bond formation, which can often only be satisfied by using mammalian systems and not other eukaryotic systems, such as yeast and insect cells. Protein expression in mammalian cells involves transfection with a plasmid carrying the gene of interest under the control of a mammalian promoter. The mammalian gene can be introduced

Alexander Shekhtman and David S. Burz (eds.), Protein NMR Techniques, Methods in Molecular Biology, vol. 831, DOI 10.1007/978-1-61779-480-3_4, © Springer Science+Business Media, LLC 2012

55

56

A. Dutta et al.

into cells either by way of transient transfection or by developing a stable cell line. The former can be achieved with both adherent and suspension cells using polyethyleneimine (PEI). The only advantage of this method over stable cell line preparation is that milligram quantities of protein can be prepared in a few days time compared to the months required by the latter method (2–4). However, there are several disadvantages associated with the transient transfection approach (1) Not every protein, in particular membrane proteins, give high yields when transiently transfected; (2) Very large quantities of DNA need to be prepared; (3) The process needs to be repeated every time protein is needed, adding labor and cost. Stable cell line creation overcomes these limitations, although 2–6 months are required to establish a high-level expression system. The effort is often justified, however, by the advantages of a stable mammalian cell line over a transient transfect (1) The yield is typically very high; (2) The process becomes fast and robust once the cell line is created for a particular protein; (3) Stable cell lines are usually created using calcium phosphate transfection, an efficient and inexpensive method; (4) Cell lines can be established in cellular backgrounds that are specifically tailored to the needs of the protein being expressed. For example, heterogenous glycosolyation is a problem in NMR studies which typically require highly homogenous material. To overcome this problem, cell lines can be used that are deficient in complex heterogenous glycosylation capabilities (5); (5) The ability to create inducible cell lines is an additional advantage. In cases where constitutive expression of a particular protein is toxic to cells, placing the gene under an inducible promoter can allow expression of otherwise toxic proteins; (6) Finally, stable cells can be easily scaled up by transferring them to suspension cultures in spinner flasks or bioreactors. Episomal vectors developed from viruses, such as Epstein–Barr virus (EBV) (6), bovine papilloma virus (BPV) (7), BK virus (BKV) (8), and Simian virus 40 (SV40) (9) are mainly used for transfection. A list of commonly used vectors can be found in (10). The advantages of using episomal vectors over integrating genes into the DNA of the host cell are numerous (1) Episomal expression will result in gene expression independent of the regulatory mechanisms of the host cell and the position of integration in the host genome, leading to higher expression levels since these factors may unfavorably influence expression. Random integration may also disrupt the characteristics of the gene of interest (11); (2) There is no interruption in host cell gene expression, which is often the case with integrative vectors, the latter leading to transformation of the host cell and undesirable effects on protein production; (3) Episomal vectors can exist in multiple copies in a cell, thus leading to amplification of the gene of interest. The main criteria for selecting a suitable episomal vector are (1) high copy number in E. coli for large-scale DNA production; (2) a strong mammalian promoter

4

Isotope Labeling in Mammalian Cells

57

for high expression levels of the gene of interest in mammalian cells; and (3) small size so that genes of different lengths can be easily cloned into the vector. Thus, the vector typically consists of a strong mammalian promoter to drive expression of the mammalian gene of interest, a viral origin of replication activated by viral early genes, which is required to carry out its replication thereby maintaining the vector in the host system, and a eukaryotic selection marker, usually an antibiotic resistance gene, such as neomycin, to select for transfected cells. The human cytomegalovirus promoter has been shown to be very powerful in both HEK293 and Chinese Hamster Ovary (CHO) cells and is also active in most mammalian cells (12). 1.2. Mammalian Cell Lines

Mammalian cell lines used for transfection with virus-based vectors carrying the gene of interest usually already carry vectors encoding viral early genes corresponding to the origin of replication used by the new vector. Some of these early genes are SV40 T antigen, EBNA-1, E1, and E2 which bind to SV40, EBV, and BPV ori, respectively. The early genes act in trans to initiate replication by binding to the ori and they also act as enhancers, thereby increasing the copy number of viral vector and the expression levels of the protein of interest (13). Since very high copy numbers of the viral genes can lead to host cell death, regulation of viral replication can be imposed by transfecting mammalian cells with a different vector carrying the early genes (13). Mammalian cell lines that are widely used for protein expression are CHO and Human Embryonic Kidney 293 (HEK293) cells. Both are suitable for use in adherent and suspension cultures. The advantage of using CHO cells is the availability of various auxotrophs that can be used as selection markers for transfection. An example of such an auxotroph is dihydrofolate reductase (DHFR) deficient cells which are triple auxotrophs for hypoxanthine, glycine, and thymidine (14). Transfection of foreign genes along with DHFR genes in these cells allows for the selection of clones in a medium devoid of the above nutrients. Another advantage of this system is that it helps amplify the foreign gene when DHFR deficient cells are grown in the presence of methotrexate, which blocks DHFR activity. This causes the transfected cells to deal with low DHFR activity by amplifying the copy number of DHFR, thus amplifying the copy number of the transfected gene of interest. The disadvantages of the CHO cell line is that these cells do not carry all of the sugar transferring enzymes (15) that are present in human cell lines. This may lead to the production of functionally nonrelevant proteins. Further, some of the posttranslational modifications of human proteins are not appropriately carried out in these cells (16). It has also been seen that expression levels are usually lower in CHO cells than in HEK293 cells (12, 17). Therefore, the human cell line, HEK293, is used for proteins that require functionally sensitive posttranslational modifications.

58

A. Dutta et al.

There are two mutant cell lines of HEK293, HEK293E, expressing EBNA-1, and HEK293T, expressing SV40 large T antigen, that increase the copy number of plasmids with EBV and SV40 origins of replication, respectively, thereby increasing expression levels of the protein of interest (13). 1.3. Expression and Labeling of Recombinant Proteins in Mammalian Cells

Uniform isotope labeling of proteins expressed in mammalian cells is still under development (18). This is because mammalian cells, like insect cells, are unable to grow in the type of minimal media that are used for bacteria where glucose or glycerol and ammonium chloride are the only sources of carbon and nitrogen, respectively. Higher eukaryotes, like the organisms they represent, require certain essential amino acids, without which they are unable to grow. Thus, the use of simple sources for 13C and 15N isotope labeling of proteins in bacteria is not possible for mammalian cells. A number of recent approaches, however, have been developed to address this complication. The first uniformly isotope-labeled protein purified from mammalian cells was obtained by Hansen et al. (19) followed by (20, 21). The method involved purification of a mixture of isotope labeled amino acids from an acid hydrolysate of algae or bacteria grown in 15NH4Cl and 13C glucose or 13CO2 to prepare uniformly labeled urokinase in a Sp2/0 mouse myeloma cell line. Different purification protocols of the labeled mixture were tested but only acid hydrolysis, which removes bacterial and algal products that are toxic to mammalian cells, proved to work. The mixture, however, needed to be supplemented with commercially available amino acids that would degrade during hydrolysis, particularly glutamine and cysteine. A commercially available form of 15 N-cysteine was used. Because of the high cost involved in purchasing 15 N-glutamate, 15N-, and 15N/13C-glutamine were synthesized from labeled glutamate (available commercially), 15NH4Cl, and ATP. Dialyzed serum was used to prevent dilution of isotope labeled amino acids with unlabeled amino acids from the serum. Such purified amino acid mixtures are commercially available today. One such commercially available mixture from Martek Biosciences Corp. was tested to support expression in CHO cells (20). The growth medium was optimized in terms of its concentration, removal of amino acids, such as aspartic acid and asparagine, and addition of amino acids, such as arginine, cysteine, and glutamic acid after isolating and purifying from the mixture so that the highest level of protein expression was obtained. However, there are serious concerns regarding the toxic effects of feeding these mixtures to mammalian cells, leading to extensive methods of purification and optimization, and making it a time and cost inefficient method. HEK293 cells, the most commonly used system of expression for preparing isotope-labeled samples, have, to date, not been grown successfully in the presence of these mixtures (Klein-Seetharaman, unpublished results). An uniformly labeled 15N TGFb1 sample was prepared from CHO cells by

4

Isotope Labeling in Mammalian Cells

59

growing them in Minimum Essential Medium (MEM, Gibco) with dialyzed serum, uniformly 15N labeled choline and uniformly 15 N-labeled amino acids (except tryptophan which was labeled only in the backbone nitrogen) (22), demonstrating the utility of CHO systems. The total cost involved was $1,000/L. Therefore, this method could prove to be expensive if the protein to be labeled does not have high expression levels, as is the case with most membrane proteins. Mammalian growth media containing a mixture of certain isotope-labeled amino acids are commercially available and have been recently used for labeling the G-protein coupled receptor, bovine rhodopsin, using 15N labeled medium in which GKLQSTVW amino acids were 15N labeled (23). If all the amino acids in the mixture do not cover the complete sequence of the protein to be labeled, then this method may not lead to complete labeling, as was the case with rhodopsin where 50% of amino acids were labeled (23). Apart from the high costs involved, an additional problem is the inability to perform perdeuteration because of the sensitivity of mammalian cells to deuterium oxide. Therefore, the most widely used method to obtain structural information on mammalian proteins by NMR has been by conducting amino acid type selective (AATS) isotope labeling. AATS labels a protein with specific isotope-labeled amino acids rather than uniform labels and yields structural information selectively for the amino acid(s) used in labeling. This has been successfully done with numerous proteins (24–26). Extensive AATS labeling has been carried out with rhodopsin, which was labeled by using 15N-isotopes of tryptophan (26), lysine (25), histidine (27) and 13C-isotopes of tryptophan (27), histidine (27) and glutamate (28), all through expression in HEK293 cells. 15N-lysine, 13C-glycine/serine double labeling has also been shown for rhodopsin (25). For example, multiple combinations of 15N- and 13C-isotope-labeled rhodopsin samples were used to assign 15N-tryptophan resonances in the NMR spectrum of rhodopsin, shown in Fig. 1 (29). Since the protocol for such a labeling method has been optimized and works very well at least for rhodopsin, we are providing a step-by-step protocol below. The first step is the creation of an inducible stable cell line of rhodopsin in HEK293 cells. Scaling up the expression level is accomplished by growing these cells in a suspension culture, thereby producing the milligram quantities required for NMR studies. The yield of rhodopsin from such a transient transfection is on the order of 50 mg/15 cm2 plate. After establishing the stable cell line with the opsin (apo form of rhodopsin) gene, the next step is to transfer these cells to suspension culture and grow them in isotope-labeled medium containing unlabeled amino acids along with the labeled amino acid(s) of interest. Up to 10 mg of rhodopsin can be obtained from such suspension cultures when amino acid supplements are provided (30), but typically 2 mg of rhodopsin are obtained from

60

A. Dutta et al.

Fig. 1. Dephased (red line) and nondephased (blue line) 15N detected 13C/15N CP REDOR spectra of selectively labeled rhodopsin in DOPC lipid bilayers at 220 K. (a) a,e-15N-Trp; (b) 13C¢-Leu/a,e-15N-Trp; (c) 13U-Thr/a,e-15N-Trp; (d) 13C¢-Cys/a,e15 N-Trp; (e) 13C¢-Pro/a,e-15N-Trp; (f) 13C¢-Gly/a,e-15N-Trp. Figure reproduced with permission from (29).

4

Isotope Labeling in Mammalian Cells

61

isotope labeled medium in which amino acid mixtures cannot be supplemented, to avoid dilution of the isotope-labeled amino acids provided in the medium.

2. Materials 2.1. Expression of Isotope-Labeled Recombinant Proteins

1. Plasmid containing the gene of interest (see Note 1). 2. HEK293 cells. 3. Dulbecco’s modified Eagle medium (DMEM F-12). 4. Blasticidin (500 mg/mL stock solution prepared in DMEM F-12). 5. Geneticin, G418 (100 mg/mL stock solution prepared in DMEM F-12). 6. 0.05% Trypsin–EDTA (Gibco). 7. Penicillin–streptomycin (PS): 100 U/mL of each. 8. Fetal bovine serum (FBS). 9. Phosphate buffer saline (PBS): Autoclaved. 10. Tetracycline: 200 mg/mL (100× stock). 11. Sodium butyrate: 500 mM (100× stock), filtered. 12. Complete medium: DMEM F-12, 10% FBS, 1% PS. 13. Induction medium: Complete medium containing 2 mg/mL tetracycline and 5 mM sodium butyrate. 14. Selection medium: Complete medium containing 5 mg/mL blasticidin and 1, 2, or 3 mg/mL of G418. 15. Cryo medium: Complete medium containing 10% DMSO. 16. 2.5 M CaCl2. 17. BES: 50 mM N,N-bis(2-hydroxyethyl)-2-aminoethanesulfonic acid, pH 7.2, 250 mM NaCl, 1.5 mM Na2HPO4, 1 M NaOH used to adjust pH. 18. 15- and 10-cm2 Tissue culture plates. 19. 6- and 24-Well cell culture plates. 20. Sterile forceps. 21. Cloning rings. 22. Vacuum grease. 23. Dodecyl maltoside.

2.2. Specific Isotope Labeling of Proteins

Suspension DMEM medium: 1. Table 1 gives the individual components of suspension DMEM in their final concentrations based on the composition of commercially available DMEM. 100× stock solutions are prepared

mg/L

84

42

105

105

146

30

66

95

16

94

Essential amino acid

Arginine  HCl

Histidine  HCl  H2O

Isoleucine

Leucine

Lysine  HCl

Methionine

Phenylalanine

Threonine

Tryptophan

Valine

42 104

Tyrosine  2Na+  2H2O

None

30

584

Serine

Proline

Glycine

Glutamine

None

63

Cystine  2HCl Glutamate

None

None

None

mg/L

Aspartate

Asparagine

Alanine

Nonessential amino acid

Table 1 Amino Acid Composition of suspension DMEM

pantothenate

0.4 4

Thiamine  HCl

4

Pyridoxal  HCl Riboflavin

4

7.2

4

4

4

mg/L

Niacinamide

i-Inositol

Folic Acid

Choline Chloride

D-Ca

Vitamins

Glucose

Phenol Red  Na+

NaH2PO4  H2O

NaCl

KCl

MgSO4

Fe(NO3)3  9H2O

CaCl2

Inorganic salts and other

4,500

15

125

6,400

400

97.7

0.1

50

mg/L

62 A. Dutta et al.

4

Isotope Labeling in Mammalian Cells

63

for most components (see Note 2). Each component is dissolved in distilled water. 2. 0.05% trypsin–EDTA (Subheading 2.1). 3. FBS (Subheading 2.1). 4. PS (Subheading 2.1). 5. Dialyzed FBS: Heat inactivated, dialyzed against PBS and then filtered. 6. 10% Pluronic F-68 (100× stock), filtered. 7. 5 mg/mL Heparin (100× stock), filtered. 8. 15-cm2 tissue culture plates. 9. 2-L spinner flasks. 10. 100× tetracycline (Subheading 2.1). 11. 100× sodium butyrate (Subheading 2.1). 12. 20% (w/v) glucose, sterile filtered. 13. 8% (w/v) NaHCO3. 14. Complete medium (Subheading 2.1). 15. Selection medium (Subheading 2.1).

3. Methods 3.1. Expression of Isotope-Labeled Recombinant Proteins

Following is a protocol for generating an inducible stable cell line of HEK293 (30–32). 1. Wake up HEK293 cells from cryostocks and maintain them in 15 cm2 plates in 25 mL of complete medium containing 5 mg/mL blasticidin. 2. Split the cells into a 10-cm2 dish at 80% confluency. 3. Split the cells 1:10 or 1:8 (~1–2 million cells per plate) into complete medium containing 5 mg/mL blasticidin the day before transfection. 4. On the day of transfection, the cells should be 30–40% confluent. To transfect the cells, prepare the following cocktail, adding components in the order given (volumes are for one 10-cm2 plate): In a falcon tube, mix 30 mg of plasmid DNA, 50 mL of 2.5 M CaCl2, 500 mL of BES, and sterile water to 1 mL. Incubate the mixture for exactly 1 min and gently add it to the cells. After 1 h, verify the efficiency of transfection by the presence of calcium phosphate precipitate which appears as fine sand particles between cells. 5. Incubate the plates at 35°C and 3% CO2 for 19 h.

64

A. Dutta et al.

6. After 19 h, wash the plates twice with complete medium containing 5 mg/mL blasticidin and add fresh complete medium containing 5 mg/mL blasticidin. 7. Incubate the cells at 37°C and 5% CO2 overnight. 8. Split the cells 1:10 (0.5–1 million cells/plate) into nine plates, three plates each for selection with G418 (1, 2, and 3 mg/mL). 9. Replace the medium after 20 h with selection medium. 10. Replace the selection medium every 2–3 days until colonies of workable size are formed. 11. On the day of picking the clones: (a) Circle the colonies that are to be picked such that the circles are big enough to place the cloning rings on its perimeter. (b) Aspirate the medium and look for more colonies on the empty plate by eye. Wash with 8 mL of PBS. (c) Using sterile forceps, dip one end (base) of each cloning ring in vacuum grease and place it over each clone (see Note 3). Make sure there is no grease on the inner walls of the rings. Make sure that the rings form a well around the clones with a leak proof seal at the bottom. (d) Add 40 mL of 0.05% trypsin–EDTA to each well and incubate for 1 min. Then, add 80 mL of fresh selection medium and gently pipette up and down. Transfer the cells (100 mL) to a 24-well plate containing 1 mL of selection medium per well. (e) Repeat for all the clones. 12. Replace the selection medium every 2–3 days until the cells are confluent. Make a note of medium changes and transfers to 6-well plates in a chart (see step 13 for details). 13. As the cells become confluent, transfer them to two 6-well plates at different cell densities. For the transfer: (a) Pipette 3 mL of selection medium into each well of a 6-well plate. (b) Aspirate the medium from the cells (24-well plate) and add 300 mL of 0.05% trypsin–EDTA to each well and incubate for 1 min. (c) Add 1,200 mL (use a 1-mL pipette and add 600 + 600 mL) of selection medium and gently pipette up down. (d) Remove 300 mL and add it to one well and place the remaining (1,200 mL) in another well. The well with higher cell density will be induced in the future to screen the clones while the one with lower density will be maintained and used for making glycerol stocks.

4

Isotope Labeling in Mammalian Cells

65

14. Change the selection medium on the cells every 2–3 days. 15. As soon as the cells in the higher density well reach confluence, induce with induction medium. 16. Harvest the cells 48-h post induction. (a) After harvesting, solubilize the cells in a detergent suitable for the membrane protein of interest (e.g., 1% dodecyl maltoside). (b) Centrifuge the solubilized cells at 126,000 × g for 20 min at room temperature and collect the supernatant. 17. Check for levels of expression by performing a Western dot blot on serial dilutions of the supernatant in PBS, containing 1% of the detergent used for solubilization, versus protein samples of known concentrations (see Note 4). 18. When the cells from the low density well reach confluence, split 1:5 into two 10-cm2 cell culture dishes. Add 0.5 mL of 0.05% trypsin–EDTA to a 6-well plate, and then add 1.5 mL of selection medium. Add 1 mL of this cell suspension to each of the two 10-cm2 plates. 19. After the cells reach confluence on the 10-cm2 plate, prepare three 1-mL glycerol stocks from each plate. To prepare glycerol stocks: (a) Wash 70–90% confluent plates twice with PBS and trypsinize with 1 mL of 0.05% trypsin–EDTA for 1 min. (b) Add 10 mL of complete medium (no selection) and collect the cells in a 15-mL falcon tube. (c) Centrifuge the cells at 600 × g for 10 min at 4°C. (d) Aspirate the medium and gently resuspend the cells in 3 mL of cryo medium. (e) Transfer the cells to cryo vials in 1-mL aliquots and place the tubes in cryo boxes. Incubate at −20°C for 1 h, and then −80°C overnight. (f) On the following day, move the tubes to liquid nitrogen storage tanks. 3.2. Specific Isotope Labeling of Proteins

After estimating the yield from above (Subheading 3.1) and identifying the highest yield clone, the following procedure is used to prepare a protein sample, in suspension culture, in which particular amino acids are specifically isotope labeled (30–32). 1. Grow the highest yielding stable cell line clone in complete medium. 2. Split the cells 1:5 into 10-cm2 dishes after ~3 days using 15 mL of selection medium.

66

A. Dutta et al.

3. Split the cells until a sufficient number of plates are obtained for setting up spinner flasks. For reference, 1 mg of protein per liter of suspension culture is usually obtained for rhodopsin and 3–4 confluent 15-cm2 plates are used to inoculate a 500-mL flask (see Notes 5 and 6). 4. Prepare labeled suspension DMEM medium from 100× stocks of the individual components (Table 1, Subheading 2.2), except for the isotope labeled amino acid(s). Supplement with 10% dialyzed FBS and 1% PS or obtain isotope-labeled medium from a commercial source. (a) Add glucose, NaCl, glutamine, isotope-labeled amino acid(s) as solids. (b) Lower the glutamine concentration to half when starting the suspension culture. (c) Add Pluronic F-68 and heparin to 1× concentration. (d) Add isotope-labeled amino acid(s) as solids. 5. Inoculate the suspension cultures and grow at 37°C, spinning at 47 rpm for 6 days. To inoculate, add 2 mL of 0.05% trypsin– EDTA to each plate and extract the cells with 8 mL of unlabeled suspension medium. Collect cells from each plate (10-mL volume) and centrifuge at 1,000 × g for 10 min at 4°C. Aspirate the medium and add 10 mL of labeled suspension DMEM medium to each plate and resuspend the cells. Add the cell suspension to the suspension culture in the spinner flask. 6. After 6 days (see Note 7), supplement the growth medium with 6 mL of 20% (w/v) glucose and 4 mL of 8% (w/v) NaHCO3 and induce expression with 5 mM sodium butyrate and 2 mg/mL of tetracycline. 7. After the 6th day, feed the cells with 6 mL of 20% glucose every 24 h. 8. Grow the cells for 2 more days after the 6th day (total 8 days in suspension). 9. Harvest at the end of 8 days.

4. Conclusions A variety of expression systems have been developed that can be used to express a protein of choice in isotope-labeled form, each system having advantages and disadvantages. These systems include bacterial, yeast, insect, and mammalian cells. The choice of the system largely depends on the type of the protein to be expressed. Other factors are the costs involved, the need for modifications and

4

Isotope Labeling in Mammalian Cells

67

yield of protein required. Bacterial expression systems are most commonly used for soluble protein expression due to ease of use and cost-effectiveness in cloning and expression. However, this system is not the system of choice for proteins that require posttranslational modification for their activity. In such cases, eukaryotic expression systems are required. Yeast being both eukaryotic and a microorganism is a simple system to use, allowing relatively inexpensive complete isotope labeling of proteins for NMR studies. Although it can carry out some posttranslational modifications, these are often not sufficient and deficiency in folding machinery can be a problem for some proteins, especially membrane proteins. For these reasons, higher eukaryotic systems, such as insect cells have been in use where posttranslational modifications are very similar to those found in mammalian cells. The greatest advantage of insect cell expression is the very large amounts of proteins that can be expressed, in most cases more than that in mammalian cells. However, disadvantages of expensive isotope-labeled media, incomplete labeling and intolerability to perdeuteration that hamper mammalian cell expression are also prevalent in insect cells. Hence, efforts are ongoing towards further developing mammalian expression systems since these are the only systems in which the full complement of folding and posttranslational modification machineries are available, optimally supporting the activity of the expressed protein.

5. Notes 1. A tetracycline (tet)-regulated mammalian expression vector should be used. 2. All solutions were prepared as 100× concentrated stock solutions, except glucose, NaCl, glutamine, and the isotopelabeled amino acid(s), which were added as solids. Typically, 500 mL of stock solutions were prepared and filtered to maintain sterility. Sterile stock solutions were kept at 4°C and were stable for several weeks. 3. Cloning rings, vacuum grease, and forceps should be autoclaved. 4. PVDF membranes should not be used. 5. Cells should be counted before adding them to spinner flasks so that appropriate number of cells (in the range between 60 and 90 million cells/500 mL) can be transferred. 6. A 2-L spinner flask is ideal for a 500-mL suspension culture. 7. The color of the medium should turn yellow after 6 days.

68

A. Dutta et al.

References 1. Nettleship, J. E., Assenberg, R., Diprose, J. M., Rahman-Huq, N., and Owens, R. J. (2010) Recent advances in the production of proteins in insect and mammalian cells for structural biology. J. Struct. Biol. 172, 55–65. 2. Aricescu, A. R., Lu, W., and Jones, E. Y. (2006) A time- and cost-efficient system for highlevel protein production in mammalian cells. Acta. Crystallogr. D Biol. Crystallogr. 62, 1243–1250. 3. Lee, J. E., Fusco, M. L., and Ollmann Saphire, E. (2009) An efficient platform for screening expression and crystallization of glycoproteins produced in human cells. Nat. Protoc. 4, 592–604. 4. Nettleship, J. E., Rahman-Huq, N., and Owens, R. J. (2009) The production of glycoproteins by transient expression in mammalian cells. Methods Mol. Biol. 498, 245–263. 5. Reeves, P. J., Callewaert, N., Contreras, R., and Khorana, H. G. (2002) Structure and function in rhodopsin: high-level expression of rhodopsin with restricted and homogeneous N-glycosylation by a tetracycline-inducible N-acetylglucosaminyltransferase I-negative HEK293S stable mammalian cell line. Proc. Natl. Acad. Sci. U.S.A. 99, 13419–13424. 6. Sclimenti, C. R., and Calos, M. P. (1998) EpsteinBarr virus vectors for gene expression and transfer. Curr. Opin. Biotechnol. 9, 476–479. 7. Sambrook, J., Rodgers, L., White, J., and Gething, M. J. (1985) Lines of BPV-transformed murine cells that constitutively express influenza virus hemagglutinin. EMBO J. 4, 91–103. 8. Grossi, M. P., Caputo, A., Rimessi, P., Chiccoli, L., Balboni, P. G., and Barbanti-Brodano, G. (1988) New BK virus episomal vector for complementary DNA expression in human cells. Arch. Virol. 102, 275–283. 9. Piechaczek, C., Fetzer, C., Baiker, A., Bode, J., and Lipps, H. J. (1999) A vector based on the SV40 origin of replication and chromosomal S/MARs replicates episomally in CHO cells. Nucleic Acids Res. 27, 426–428. 10. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, CSH Laboratory Press, Cold Spring Harbor, NY. 11. Doerfler, W., Schubbert, R., Heller, H., Kammer, C., Hilger-Eversheim, K., Knoblauch, M., and Remus, R. (1997) Integration of foreign DNA and its consequences in mammalian systems. Trends Biotechnol. 15, 297–301. 12. Xia, W., Bringmann, P., McClary, J., Jones, P. P., Manzana, W., Zhu, Y., Wang, S., Liu, Y., Harvey, S., Madlansacay, M. R., McLean, K.,

Rosser, M. P., MacRobbie, J., Olsen, C. L., and Cobb, R. R. (2006) High levels of protein expression using different mammalian CMV promoters in several cell lines. Protein Expr. Purif. 45, 115–124. 13. Van Craenenbroeck, K., Vanhoenacker, P., and Haegeman, G. (2000) Episomal vectors for gene expression in mammalian cells. Eur. J. Biochem. 267, 5665–5678. 14. Urlaub, G., and Chasin, L. A. (1980) Isolation of Chinese hamster cell mutants deficient in dihydrofolate reductase activity. Proc. Natl. Acad. Sci. U.S.A. 77, 4216–4220. 15. Grabenhorst, E., Schlenke, P., Pohl, S., Nimtz, M., and Conradt, H. S. (1999) Genetic engineering of recombinant glycoproteins and the glycosylation pathway in mammalian host cells. Glycoconj. J. 16, 81–97. 16. Suttie, J. W. (1986) Report of Workshop on expression of vitamin K-dependent proteins in bacterial and mammalian cells. Madison, Wisconsin, USA, April 1986, Thromb. Res. 44, 129–134. 17. Schlaeger, E. J., Kitas, E. A., and Dorn, A. (2003) SEAP expression in transiently transfected mammalian cells grown in serum-free suspension culture. Cytotechnology 42, 47–55. 18. Takahashi, H., and Shimada, I. (2010) Production of isotopically labeled heterologous proteins in non-E. coli prokaryotic and eukaryotic cells. J. Biomol. NMR 46, 3–10. 19. Hansen, A. P., Petros, A. M., Mazar, A. P., Pederson, T. M., Rueter, A., and Fesik, S. W. (1992) A practical method for uniform isotopic labeling of recombinant proteins in mammalian cells. Biochemistry 31, 12713–12718. 20. Lustbader, J. W., Birken, S., Pollak, S., Pound, A., Chait, B. T., Mirza, U. A., Ramnarain, S., Canfield, R. E., and Brown, J. M. (1996) Expression of human chorionic gonadotropin uniformly labeled with NMR isotopes in Chinese hamster ovary cells: an advance toward rapid determination of glycoprotein structures. J. Biomol. NMR 7, 295–304. 21. Shindo, K., Masuda, K., Takahashi, H., Arata, Y., and Shimada, I. (2000) Backbone 1H, 13C, and 15N resonance assignments of the anti-dansyl antibody Fv fragment. J. Biomol. NMR 17, 357–358. 22. Archer, S. J., Bax, A., Roberts, A. B., Sporn, M. B., Ogawa, Y., Piez, K. A., Weatherbee, J. A., Tsang, M. L., Lucas, R., Zheng, B. L., and et al. (1993) Transforming growth factor beta 1: NMR signal assignments of the recombinant protein expressed and isotopically enriched using Chinese hamster ovary cells. Biochemistry 32, 1152–1163.

4 23. Werner, K., Richter, C., Klein-Seetharaman, J., and Schwalbe, H. (2008) Isotope labeling of mammalian GPCRs in HEK293 cells and characterization of the C-terminus of bovine rhodopsin by high resolution liquid NMR spectroscopy. J. Biomol. NMR 40, 49–53. 24. Arata, Y., Kato, K., Takahashi, H., and Shimada, I. (1994) Nuclear magnetic resonance study of antibodies: a multinuclear approach. Methods Enzymol. 239, 440–464. 25. Klein-Seetharaman, J., Reeves, P. J., Loewen, M. C., Getmanova, E. V., Chung, J., Schwalbe, H., Wright, P. E., and Khorana, H. G. (2002) Solution NMR spectroscopy of (alpha -15N) lysine-labeled rhodopsin: The single peak observed in both conventional and TROSYtype HSQC spectra is ascribed to Lys-339 in the carboxyl-terminal peptide sequence. Proc. Natl. Acad. Sci. U.S.A. 99, 3452–3457. 26. Klein-Seetharaman, J., Yanamala, N. V., Javeed, F., Reeves, P. J., Getmanova, E. V., Loewen, M. C., Schwalbe, H., and Khorana, H. G. (2004) Differential dynamics in the G protein-coupled receptor rhodopsin revealed by solution NMR. Proc. Natl. Acad. Sci. U.S.A. 101, 3409–3413. 27. Patel, A. B., Crocker, E., Reeves, P. J., Getmanova, E. V., Eilers, M., Khorana, H. G., and Smith, S. O. (2005) Changes in interhelical hydrogen bonding upon rhodopsin activation. J. Mol. Biol. 347, 803–812.

Isotope Labeling in Mammalian Cells

69

28. Han, M., and Smith, S. O. (1995) Highresolution structural studies of the retinal– Glu113 interaction in rhodopsin. Biophys. Chem. 56, 23–29. 29. Werner, K., Lehner, I., Dhiman, H. K., Richter, C., Glaubitz, C., Schwalbe, H., KleinSeetharaman, J., and Khorana, H. G. (2007) Combined solid state and solution NMR studies of alpha,epsilon-15N labeled bovine rhodopsin. J. Biomol. NMR 37, 303–312. 30. Reeves, P. J., Kim, J. M., and Khorana, H. G. (2002) Structure and function in rhodopsin: a tetracycline-inducible system in stable mammalian cell lines for high-level expression of opsin mutants. Proc. Natl. Acad. Sci. U.S.A. 99, 13413–13418. 31. Reeves, P. J., Callewaert, N., Contreras, R., and Khorana, H. G. (2002) Structure and function in rhodopsin: high-level expression of rhodopsin with restricted and homogeneous N-glycosylation by a tetracycline-inducible N-acetylglucosaminyltransferase I-negative HEK293S stable mammalian cell line 452. Proc. Natl. Acad. Sci. U.S.A. 99, 13419–13424. 32. Reeves, P. J., Thurmond, R. L., and Khorana, H. G. (1996) Structure and function in rhodopsin: high level expression of a synthetic bovine opsin gene and its mutants in stable mammalian cell lines. Proc. Natl. Acad. Sci. U.S.A. 93, 11487–11492.

Chapter 5 Cell-Free Protein Production for NMR Studies Mitsuhiro Takeda and Masatsune Kainosho Abstract The cell-free expression system using an Escherichia coli extract is a practical method for producing isotope-labeled proteins. The advantage of the cell-free system over cellular expression is that any isotopelabeled amino acid can be incorporated into the target protein with minimal scrambling, thus providing opportunities for advanced isotope labeling of proteins. We have modified the standard protocol for E. coli cell-free expression to cope with two problems specific to NMR sample preparation. First, endogenous amino acids present in the E. coli S30 extract lead to dilution of the added isotope. To minimize the content of the remaining amino acids, a gel filtration step is included in the preparation of the E. coli extract. Second, proteins produced by the cell-free system are not necessarily homogeneous due to incomplete processing of the N-terminal formyl-methionine residue, which complicates NMR spectra. Therefore, the protein of interest is engineered to contain a cleavable N-terminal histidine-tag, which generates a homogeneous protein after the digestion of the tag. Here, we describe the protocol for modified E. coli cell-free expression. Key words: Cell-free synthesis, S30 extract from E. coli, NMR, Stable isotope labeling

1. Introduction The NMR study of a protein starts by expressing the isotope-labeled protein. It is now widely accepted that the cell-free expression system is one of the most practical methods available for producing isotope-labeled proteins, along with the cellular expression system (1, 2). The cell-free system employs a cell extract from an organism such as Escherichia coli (3–7), an extract from wheat germ (8) or a mixture of recombinant proteins (9). As a consequence of its reduced amino acid metabolic activity in vitro, isotope-labeled amino acids can be efficiently incorporated into a target protein with minimal scrambling (10), which generates new opportunities for advanced selective isotope labeling of proteins, such as stereoarray isotope-labeled proteins (11–13). Alexander Shekhtman and David S. Burz (eds.), Protein NMR Techniques, Methods in Molecular Biology, vol. 831, DOI 10.1007/978-1-61779-480-3_5, © Springer Science+Business Media, LLC 2012

71

72

M. Takeda and M. Kainosho

We utilize a cell-free system, in which the transcription and translation systems are reconstituted in a vessel by combining an E. coli extract and several components. The E. coli cell-free system has a long history and the level of expression has been improving due to extensive investigations. The E. coli cell-free protein expression system is generally regarded as a practical method for producing isotope-labeled proteins, along with the cellular expression system. E. coli extract is now commercially available. In many of the E. coli cell-free systems, transcription is mediated by T7 RNA polymerase, and thus any plasmid encoding a DNA sequence under the control of the T7 promoter, such as the pET vectors (Novagen), can be used. Therefore, it is now relatively easy to test for expression of proteins of interest by using the E. coli cell-free system. We encountered two problems in the standard protocol of the E. coli cell-free procedure in terms of NMR sample preparation, and thus addressed the problems (14). First, the endogenous amino acids present in the E. coli S30 extract are incorporated into the target protein, which leads to isotope dilution. When (2H, 13C, 15 N)-calmodulin (CaM) protein was produced by using an E. coli S30 extract prepared by the standard protocol, the labeling efficiency was only ~90%. Therefore, to minimize the residual amino acids, we introduced a gel filtration step into the protocol for preparing the E. coli S30 extract. As a result, the labeling efficiency increased to 96% without loss of activity in the extract (14). Second, the formyl group of the N-terminal methionine in proteins produced by the cell-free system is not processed completely. The presence of the formyl moiety causes large, chemical shift changes in nearby residues, giving rise to doubled peaks, which correspond to formylated and deformylated forms. For example, the NMR signals from CaM, which has a flexible N terminus, are affected by the heterogeneity over an extensive region (14). Larger spectral complications may arise from heterogeneity of the peptide chain in proteins with a structured N terminus. We solved this problem by engineering the target protein with a cleavable N-terminal tag. After cleaving the tag, the protein was homogeneous, and the (1H, 15N) heteronuclear single quantum correlation (HSQC) spectrum of this CaM preparation lacked detectable extra peaks (14). To engineer proteins with an N-terminal cleavable tag, a template vector encoding an N-terminal tag followed by the multicloning site is useful. We constructed a family of vectors by modifying a pIVEX2.3d vector (Roche) to encode the N-terminal tag. In these vectors, the amino acid sequences are identical, but the DNA sequence in the region of the N-terminal tag is different. Introducing silent mutations into the DNA sequence in the N-terminal region can lead to different expression levels of the target protein (14). Therefore, plasmids with a higher expression level can be isolated when the target DNA sequence is introduced into the modified

5

Cell-Free Protein Production for NMR Studies

73

pIVEX vectors. In the case of CaM, for example, improvements of the protein yields relative to original sequence were observed for two of the ten pIVEX vectors. In the large-scale reaction performed with the best sequence and 51.1 mg of amino acid mixture (1.7 mg/mL), the amount of CaM synthesized was 5.2 mg (10 wt%). This was twice the yield from the construct prior to the silent mutagenesis (14).

2. Materials 2.1. Preparation of E. coli S30 Extract

1. RNase-free Milli-Q PF Plus water (Millipore) (see Note 1). 2. LB medium: Dissolve 20 g of Luria Broth powder in 1 L of water. 3. Incomplete rich medium: Dissolve 5.6 g of KH2PO4, 28.9 g of K2HPO4, 1 g of Bacto yeast extract, and 1.5 mg of thiamine hydrochloride in 1 L of water, and autoclave the solution. Cool to room temperature and add 50 mL of 40% (w/v) D-glucose and 10 mL of 100 mM Mg(OAc)2. 4. BL21 Star (DE3) cells (Invitrogen) (see Note 2). 5. French press (see Note 3). 6. S30 buffer: Combine 10 mL of 1 M Tris–OAc, pH 8.2, 10 mL of 1.4 M Mg(OAc) 2, and 10 mL of 6 M KOAc in a final volume of 1 L of RNase-free water, atuoclave, and add 1 mL of 1 M DTT. 7. Gel filtration apparatus: Pack Sephadex G25 resin into a 2.5 × 20-cm chromatography column and set the column vertically at 4°C. After the resin settles, attach a funnel to the top of the column and equilibrate with 500 mL of S30 buffer (see Note 4). 8. Dialysis tubing (molecular weight cutoff: 6–8 kDa). 9. Polyethylene glycol (PEG)-8000: Avg. MW 8,000 Da. 10. 2-mercaptoethanol. 11. Diethyl pyrocarbonate (DEPC). 12. Centrifugation tube: Wash the tubes with DEPC water and autoclave to completely remove RNase. 13. 40% (w/v) D-glucose. 14. 100 mM Mg(OAc)2. 15. 1 M Tris–OAc, pH 8.2. 16. 1.4 M Mg(OAc)2. 17. 6 M KOAc. 18. 1 M DTT.

74

M. Takeda and M. Kainosho

2.2. Preparation of T7 RNA Polymerase

1. M9 medium supplemented with tryptone peptone: Combine 1 g of Tryptone peptone (Bectone), 5 g of NaCl, 1 g of NH4Cl, 3 g of KH2PO4, and 6 g of Na2HPO4, and add water to 1 L. After autoclaving, add 10 mL of sterile-filtered 40% (w/v) glucose and 1 mL of sterile 1 M MgSO4. 2. Sonication buffer A: 50 mM Tris–HCl, pH 8.1, 20 mM NaCl, 2 mM EDTA, and 1 mM DTT. 3. Sonication buffer B: 50 mM Tris–HCl, pH 8.1, 20 mM NaCl, 2 mM EDTA, 1 mM DTT, 1.5 mg/mL lysozyme, 20 μg/mL PMSF, 10 μg/mL bacitracin, and 0.1 mM benzamidine. 4. Dialysis buffer: 20 mM sodium phosphate–NaOH, pH 7.7, 1 mM EDTA, 1 mM DTT, and 5% glycerol. 5. SP-Sepharose Fast Flow (GE Healthcare): This resin is packed into a column (3 × 30 cm). 6. Elution buffer: 20 mM sodium phosphate–NaOH, pH 7.7, 1 mM EDTA, 1 mM DTT, 5% glycerol, 200 mM NaCl, and 20 μg/mL PMSF. 7. Isopropyl-beta-D-thiogalactopyranoside (IPTG). 8. 0.8% (w/v) sodium deoxycholate. 9. 2 M Ammonium sulfate. 10. 10% (w/v) Polyethleneimine–HCl, pH 7.0. 11. Saturated ammonium sulfate: Adjust pH to 7.0 using 1 M Tris. 12. E. coli strain BL21 (DE3) transformed with a plasmid encoding T7 RNA polymerase. 13. LB medium (Subheading 2.1). 14. 40% D-Glucose (Subheading 2.1). 15. 1 M MgSO4: Sterilize by autoclaving.

2.3. Cell-Free Synthesis Reaction

1. LM mixture: To prepare the low-molecular-weight mixture, combine 22 mL of 2 M HEPES–KOH, pH 7.5, 33.4 mL of 6 M K(OAc), 210 mg of DTT, 530 mg of ATP, 338 mg of CTP, 335 mg of GTP, 310 mg of UTP, 172 mg of cAMP, 28 mg of folinic acid, and 140 mg of tRNA, and, if needed, 64 mL 50% (w/v) PEG-8000 (see Note 5), and then add RNase-free water up to 200 mL (see Note 6). 2. Dialysis membrane for cell-free reaction: A size 8 cellulose membrane (Viskase Sales Corporation) ensures a reproducible expression level. Cut the membrane to an appropriate size. First, soak the membrane in RNase-free water and then warm it to 70–80°C. After it has cooled down, wash the membrane with RNase-free water. Second, soak the membrane in water containing ~50 mM of NaHCO3, warm it in a microwave to remove glycerol, and then wash it with RNase-free water. Third, soak the membrane in water containing ~1 mM EDTA, warm it in a microwave to further remove glycerol, and then

5

Cell-Free Protein Production for NMR Studies

75

wash it with RNase-free water three times. Finally, soak the membrane in water and autoclave it. 3. 0.645 M Creatine phosphate (see Note 7). 4. 10 mg/mL Creatine kinase (see Note 7). 5. RNase inhibitor (Porcine liver). 6. 1.4 M NH4OAc or (15N) NH4OAc (see Note 8). 7. T7 RNA polymerase at ~11 mg/mL (see Note 9). 8. Template DNA: A plasmid containing the sequence encoding the target protein at ~1 mg/mL (see Note 10). 9. 2× Loading buffer: 2 mL of 0.5 M Tris–HCl, pH 6.8, 4 mL of 10% sodium dodecyl sulfate (SDS), 1.2 mL of 2-mercaptoethanol, 2 mL of glycerol, 1–2 mg of bromophenol blue, and 0.8 mL of water. 10. RNase-free water (Subheading 2.1). 11. 0.5 M Mg(OAc)2. 12. S30 extract (Subheading 3.1). 13. Reaction solution: Mix 112 μL of RNase-free water, 9.8 μL of 1.4 M NH4OAc, 15 μL of 0.5 M Mg(OAc)2, 20 μL of SAIL amino acids, 40 μL of 0.645 M creatine phosphate, 125 μL of LM mixture, 10 μL of 1 mg/mL template DNA, 4.5 μL of 11 mg/mL T7 RNA polymerase, 1.25 μL of 40 U/mL RNase inhibitor, 12.5 μL of 10 mg/mL creatine kinase, and 150 μL of S30 extract for 0.5 mL total volume. 14. Dialysis solution: Mix 1160.8 μL of RNase-free water, 39.2 μL of 1.4 M NH4OAc, 60 μL of 0.5 M Mg(OAc)2, 80 μL of SAIL amino acids, 160 μL of 0.645 M creatine phosphate, and 500 μL of LM mixture for 2 mL total volume. 15. Phosphate buffer: 10 mM potassium phosphate, pH 7.0. Adjust pH using NaOH or HCl. 16. 6 M KOAc (Subheading 2.1). 17. 50% (w/v) PEG-8000. 18. 10% SDS. 19. 0.5 M Tris–HCl, pH 6.8. 20. 2-Mercaptoethanol.

3. Methods The E. coli S30 extract preparation protocol essentially follows previously reported procedures (3, 7, 15), but contains a gel filtration step to minimize the endogenous amino acids. Typically, preparation of the S30 extract takes 2–3 days for one person. We strongly recommend saving an aliquot of the extract at each step to monitor

76

M. Takeda and M. Kainosho

the activity throughout the purification. T7 RNA polymerase is also prepared following established methods (16–18). The preparation of T7 RNA polymerase takes 1 week for one person. The cell-free expression strategy for new proteins is shown in Fig. 1 . As the first step, we recommend starting with E. coli cellular expression of the target protein with uniform 15N labeling.

Fig. 1. Flowchart of the cell-free strategy (reproduced from ref. 14 with permission from Elsevier Science).

5

Cell-Free Protein Production for NMR Studies

77

It is less costly and more efficient to produce proteins from E. coli cells than by using the cell-free method, and cellular expression is better suited for optimizing the purification protocol and the NMR buffer conditions. In general, a protein that exhibits poor cellular expression is also likely to show poor cell-free expression, unless the protein produced is toxic to E. coli cells. Next, several parameters are optimized to maximize the expression level in the cell-free reaction based on small-scale reactions under different conditions. There are various parameters to consider in the optimization process, including the concentration of magnesium, incubation temperature, incubation time, and concentration of PEG. If necessary, the target DNA sequence can be transferred to the line of pIVEX vectors with different silent mutations, as described in the introduction. When an acceptable expression level is attained, a large quantity of the 15N-labeled protein is produced and its (1H, 15N)-HSQC spectrum is compared with that of the protein produced by cellular expression. This comparison helps to detect any possible difference between in vivo and in vitro expression. Special attention should be paid to the incomplete deformylation of the N terminus of proteins in the cell-free reaction. The presence of formylated and deformylated species complicates NMR spectra. To overcome this problem, the use of an engineered N-terminal cleavable tag is helpful, as shown in Fig. 2. 3.1. Preparation of E. coli S30 Extract

1. Inoculate 10 mL of LB medium with the E. coli stock cells (BL21 star (DE3) strain) and incubate the preculture overnight at 37°C with shaking. 2. Add the preculture to 1 L of incomplete rich medium prewarmed to 37°C, and incubate the culture at the same temperature with shaking to an optical density at 600 nm (OD600) of 0.7 (see Note 11). 3. Centrifuge the cells at 5,000 × g for 10 min at 4°C and then wash them three times with 200 mL of ice-cold S30 buffer supplemented with 0.05% 2-mercaptoethanol (see Note 12). Store the cells as a pellet at −80°C. 4. Gently resuspend the cell pellet in 200 mL of ice-cold S30 buffer supplemented with 0.05% 2-mercaptoethanol. Centrifuge at 5,000 × g for 10 min at 4°C and weigh the pellet. Resuspend the pellet in 1.27 mL of S30 buffer per gram of cells. 5. Disrupt the cells with a French Press at 20,000 psi (1,400 kg cm−2) (see Note 13). Immediately after the disruption, add 30 μL of 1 M DTT and centrifuge the lysate at 30,000 × g for 30 min at 4°C in DEPC-treated/autoclaved centrifuge tubes. Carefully collect ~1.4 mL of supernatant per gram of E. coli without mixing with the precipitate.

M. Takeda and M. Kainosho

a

105

110

Q3E2

Q3E2

T5 Q3E2

Q3E2

T70 D2 D2

115

T70 R74

R74 I9 I9

Q3 Q3 K77 K77

D78 D78

M71 M71

A73 A73

K13 K13

120

15N (p.p.m.)

T5

125 A1

L4

L4

M0

130

11

10

9

8

6

7

1H (p.p.m.)

b

105 G113 G33 G59

G40

N137D2

N137D2

Q135E2

Q135E2

T62 G96

N53D2

T5 G98 G25

G134

T29 T44

G61

G132 V55 G23 Q8E2 Q41E2 T146

D95

T117

T26 D58

T110

N53D2

N111D2

Q143E2 N42D2 S17

Q3E2 Q49E2

110

Q41E2 Q8E2 N111D2 Q143E2 N42D2 Q3E2 Q49E2

D22 M145 T79 N60D2

T70 F19 Q135 N60D2 E127 N97D2 R90 E54 M109 Y99 N97D2 M72 D93 D131 D129 F92 S81 D20 R74 E139 D2E67 R106 Q49 N60 T34 N53 I125 M76 I52 Q143 Y138 A103 V91 Q3 M36 Q41 K75 F65 F89 L69 R126 E87 E47 H107 E119 A128 R37 E84 S38V108 L112 M51 F16 E11 D122 M124 V142 E140 E104 N97 E123 Q8 L32 E7 I9 E83D50 M144 L39 E6 E120 E45 F12 D24 L18 L48 E114 K77 E14 K30 D133 A46 A10 E31 D118 L105 V121 D56 R86 D78 M71 V35 A102 I85 A88 L4 F68 A73 E82 A15 N111 K13 D80 A1 K21 S101 I63 K115 K148 V136 F141 L116 K94 N42

D64 I100

115

T28

120

125

A147

I130

I27

130

N137 A57

11

15N (p.p.m.)

78

10

9

8

7

6

Fig. 2. (1H, 15N) HSQC experiments of (U-13C, 15N) CaM. (a) CaM synthesized by the E. coli cell-free system. (b) CaM synthesized with the N-terminal tag following its removal by thrombin digestion. The extra peaks in (a) and the peaks in (b) are labeled with their assignments (reproduced from ref. 14 with permission from Elsevier Science).

5

Cell-Free Protein Production for NMR Studies

79

6. Transfer the collected supernatant to fresh DEPC-treated/ autoclaved tubes. Centrifuge at 30,000 × g for 30 min at 4°C and collect ~1.0 mL of the supernatant per gram of E. coli and place it into a 50-mL tube. 7. Incubate the 50-mL tube at 37°C for 80 min with shaking (see Note 14). 8. Dialyze the solution for 45 min against 2 L of S30 buffer at 4°C using dialysis tubing with a MWCO of 6–8,000 kDa. Repeat the dialysis, and then centrifuge the solution at 15,000 × g for 10 min at 4°C. 9. Apply the supernatant to a gel-filtration column preequilibrated at 4°C with S30 buffer. Load the supernatant onto the column. Maintain a continuous flow of S30 buffer in the column. When the sample first begins to exit the column, as judged from its color and turbidity (see Note 15), collect, in bulk, 1.4 times the total volume applied to the column. 10. Dialyze the collected bulk solution at 4°C for 40–50 min against 700 mL of an equal weight mixture of PEG-8000 and S30 buffer (see Note 16). Adjust the dialysis time so as to concentrate the extract to 0.86 times the initial volume. Dialyze the extract against 2 L of S30 buffer at 4°C for 60 min to remove PEG-8000. 11. Transfer the dialyzed S30 extract to 1.5-mL tubes, freeze it in liquid nitrogen, and store it at −80°C (see Note 17). 12. Assess the extent to which the endogenous amino acids in the extract are eliminated by preparing a protein by using an 2H-, 13 C-, and 15N-labeled amino acid mixture, e.g., SAIL amino acids set. The remaining endogenous amino acids give rise to proton signals in 13C-filtered 1H NMR experiments, as shown in Fig. 3. 3.2. Preparation of T7 RNA Polymerase

1. Using a glycerol stock of E. coli strain BL21 (DE3) transformed with a plasmid encoding T7 RNA polymerase, inoculate 10 mL of LB medium containing the appropriate antibiotic in a 50-mL tube and then incubate with shaking at 37°C overnight. 2. Inoculate 1 L of M9 medium supplemented with tryptone peptone, containing the appropriate antibiotic, with the 10 mL culture from step 1 into a 2-L flask that has been prewarmed to 37°C. Incubate the cells with shaking at 37°C until the culture reaches an OD600 of 0.5. 3. Induce the expression of T7 RNA polymerase by adding IPTG to a final concentration of 0.5 mM. Incubate the cells with shaking for 8 h at 37°C and then centrifuge the culture at 5,000 × g for 10 min at 4°C. Store the cell pellet at −80°C.

80

M. Takeda and M. Kainosho

Fig. 3. 13C-filtered 1H-NMR spectra of CaM. (a) 15N-labeled CaM synthesized with the conventional extract. (b) 2H-, 13C-, 15N-labeled CaM synthesized with the conventional S30 extract. (c) 2H-, 13C-, 15N-labeled CaM synthesized with the improved, dialyzed S30 extract. All of the spectra are adjusted for the intensities of the amide region. Since the peaks of the protons attached to 13C are filtered, only the protons attached to 12C give rise to resonances in the aliphatic region (reproduced from ref. 14 with permission from Elsevier Science).

4. Suspend the frozen cell pellet in 72 mL of sonication buffer A. Add 18 mL of sonication buffer B and stir the mixture for 15 min at 4°C (see Note 18). Add 7.5 mL of 0.8% sodium deoxycholate and stir the mixture for 15 min at 4°C. Disrupt the cells with an ultrasonic generator (five rounds of 1-min sonication at 5-min intervals). 5. Add 15 mL of 2 M ammonium sulfate and bring the volume of the lysate to 150 mL by adding sonication buffer A. Then, gradually add 15 mL of 10% polyethyleneimine and stir the mixture on ice for 20 min. Centrifuge at 39,000 × g for 15 min at 4°C.

5

Cell-Free Protein Production for NMR Studies

81

6. Gradually add 0.82 volume of saturated ammonium sulfate to the supernatant. Stir on ice for 15 min and centrifuge the mixture at 12,000 × g for 15 min at 4°C. 7. Resuspend the precipitate in 45 mL of dialysis buffer containing 100 mM NaCl and 20 μg/mL PMSF. Dialyze the suspension against 2 L of dialysis buffer containing 100 mM NaCl and 20 μg/mL PMSF for 3 h using a membrane with a molecular weight cutoff of 6,000–8,000. Repeat the dialysis for 3 h and overnight. 8. Centrifuge the dialysate at 12,000 × g for 10 min at 4°C. Dilute the dialysate with one volume of dialysis buffer containing 20 μg/mL of PMSF to produce conductivity equal to that of the dialysis buffer plus 50 mM NaCl. Load the dialysate on an SP-Sepharose FF column (3 × 30 cm), equilibrated with dialysis buffer containing 50 mM NaCl and 20 μg/mL of PMSF, and run the 2-column volumes of dialysis buffer containing 50 mM NaCl and 20 μg/mL of PMSF at a flow rate of 1 mL/min to elute impurities. If necessary, to further remove impurities remaining in the column, load up to 500 mL of dialysis buffer containing 50 mM NaCl and 20 μg/mL PMSF at a flow rate of 2 mL/min (see Note 19). 9. Elute the T7 RNA polymerase with elution buffer and dialyze it against dialysis buffer containing 100 mM NaCl and 20 μg/mL of PMSF at 4°C overnight. 10. Repeat the dialysis for 6 h (see Note 20). Finally, dialyze the T7 RNA polymerase against dialysis buffer containing 100 mM NaCl and 50% glycerol at 4°C overnight. Centrifuge at 18,000 × g for 10 min at 4°C, collect the supernatant, and determine the concentration of the T7 RNA polymerase by measuring the optical density at 280 nm (ε280 = 1.4 × 105 M−1 cm−1) (see Note 21). Transfer the protein to 1.5-mL tubes and store it at −20°C. 3.3. Cell-Free Synthesis Reaction

1. Wear sanitary gloves (see Note 22) and thaw the LM mixture and the S30 extract (see Note 23). 2. Prepare the reaction and dialysis solutions (Subheading 2.3). In the case of a small-scale reaction for evaluating expression levels, the typical volumes of the reaction and dialysis solutions are 0.5 and 2.0 mL, respectively. For large production quantities, each volume is scaled up while maintaining the volume ratio of the reaction solution to the dialysis solution (see Note 24). 3. Transfer the dialysis solution to an RNase-free vessel. Tie one end of the dialysis tube firmly. Transfer the reaction solution to the dialysis tube, and tie off the other end. Fold the tubing 4–6 times and place it into the vessel such that the dialysis tube is completely submerged in the dialysis solution.

82

M. Takeda and M. Kainosho

4. Incubate the vessel with shaking for 4–8 h at 37°C (see Note 25). 5. Retrieve the reaction and dialysis solutions. If the expressed protein has a molecular weight smaller than molecular weight cutoff of the membrane, then it will be found in both the reaction and dialysis solutions. 6. To evaluate the expression level, centrifuge 50-μL aliquots of the reaction and dialysis solutions at 5,000 × g for 1 min and 4°C for. Mix the supernatant with an equal volume of 2× loading buffer, incubate for 5 min at 95°C, and load 10 μL on a polyacrylamide gel. Dissolve the precipitate in 50 μL of phosphate buffer (pH 7.0), mix it with an equal volume of 2× loading buffer, and load 10 μL on a polyacrylamide gel.

4. Notes 1. Water, treated with DEPC and then autoclaved, can also be used as RNase-free water (19). However, handle it carefully, since DEPC is a carcinogen. 2. The expression level of BL21 star is 1.2–1.4 times higher than that of the A19 strain. 3. The disruption of the E. coli cells can be accomplished by using equipment other than a French press. 4. The uniformity of the filled resin affects the resolution of gel filtration. The resin is used only once for each preparation. 5. While the use of PEG increases the expression level in some cases, it hampers SDS-PAGE analysis. Therefore, PEG should be removed by ethanol precipitation when the sample is subjected to SDS-PAGE analysis. 6. The prepared LM mixture can be frozen at −20°C for 1 month or more. 7. The creatine phosphate and creatine kinase should be prepared just prior to use. 8. If 15N labeling of the side-chain amide groups in asparagine and glutamine residues is intended, use (15N)-NH4OAc. 9. While we prepare the T7 RNA polymerase ourselves, commercially available T7 RNA polymerase can also be used. 10. The cell-free system uses a large amount of the plasmid, and thus a high-copy plasmid saves both cost and time. 11. The growth rate of the cells in the culture correlates with the activity of the resulting extract (20). We strongly recommend monitoring the growth rate. If the grow rate is slow, then it

5

Cell-Free Protein Production for NMR Studies

83

might be better to restart the culture. On the other hand, overgrowth, i.e., OD600 larger than 0.9, leads to reduced activity of the cells. 12. Do not allow the suspension to foam. 13. The cell disruption is a critical step that affects the activity of the extract. 14. This runoff reaction was performed with a preincubation mixture (14). The expression level does not change, even without the supplement (21). 15. The fraction to be collected looks yellow. 16. Before use, stir the PEG-S30 buffer at 4°C with a large magnetic stir bar to avoid PEG deposition. 17. The presence of DTT is necessary for preserving the activity over time. The activity of an S30 extract prepared without DTT decreases by 10–20% after 3 months of storage at −80°C. 18. The viscosity of the solution increases because of the lysozyme. 19. Wash until the optical density at 280 nm becomes constant. 20. The protein may precipitate during the dialysis. 21. If necessary, concentrate the solution up to about 10 mg/mL with an ultrafiltration device. 22. When setting up the cell-free reaction, use sanitary gloves to prevent contamination with RNases. 23. The frozen S30 extract should be thawed gently on ice. 24. Tryptophan and tyrosine residues are unlikely to be soluble. If needed, warm the amino acid solutions to 60°C. Do not vigorously shake the solution after the template DNA has been added. 25. PEG is assumed to play a role as a stabilizer of the system. For example, in the case of CaM, the ratio of protein yields with and without PEG was 1.6. However, even a small amount of PEG can disturb the SDS-PAGE quantitation of the production level of a target protein. If the solution contains PEG, then precipitate the supernatant with ethanol. Empirically, however, it is difficult to completely remove PEG even by ethanol precipitation. Therefore, optimizing the PEG concentration is somewhat difficult in small-scale cell-free reactions.

Acknowledgments This work was supported by a grant from the Targeted Protein Research Program (MEXT) to M. Kainosho and a Grant-in-Aid for Young Scientists (B) (21770110) to M. Takeda.

84

M. Takeda and M. Kainosho

References 1. Clemens, M.J., and Prujin, G.J. (1999) Protein synthesis in Eukaryotic Cell-free systems. Oxford University Press, New York, pp. 129–165. 2. Kramer, G., Kudlicki, W., and Hardesty, B. (1999) Cell-free Coupled TranscriptionTranslation systems from Escherichia coli. Oxford University Press, New York, pp. 129–165. 3. Zubay, G. (1973) In vitro synthesis of protein in microbial systems. Ann. Rev. Gen. 7, 267–287. 4. Spirin, A.S., Barano, V.I., Ryabova, L.A., Ovodov, S.Y., and Alakhov, Y.B. (1988) A continuous cell-free translation system capable of producing polypeptides in high yield. Science 242, 1162–1164. 5. Kim, D.M., Kigawa, T., Choi, C.Y., and Yokoyama, S. (1996) A highly efficient cell-free protein synthesis system from Escherichia coli. Eur. J. Biochem. 239, 881–886. 6. Kim, D.M., and Swartz, J.R. (2000) Prolonging cell-free protein synthesis by selective reagent additions. Biotechnol. Prog. 16, 385–390. 7. Kigawa, T., Yabuki, T., Yoshida, Y., Tsutsui, M., Ito, Y., Shibata, T., and Yokoyama, S. (1999) Cell-free production and stable-isotope labeling of milligram quantities of proteins. FEBS Lett. 442,15-19. 8. Madin, K., Sawasaki, T., Ogasawara, T., and Endo, Y. (2000) A highly efficient and robust cell-free protein synthesis system prepared from wheat embryos: plants apparently contain a suicide system directed at ribosomes. Proc. Natl. Acad. Sci. U.S.A. 97, 559–564. 9. Shimizu, Y., Inoue, A., Tomari, Y., Suzuki, T., Yokogawa, T., Nishikawa, K., and Ueda, T. (2001) Cell-free translation reconstituted with purified components. Nat. Biotechnol. 19, 751–755. 10. Kigawa, T., Muto, Y., and Yokoyama, S. (1995) Cell-free synthesis and amino acid-selective stable isotope labeling of proteins for NMR analysis. J. Biomol. NMR 6, 129–134. 11. Kainosho, M., Torizawa, T., Iwashita, Y., Terauchi, T., Ono. A,M., and Güntert, P. (2006) Optimal isotope labelling for NMR

12.

13.

14.

15.

16.

17.

18.

19.

20.

21.

protein structure determinations. Nature 440, 52–57. Takeda, M., Ikeya, T., Güntert, P., and Kainosho, M. (2007) Automated structure determination of proteins with the SAIL-FLYA NMR method. Nature Protocol 2, 2896–2902. Kainosho, M., and Güntert, P. (2010) SAILStereo-array isotope labeling. Q. Rev. Biophys. 7, 1–54. Torizawa, T., Shimizu, M., Taoka, M., Miyano, H., and Kainosho, M. (2004) Efficient production of isotopically labeled proteins by cell-free synthesis: A practical protocol. J. Biomol. NMR 30, 311–325. Pratt, J.M. (1984) Transcription and Translation: A Practical Approach, IRL Press, New York, pp. 179–209. Davanloo, P., Rosenberg, A.H., Dunn, J.J., and Studier, F.W. (1984) Cloning and expression of the gene for bacteriophage T7 RNA polymerase. Proc. Natl. Acad. Sci. U.S.A. 81, 2035–2039. Zawadzki, V., and Gross, H.J. (1991) Rapid and simple purification of T7 RNA polymerase. Nucl. Acid Res. 19,1948. Grodberg, J., and Dunn, J.J. (1988) ompT encodes the Escherichia coli outer membrane protease that cleaves T7 RNA polymerase during purification. J. Bacteriol. 170, 1245–1253. Huang, Y.H., Leblanc, P., Apostolou, V., Stewart, B., and Moreland, R.B. (1995) Comparison of Milli-Q PF Plus water to DEPC-treated water in the preparation and analysis of RNA. Biotechniques 19, 656–661. Zawada, J., and Swartz, J.R. (2006) Effects of growth rate on cell extract performance in cellfree protein synthesis. Biotechnol. Bioeng. 94, 618–624. Liu, D.V., Zawada J.F., and Swartz, J.R. (2005) Streamlining Escherichia coli S30 extract preparation for economical cellfree protein synthesis. Biotechnol. Prog. 21, 460–465.

Chapter 6 Cell-Free Membrane Protein Expression for Solid-State NMR Alaa Abdine, Kyu-Ho Park, and Dror E. Warschawski Abstract Although cell-free expression is a relative newcomer to the biochemical toolbox, it has already been reviewed extensively, even in the more specialized cases such as membrane protein expression, nanolipoprotein particles, and applications to crystallography and nuclear magnetic resonance (NMR). Solid-state NMR is also a newcomer to the structural biology toolbox, with its own specificities in terms of sample preparation. Cell-free expression and solid-state NMR are a promising combination that has already proven useful for the structural study of membrane proteins in their native environment, the hydrated lipid bilayer. We describe below several protocols for preparing MscL, a mechanosensitive membrane channel, using cell-free expression destined for a solid-state NMR study. These protocols are flexible and can easily be applied to other membrane proteins, with minor adjustments. Key words: In vitro synthesis, Integral membrane proteins, Membrane protein reconstitution, Liposomes, Solid-state NMR

1. Introduction Cell-free expression is one of the major new developments in structural biology, both for nuclear magnetic resonance (NMR) and X-ray crystallography, because it allows for overproduction of proteins, both wild-type and mutant, that are often produced with prohibitively low yields using classical biosynthetic methods. Proteins that are expressed cell-free end up in the reaction container and hence in water where membrane proteins precipitate, often irreversibly. Cell-free expression was first thought to be impractical for membrane protein expression because the system did not include a biological apparatus for targeting the protein to the membrane (1). Providing the medium with detergent was also considered risky since detergent could perturb protein expression by interfering with the ribosome, polymerase or any other essential

Alexander Shekhtman and David S. Burz (eds.), Protein NMR Techniques, Methods in Molecular Biology, vol. 831, DOI 10.1007/978-1-61779-480-3_6, © Springer Science+Business Media, LLC 2012

85

86

A. Abdine et al.

component of the cell-free expression system. These obstacles were overcome in 2004, when several groups, using optimized cell lysates, developed protocols for membrane protein expression in vitro, in the presence of detergents (2–4). For X-ray crystallography and solution-state NMR, the major techniques used in structural biology today, membrane protein structure determination is still a challenge: first, because the aforementioned necessity of obtaining large quantity of functional proteins is aggravated, in vivo, by the limited membrane surface available (5) and second, because membrane proteins are hydrophobic and have to be manipulated in detergents at all times when they are extracted from their native environment, the lipid bilayer (6). Detergents or other surfactants are used for membrane solubilization and during protein purification (which often requires an affinity tag, such as a polyhistidine stretch). They can interfere with protein crystallization or make aggregates that are too large for solution-state NMR. Last but not the least, detergents can interfere with protein folding or function, and the protein structure or dynamics determined in a detergent environment is not necessarily representative of the native structure or dynamics (7, 8). Nevertheless, since 1985, over 200 membrane protein structures have been determined by using X-ray crystallography (9) and, since 1997, about 30 structures by using solution-state NMR (10). In this context, membrane protein cell-free expression has been developed and reviewed, especially for isotope labeling strategies, another issue for NMR, where in vitro synthesis offers a very efficient and versatile alternative that greatly minimizes amino acid scrambling (11, 12). Solid-state NMR is an alternative for membrane protein structure determination inside the hydrated lipid bilayer, where the protein is correctly folded and stable for a long time. Sample preparation for solid-state NMR is therefore different than for solutionstate NMR. If the protein is expressed cell-free in the presence of detergent, it needs to be purified, renatured and reconstituted in a membrane bilayer. Compared to a protein expressed in a cell, the solubilization step is avoided and the purification step is greatly simplified and accelerated, reducing the chances of spurious proteolysis. Reconstitution of the protein in a lipid membrane is the price to pay to obtain a sample where the membrane protein is almost certainly in its native state, and where the protein function can be checked and monitored (13, 14). All these steps are quite flexible, and they are described below for the expression of the mechanosensitive channel MscL. This protocol is general for membrane proteins, although details such as the nature or the concentration of detergent can vary from one protein to the next. Cell-free expression in presence of liposomes or nanodiscs is an alternative sample preparation approach that has been developed more recently and that has yet to be proven general for membrane

6

Cell-Free Membrane Protein Expression for Solid-State NMR

87

proteins (15–17). When possible, it presents many advantages for solid-state NMR studies. First and foremost, the protein is already in its final state, saving time and avoiding many biochemical steps where the protein may be partially lost or inactivated. Second, the protein is purified simply by centrifugation, alleviating the necessity of adding an affinity tag. In addition, no detergent is used that can interfere and that needs to be removed. Importantly, the protein is in its native environment at all times, where it is well folded, functional and stable (18). Finally, this approach is quite flexible, allowing for a variety of labeling strategies (19). In our hands, cell-free expression of MscL in the presence of liposomes has proven efficient (3, 14, 18, 19) and we describe it below, as it is the best method so far for providing a solid-state NMR sample. The protocols presented here make use of commercial kits for cell-free expression, which we found advantageous for its convenience, reliability, for saving the time and manpower to make the lysate, and also for managing the stocks. Cell lysates can also be prepared, following published protocols from various cell types such as bacteria, wheat germ, or others (11, 20). We describe below an optimized procedure for expressing the membrane protein MscL, using the commercial Roche/5Prime continuous exchange vessel and kits, with an Escherichia coli extract. Since, for each sample preparation protocol, it is necessary to check for protein integrity, we also describe several tests that should be performed to assess the quality of the sample.

2. Materials 2.1. Cell-Free Expression of the Mechanosensitive Channel MscL in Detergent Micelles

All solutions are prepared with autoclaved nuclease-free Milli-Q water (see Note 1). 1. Thermoregulated shaker for the cell-free reaction vessel. 2. Nuclease-free 50-mL tubes. 3. Nuclease-free 1.5-mL tubes. 4. Nuclease-free glass pipettes. 5. Nuclease-free pipette tips (0–10, 10–200, 200–1,000 μL, and 1–5 mL), autoclaved at 121°C for 20 min. 6. pIVEX-2.3-mscL plasmid (3). Encodes the E. coli MscL C-terminally fused to a His6 tag under the control of the T7 promoter. Dissolve in pure nuclease-free water and store aliquoted at −20°C, at 0.5 μg/μL. 7. RTS 9000 cell-free expression kit (Roche/5Prime). Contains lyophilized E. coli lysate, reaction mix, feeding mix, reconstitution buffer, continuous exchange reaction vessel, and a syringe (see Fig. 1, Notes 2–5). Store at −80°C.

88

A. Abdine et al.

Fig. 1. Roche/5Prime RTS 9000 reaction vessel. The reaction compartment (10 mL) and the feeding compartment (100 mL) are each accessible through two screws. The reaction compartment contains the cell lysate, plasmid DNA, detergent or preformed liposomes and is where the coupled transcription/translation reaction takes place. The separate feeding compartment provides additional ions, energy substrates, nucleotides and amino acids, through a semipermeable membrane (MW cutoff 10 kDa). Simultaneously, by-products that may inhibit the reaction are diluted through the same membrane into the feeding compartment. This continuous exchange allows cell-free expression to last for up to 24 h.

8. Unlabeled amino acids (powder), store at −20°C. 9.

13

C/15N labeled amino acids (powder), store at −20°C.

10. Dithiothreitol (DTT): Prepare 40 mM stock, store at −20°C. 11. 20% (w/v) Triton X-100: Prepare 200 mL, stir until homogeneous, store at 4°C for up to 6 months. 12. Unlabeled amino acid solutions: Prepare 168 mM solutions of each amino acid except for leucine (140 mM) and the labeled amino acids, Ile and Thr, with the appropriate solution (see Note 6). The amount of amino acid to be incorporated is calculated in Subheading 3.1 and is indicated in Table 1. Transfer the appropriate volume of each unlabeled amino acid to a 1.5-mL vial, and sonicate until the solution looks clear. Transfer the individual solutions to a 50-mL tube. Add Tyr and Leu last to avoid precipitation; prepare fresh before using and keep on ice. 13. Labeled amino acid solutions, Ile and Thr: Dissolve 23 mg of Ile and 3.9 mg of Thr in 1.0 and 0.20 mL of reconstitution buffer, respectively (~168 mM each), into 1.5-mL tubes; prepare fresh before using and keep on ice (see Subheading 3.1, Table 1 and Note 7). 14. FPLC purification system. 15. 1 M NaOH: To adjust pH, store at room temperature. 16. 100 mM NiSO4: Store at room temperature. 17. 20% Ethanol: Store at room temperature for up to a month. 18. 4-(2-Aminoethyl) benzene sulfonyl fluoride hydrochloride (AEBSF): 100 mM stock solution in water, store at 4°C for up to 6 months.

6

Cell-Free Membrane Protein Expression for Solid-State NMR

89

Table 1 Preparation of amino acid solutions for MscL cell-free expression in a volume V = 110 mL (see Subheading 3.1 and Fig. 1) Amino acid

MWi (g/mol)

n

Amount (mg)

Volume (mL)

Ala (A)

89

15

15

0.98

Arg (R)

174

6

11

0.39

Asn (N)

132

6

Asp (D)

133

7

10

0.46

Cys (C)

121

0

0

0.10

Gln (Q)

146

4

6.4

0.26

Glu (E)

147

8

13

0.52

Gly (G)

75

13

11

0.85

His (H)

155

7

12

0.46

Ile (I)

131

16

23

1.0

Leu (L)

131

13

19

1.0

Lys (K)

146

9

14

0.59

Met (M)

149

5

Phe (F)

165

10

Pro (P)

115

7

8.9

0.46

Ser (S)

105

4

4.6

0.26

Thr (T)

119

3

3.9

0.20

Trp (W)

204

0

0

0.10

Tyr (Y)

181

1

2.0

0.10

Val (V)

117

13

8.7

8.2 18

17

0.39

0.33 0.65

0.85

n is the number of each amino acid type, of molecular weight MWi, in the protein sequence. The weight of each amino acid is (n × MWi × 1.1 × 10−2), expressed in mg. Since each amino acid is solubilized at 168 mM, except for leucine, at 140 mM, the corresponding volume of each amino acid solution is (n × 11)/168, except for leucine where it is (n × 11)/140, expressed in mL

19. Chelating column (5 mL): Stored in 20% ethanol at 4°C. 20. FPLC buffer A1: 50 mM NaH2PO4–NaOH (7.1 g/L), pH 8, 300 mM NaCl (17.5 g/L), 10 mM imidazole (680 mg/L), 4.0% Triton X-100 (40 g/L), store at 4°C. 21. FPLC buffer A2: 50 mM NaH2PO4–NaOH (7.1 g/L), pH 8, 300 mM NaCl (17.5 g/L), 10 mM imidazole (680 mg/L), 0.2% Triton X-100 (2 g/L), store at 4°C.

90

A. Abdine et al.

22. FPLC buffer B: 50 mM NaH2PO4–NaOH (7.1 g/L), pH 8, 300 mM NaCl (17.5 g/L), 500 mM imidazole (34 g/L), 0.2% Triton X-100 (2 g/L), store at 4°C. 23. FPLC sample loading buffer: 50 mM NaH2PO4–NaOH (7.1 g/L), pH 8, 300 mM NaCl (17.5 g/L), 10 mM imidazole (680 mg/L), 1% Triton X-100 (10 g/L), store at 4°C. 24. 10× Dialysis buffer: 0.1 M HEPES–KOH, pH 7.5, 1 M KCl, and 2% (w/v) Triton X-100, store at 4°C for up to 6 months. 25. 1× Dialysis buffer: Prepare 2 L by diluting 200 mL of 10× buffer in 1,750 mL of water, adjusting the pH to 7.5 using 4 M KOH, and adding water to 2 L final volume. Store at 4°C for up to 6 months. 26. 2 dialysis cassettes with a 10 kDa MW cutoff. 27. Methanol. 28. DOPC: 1,2-dioleoyl-sn-glycero-3-phosphocholine powder, store at −20°C. 29. 4 M KOH: To adjust pH, store at room temperature. 30. 0.8% Triton X-100: Prepare 10 mL, dissolve and stir until homogeneous, store at 4°C for up to 6 months. 31. Wet polystyrene beads: Wash 5 g of 300–1,200 μm polystyrene beads with 25 mL of pure methanol and then wash four times with 25 mL of Milli-Q water. The beads are kept in water at 4°C and are stable for months, provided a weekly renewal of Milli-Q water. 32. HEPES–KCl buffer: 10 mM HEPES–KOH, pH 7.5, 100 mM KCl, store at 4°C for up to 6 months. 33. HEPES solutions for solubilizing Tyr, Trp, and Phe (Subheading 2.1, see Notes 6 and 7): 60 mM HEPES, pH 13 for Tyr, pH 1 for Trp, and pH 7.5 for Phe. Adjust the pH using KOH or HCl. Keep on ice. 2.2. Cell-Free Expression of the Mechanosensitive Channel MscL in Liposomes

All solutions are prepared with autoclaved nuclease-free Milli-Q water (see Note 1). 1. Thermoregulated shaker for the cell-free reaction vessel. 2. Mini-Extruder with two 1-mL gastight microsyringes and 0.1-μm cutoff polycarbonate membranes. 3. Nuclease-free 50-mL tubes. 4. Nuclease-free 1.5-mL tubes. 5. Nuclease-free glass pipettes. 6. Nuclease-free pipette tips (0–10, 10–200, 200–1000 µL and 1–5 mL). Autoclave at 121°C for 20 min.

6

Cell-Free Membrane Protein Expression for Solid-State NMR

91

7. pIVEX-2.3-mscL plasmid: For protein expression in liposomes, an affinity tag such as a polyhistidine stretch is preferable but not mandatory. 8. RTS 9000 cell-free expression kit (Roche/5Prime). Contains lyophilized E. coli lysate, reaction mix, feeding mix, reconstitution buffer, a continuous exchange reaction vessel, and a syringe (see Fig. 1, Notes 2–5), store at −80°C. 9. Unlabeled amino acids (powder), store at −20°C. 10.

13

C/15N labeled amino acids (powder), store at −20°C.

11. DTT (dithiothreitol): Prepare 40 mM stock, store at −20°C. 12. 4 M KOH: To adjust pH, store at room temperature. 13. AEBSF (Subheading 2.1). 14. DOPC. 15. Chloroform. 16. HEPES–KCl buffer (Subheading 2.1). 17. DOPC liposome preparation: Prepare a chloroform solution of 10 mg/mL of DOPC (100 mg of DOPC in 10 mL of dry chloroform). Dry the lipids and then resuspend with RTS reconstitution buffer to obtain a 20 mg/mL aqueous solution. Sonicate 2.5 mL of the solution for 5 min, and then extrude it 13 times with a 0.1-μm filter, to obtain a liposome solution containing 40–50 mg of lipids, and store at 4°C. This is the amount of lipids that is expected in the final sample. DOPC can be replaced by other lipids or lipid mixtures, at a higher or lower concentration. 18. HEPES solutions for solubilizing Tyr, Trp, and Phe (Subheading 2.1). 19. Unlabeled amino acid solutions: Prepare 168 mM solutions of each amino acid except for leucine (140 mM) and the labeled amino acids (item 20), with the appropriate solution (see Note 6). The amount of amino acid to be incorporated is calculated in Subheading 3.1 and is indicated in Table 1. The appropriate volume of each unlabeled amino acid is transferred to a 1.5-mL vial, and sonicated until the solution looks clear. The vials are transferred into a 50-mL tube. Tyr and Leu are added last, to avoid precipitation; prepare fresh before using and keep on ice. 20. Labeled amino acid solutions, Arg, Ile, Pro, Met, and Phe: Weigh 11 mg of Arg, 23 mg of Ile, and 8.9 mg of Pro and dissolve in 0.39, 1.0, and 0.46 mL of reconstitution buffer, respectively (~168 mM). Weigh 8.2 mg of Met and dissolve in 0.33 mL of reconstitution buffer plus 50 μL of 40 mM DTT. Weigh 18 mg of Phe and dissolve in 0.65 mL of the appropriate HEPES solution (Subheading 2.1), and sonicate for 1 min. Prepare each amino acid solution fresh before using and keep on ice (see Subheading 3.1, Table 1 and Note 7).

92

A. Abdine et al.

2.3. Sample Characterization

All solutions described should be prepared using Milli-Q water and stored at room temperature unless stated otherwise. 1. FA diluted solution: 37% formaldehyde in water, store at 4°C. 2. Dimethyl sulfoxide (DMSO). 3. 1 M NaOH: To adjust pH. 4. 1 M HCl: To adjust pH. 5. 1 M Tris base, pH 8. 6. SDS running buffer: 100 mM Tris base, 100 mM HEPES, 0.1% SDS, adjust to pH 8 using NaOH or HCl. 7. Staining buffer: 0.5% Coomassie Brilliant Blue R-250 dye, 50% ethanol and 10% acetic acid. 8. Destaining buffer: 20% ethanol and 10% acetic acid. 9. 2× SDS loading buffer: 0.2 M Tris base, pH 8, 8% SDS, 40% glycerol, 0.4% bromophenol blue, 0.4 M DTT, adjust pH 8 using 1 M NaOH or 1 M HCl. 10. 1× SDS loading buffer: dilute 1 mL of 2× SDS loading buffer with 1 mL of water. 11. DSS solution: 50 mM disuccinimidylsuberate in DMSO. 12. HEPES–KCl buffer (see Subheading 2.1). 13. Polyvinylidene fluoride (PVDF) membranes. 14. TBS-Tween buffer 1: 50 mM Tris base, pH 7.4, 150 mM NaCl, 0.05% Tween-20. 15. Monoclonal anti-histidineperoxidase-conjugate antibody solution (for the dot blot): Dilute 2,000× from the stock solution with TBS-Tween buffer 1, prepare fresh. 16. Diaminobenzidine solution: Dissolve one tablet of diaminobenzidine and one tablet of urea hydrogen peroxide in 5 mL of water, prepare fresh. 17. Western-blotting detection kit. 18. Nonfat dry milk (powder). 19. Chemiluminescence reagents (H2O2 and luminol), store at 4°C. 20. Towbin buffer: 25 mM Tris base, 192 mM glycine, 20% isopropanol, and 0.1% SDS. Store without isopropanol at 4°C, add isopropanol just before use. 21. Ponceau S solution: 0.1% Ponceau S, 5% acetic acid. 22. TBS-Tween buffer 2: TBS-Tween buffer 1 with 5% (w/v) nonfat dry milk, freshly made. 23. Mouse anti-histidine antibody solution (for the Western-blot): Dilute 5,000× from the stock solution with TBS-Tween buffer 1, prepare fresh. 24. Anti-mouseperoxidase-coupled antibody solution (for the Western-blot): Dilute 10,000× from the stock solution with TBS-tween buffer 1, prepare fresh.

6

Cell-Free Membrane Protein Expression for Solid-State NMR

93

25. Micro BCA Protein Assay kit (Pierce). 26. Nuclease-free 1.5-mL tubes. 27. Bovine serum albumine (BSA): 1 mg/mL stock solution. 28. Solubilization buffer: 0.2% Triton X-100, solubilized in HEPES–KCl buffer, store at 4°C for up to 6 months. 29. Acetone, stored at −20°C. 30. Nitric acid solution: 1%, store at room temperature in a safety cabinet. Handle under the hood. 31. Perchloric acid solution: 70%, store at room temperature in a safety cabinet. Handle under the hood. 32. Ammonium molybdate solution: Dissolve 2.5 g in 100 mL of water, store at 4°C. 33. Ascorbic acid solution: Dissolve 10 g in 100 mL of water, prepare fresh, and do not keep for more than 1 week at 4°C. 34. Phosphate stock solution: Dissolve 10 mg of KH2PO4 in 100 mL of water and store at 4°C. 35. Nuclease-free 3-mL ultracentrifugation tubes. 36. Sucrose. 37. Sucrose solutions: 10, 20, and 45% (w/v) sucrose, solubilized in HEPES–KCl buffer, store at 4°C. 38. Nitrocellulose membrane (for Western blotting). 39. X-ray film (to develop the proteoliposomes must be mixed with sucrose Western blot). 40. Film developer. 2.4. NMR Sample Preparation

Four millimeter diameter rotors for high-resolution magic-angle spinning solid-state NMR (see Fig. 2).

Fig. 2. Four millimeter diameter rotors for high-resolution magic-angle spinning solidstate NMR. The sample (approximately 50 μL) is contained in the center of the zirconium rotor, thanks to the Teflon insert bottom and top, and is kept tightly in place by the top screw and the Kel-F cap.

94

A. Abdine et al.

3. Methods The methods outlined below describe the following: (1) calculation of the amount of incorporated amino acids, (2) cell-free expression of the mechanosensitive channel MscL in detergent micelles, purification of protein/detergent micelles, protein reconstitution into liposomes, (3) cell-free expression directly into liposomes, (4) sample characterization, and (5) NMR sample preparation. 3.1. Cell-Free Expression of the Mechanosensitive Channel MscL in Detergent Micelles

The method below describes the production of an MscL sample for solid-state NMR, using a cell-free protein expression system in detergent micelles. Although any labeling scheme could be performed, the sample described below was labeled on isoleucines and threonines, using 13C/15N-labeled amino acids, and was subsequently used in solid-state NMR experiments (14).

3.1.1. Day 1: Cell-Free Protein Expression in Detergent Micelles

1. Calculate the amount of amino acids to be incorporated during protein expression. The concentration of each amino acid in the protein cell-free expression system (V = 110 mL, see Fig. 1) is usually ~1 mM. For a given protein, each amino acid concentration depends on its occurrence, n, in the protein sequence. In our case, we have obtained good results with a final concentration of each amino acid equal to n/10, expressed in mM. The corresponding weight of each amino acid, of molecular weight MWi, is therefore (n × V × MWi/10), expressed in mg. The corresponding volume of each amino acid solution of concentration Ci, where Ci is 168 or 140 μM (Subheading 2.1), is (n × V/10)/Ci, expressed in mL. The weights and final volumes calculated for this protocol are summarized in Table 1. 2. Thaw the DTT, RTS reconstitution buffer, and MscL plasmid at room temperature. 3. Thaw the other components of the RTS kit (E. coli lysate, reaction mix, and feeding mix) on ice. 4. Prepare the unlabeled amino acid solutions (see Subheading 2.1). 5. Prepare the labeled amino acid solutions (see Subheading 2.1). 6. Add the labeled amino acid solution to the unlabeled amino acid solution. 7. Add 3 mL of 40 mM DTT, sonicate (80 W for 5 min or until clear), and store on ice. 8. Reconstitute the lyophilized E. coli lysate in 5.2 mL of reconstitution buffer. Shake the bottle gently (see Note 8). 9. Reconstitute the lyophilized reaction mix in 2.2 mL of reconstitution buffer. Shake the bottle gently (see Note 8).

6

Cell-Free Membrane Protein Expression for Solid-State NMR

95

10. Reconstitute the lyophilized feeding mix in 80 mL of reconstitution buffer. Shake the bottle gently (see Note 8). 11. Prepare the feeding solution by adding 26 mL of the reconstituted amino acid solutions (mixture of unlabeled and labeled amino acid solutions) and 3 mL of 40 mM DTT to the feeding mix (80 mL). 12. Prepare the reaction solution by adding the reconstituted reaction mix (2.2 mL), 2.7 mL of the reconstituted amino acid solution and 0.3 mL of 40 mM DTT to the reconstituted E. coli lysate (5.2 mL). Remove a 20 μL aliquot of this solution for later comparison on gel electrophoresis. 13. Add 300 μL of the MscL plasmid (~150 μg) to the reaction solution. 14. Add 200 μL of 20% Triton X-100 solution to the reaction solution. 15. Add 2.2 mL of 20% Triton X-100 solution to the feeding solution. 16. Open both screws of the reaction compartment and fill the reaction compartment with the reaction solution using a nuclease-free pipette (see Fig. 1). Remove air bubbles by tapping the vessel lightly. 17. Open both screws of the feeding compartment and fill the feeding compartment with the feeding solution, using the provided syringe (see Fig. 1). Remove air bubbles by tapping the vessel lightly. 18. Insert the reaction vessel into a thermoregulated shaker. 19. Set the shaking speed to 800 rpm. 20. Set the temperature to 30°C. 21. Incubate the reaction for 22 h. 3.1.2. Day 2: Purification of Detergent/Protein Micelles

1. After 22 h, stop the cell-free protein expression. Remove a 20 μL aliquot of the reaction run-on solution for gel electrophoresis. 2. Remove the reaction run-on mix using a pipette, add 100 μL of the AEBSF solution to prevent protease digestion, and stir at room temperature for 15 min (see Note 9). 3. Dilute the sample to a final volume of 50 mL using FPLC buffer A1 and centrifuge at 10,000 × g for 15 min at 4°C. 4. Connect the chelating column to the FPLC system (see Note 10). Pass 4–5 column volumes (CV) of distilled water through the column, to wash away the ethanol solution in which it has been kept. 5. Degas the FPLC buffers for 10 min in an ultrasonic bath. Wash the pump inlets with the degassed solutions. Wash the system with FPLC buffer A1 until the UV and conductivity baselines are stable.

96

A. Abdine et al.

6. At the end of the sample centrifugation (step 3), dilute the supernatant (reaction run-on mix) with FPLC sample loading buffer to a final volume of 100 mL (final dilution ~10×). 7. Filter this solution through a 0.45-μm filter. 8. Resuspend the pellet with 5 mL of FPLC buffer A1 and remove a 20 μL aliquot for gel electrophoresis. 9. Introduce half (50 mL) of the diluted reaction run-on mix into the column (see Note 11). Collect the flow-through in the first 50-mL tube. 10. Wash off the nonspecifically bound material by raising the concentration of imidazole to 100 mM, by mixing 80% of FPLC buffer A1 with 20% of FPLC buffer B (6 CV total). The concentration of detergent remains high (around 3%). Collect the flow-through in the second 50-mL tube. 11. To elute the protein, switch FPLC buffer A1 to buffer A2, to reduce the detergent concentration to 0.2%. Collect the flowthrough in the second 50-mL tube. 12. Increase the imidazole concentration to 200 mM by mixing 60% of solution A2 with 40% of solution B (2 CV total), and collect eluant fractions of 2 mL each (see Note 12). 13. Increase the imidazole concentration to 500 mM by switching to 100% FPLC buffer B (6 CV total) and collect eluant fractions of 2 mL each. 14. Wash the column with FPLC buffer A1 (10 CV or until the UV and conductivity baselines are stable), then repeat steps 9–13 with the remaining 50 mL (second half of the diluted reaction run-on mix, see Note 11). 15. At the end of the FPLC protein purification run (see Note 10), the potentially interesting fractions are separated into approximately 40 tubes, whereas the flow-though is contained in two 50-mL tubes. Remove a 20 μL aliquot from each of these 42 tubes, for gel electrophoresis. 16. Determine which fractions contain the histidine-tagged protein by performing a dot blot (see Subheading 3.3.4). 17. Wash the two dialysis cassettes in 1× dialysis buffer. 18. For each half of the solution, pool the fractions containing the protein into a dialysis cassette. 19. Dialyze against 1 L of 1× dialysis buffer, overnight at 4°C. 3.1.3. Day 3: Sample Characterization and Reconstitution into Liposomes

1. Dialyze a second time against 1 L of 1× dialysis buffer for 2 h at 4°C. Remove a 20 μL aliquot for gel electrophoresis. 2. Assess the cell-free expression level, FPLC purification efficiency, and oligomeric state by SDS-PAGE (see Subheadings 3.3.1 and 3.3.3).

6

Cell-Free Membrane Protein Expression for Solid-State NMR

97

3. After SDS-PAGE, wash the gel in water and perform a Westernblot transfer (see Subheading 3.3.5). 4. Estimate the protein concentration. Since MscL does not contain any tryptophan it has to be quantified by using the BCA method (see Subheading 3.3.6 and Note 13). 5. BCA quantification of our MscL preparation indicated that each half of the solution contains approximately 4.8 mg of protein in a 10 mL volume that will be reconstituted in 19.2 mg of DOPC liposomes (protein–lipid ratio of 1:4). DOPC can be replaced by other lipids or lipid mixtures, at a higher or lower concentration (21). Lipids can also be replaced by other mixtures to reconstitute the protein into bicelles or nanodiscs (22, 23). 6. Split the solutions into four fractions of 5 mL, each containing 2.4 mg of protein and 10 mg of Triton X-100. To each fraction, add 9.6 mg of DOPC solubilized in 2.5 mL of the 0.8% Triton X-100 solution, such that the final lipid–detergent ratio is 1:3. Stir slowly for 15 min at room temperature. 7. To each fraction of 7.5 mL add 300 mg of wet polystyrene beads (detergent–bead ratio is 1:10) and put the four tubes on a rotator, to remove the detergent by adsorption onto the beads. Incubate overnight at 4°C. For alternative methods (21, 24), see Note 14. 3.1.4. Day 4: Preparation of the Sample for Solid-State NMR

1. Add another 300 mg of wet polystyrene beads to each tube. Incubate for another 2 h, at 4°C. 2. After 2 h, set the tubes vertically for 5 min and discard the pellet and the beads. 3. Divide the supernatant into four tubes and centrifuge at 100,000 × g for 30 min at 4°C. Remove a 20 μL aliquot of one supernatant for gel electrophoresis, discard the rest and resuspend the pellets with 1 mL of HEPES–KCl buffer for each pellet. Split the new resuspended pellets into two tubes and centrifuge at 100,000 × g for 30 min at 4°C. Repeat one more time so that the entire sample fits into a single tube. Remove a final 20 μL aliquot of the supernatant for gel electrophoresis. Dry the final pellet under argon and store it at −20°C. 4. Assess the reconstitution by SDS-PAGE (see Subheading 3.3.1), and by proteoliposome density characterization on a sucrose gradient (see Subheading 3.3.9). Also, estimate the lipid and water content (see Subheadings 3.3.8 and 3.3.10, and Note 15). 5. The average quantity of protein obtained with the RTS 9000 kit is 10 mg in 10 mL of reaction mix, which is generally sufficient to prepare two or three NMR samples. Transfer ~30 mg of pelleted sample to the NMR rotor (see Subheading 3.4). 6. Store the remaining pellet at −20°C.

98

A. Abdine et al.

3.2. Cell-Free Expression of the Mechanosensitive Channel MscL in Liposomes

The method below describes the production of an MscL sample for solid-state NMR, using a cell-free protein expression system directly into lipid vesicles. Although any labeling scheme could be performed, the sample described below was labeled on arginines, isoleucines, methionines, phenylalanines and prolines, using 13 C/15N-labeled amino acids, and was subsequently used in solidstate NMR experiments (19). Lipids used were synthetic DOPC, but other lipids or complex mixtures can also be used, such as asolectin (18) or nanodiscs (17).

3.2.1. Day 1: Cell-Free Protein Expression

1. Thaw the DTT solution, RTS reconstitution buffer and MscL plasmid at room temperature. 2. Thaw the other components of the RTS kit (E. coli lysate, reaction mix, and feeding mix) on ice. 3. Prepare the DOPC liposomes (Subheading 2.2). 4. Prepare the HEPES solutions (Subheading 2.1). 5. Prepare the unlabeled amino acid solutions (Subheading 2.1). 6. Prepare the labeled amino acid solutions (Subheading 2.1). 7. Add the labeled amino acid solutions to the unlabeled amino acid solutions. 8. Add 3 mL of 40 mM DTT, sonicate (80 W for 5 min or until clear), and store on ice. 9. Reconstitute the lyophilized E. coli lysate in 2.7 mL of reconstitution buffer. Shake the bottle gently (see Note 8). 10. Add the liposome solution (2.5 mL) to the E. coli lysate once the latter is completely dissolved. 11. Reconstitute the lyophilized reaction mix in 2.2 mL of reconstitution buffer. Shake the bottle gently (see Note 8). 12. Reconstitute the lyophilized feeding mix in 80 mL of reconstitution buffer. Shake the bottle gently (see Note 8). 13. Prepare the feeding solution by adding 26 mL of the reconstituted amino acid solution (mixture of unlabeled and labeled amino acid solutions) and 3 mL of 40 mM DTT to the feeding mix solution (80 mL). 14. Prepare the reaction solution by adding the reconstituted reaction mix (2.2 mL), 2.7 mL of the reconstituted amino acid solution and 0.3 mL of 40 mM DTT to the reconstituted E. coli lysate with the liposomes (5.2 mL). Take a 20 μL aliquot of this reaction solution for later comparison on gel electrophoresis. 15. Add 300 μL of the MscL plasmid (~150 μg) to the reaction solution. 16. Open both screws of the reaction compartment and fill the reaction compartment with the reaction solution using a nuclease-free pipette (see Fig. 1). Remove any air bubbles by tapping the vessel lightly.

6

Cell-Free Membrane Protein Expression for Solid-State NMR

99

17. Open both screws of the feeding compartment and fill the feeding compartment with the feeding solution, using the provided syringe (see Fig. 1). Remove any air bubbles by tapping the vessel lightly. 18. Insert the reaction vessel into a thermoregulated shaker. 19. Set the shaking speed to 800 rpm. 20. Set the temperature to 30°C. 21. Incubate the reaction for 22 h. 3.2.2. Day 2: Preparation of the Sample for Solid-State NMR

1. After 22 h, stop the cell-free protein expression. Remove a 20 μL aliquot of the reaction run-on solution for gel electrophoresis. 2. Remove the reaction run-on mix using a pipette, add 100 μL of the AEBSF solution to prevent protease digestion, and stir at room temperature for 15 min (see Note 9). 3. Split the sample into six tubes and centrifuge at 100,000 × g for 30 min at 4°C. Remove a 20 μL aliquot of one supernatant for gel electrophoresis, discard the rest and resuspend each pellet with 1 mL of HEPES–KCl buffer. Split the resuspended pellets into three tubes and centrifuge at 100,000 × g for 30 min at 4°C. Repeat one more time so that the entire sample fits into a single tube. Remove a final 20 μL aliquot of the supernatant for gel electrophoresis. Dry the final pellet under argon and store it at −20°C. 4. Assess cell-free expression efficiency and oligomeric state by SDS-PAGE (see Subheadings 3.3.2 and 3.3.3). At this point, the expressed protein is generally almost pure. 5. If the protein was expressed with a polyhistidine tag, after SDSPAGE, wash the gel in water and perform a Western-blot transfer (see Subheading 3.3.5). 6. Estimate the protein and lipid concentration using the BCA and Rouser methods respectively (see Subheadings 3.3.7 and 3.3.8, and Note 13). 7. Characterize the proteoliposome density on a sucrose gradient (see Subheading 3.3.9), and estimate the water content (see Subheading 3.3.10 and Note 15). 8. The average quantity of protein obtained with the RTS 9000 kit is 10 mg in 10 mL of reaction mix, which is generally sufficient to prepare two or three NMR samples. About 30 mg of pelleted sample is transferred into the NMR rotor (see Subheading 3.4). 9. Store the remaining pellet at −20°C.

100

A. Abdine et al.

3.3. Sample Characterization 3.3.1. Electrophoresis of Protein Expressed in Detergent Micelles

1. Perform SDS-PAGE on the following samples: (a) Aliquots from the reaction mix before and after the cell-free expression reaction, to check for expression of the target protein. (b) Aliquots from the centrifugation pellet and supernatant, to check whether the protein was totally solubilized in detergent micelles or if there are some aggregates. (c) Aliquots from the purification flow-through and the purified fractions. (d) Analyze the incorporation of the protein by comparing aliquots of the protein-detergent micelles with the proteoliposomes, after the reconstitution step. (e) Aliquots of the supernatant of all the wash steps. (f) Also check the integrity of the purified protein after dialysis. 2. Dilute all SDS PAGE samples 1:1 with 2× SDS loading buffer. 3. Heat the samples to 90°C for 5 min. 4. Cool the samples and load on a gel. 5. Run the gel using SDS running buffer. 6. Wash the gel in water and stain/destain the gel using staining/ destaining buffers, respectively.

3.3.2. Electrophoresis of Protein Expressed in Liposomes

1. Perform SDS-PAGE on the following samples: (a) Aliquots from the reaction mix before and after the cellfree expression reaction, to check for expression of the target protein. (b) Also check the centrifugation supernatants. 2. Follow steps 2–6 in Subheading 3.3.1.

3.3.3. Electrophoresis for Oligomeric State Characterization

If the protein is an oligomer (like MscL), assess its integrity by cross-linking the protein with different free primary amine targeting reagents, like formaldehyde (FA) or disuccinimidylsuberate (DSS) (3, 25). NB: the protein buffer must not contain primary amines (such as in Tris). 1. Mix 5 μg of purified MscL, either in detergent or liposomes, in 10 μL of HEPES–KCl buffer and 0.54 μL of the FA diluted solution. 2. Mix 5 μg of purified MscL, either in detergent or liposomes, in 10 μL of HEPES–KCl buffer and 0.2 μL of 50 mM DSS. 3. Incubate both samples, without shaking, for 30 min at room temperature.

6

Cell-Free Membrane Protein Expression for Solid-State NMR

101

Fig. 3. Coomassie blue stained SDS-PAGE of MscL: (1) purified in detergent, (2) crosslinked with formaldehyde, (3) cross-linked with disuccinimidylsuberate. Formaldehyde generates five protein bands on the gel, corresponding to various oligomeric forms, from monomers to pentamers, while disuccinimidylsuberate generates mostly pentamers, confirming the pentameric nature of Escherichia coli MscL (3, 25).

4. Stop the cross-linking reactions by adding 2 μL of 1 M Tris base, pH 8, to each sample. Gently mix and incubate for 10 min at room temperature. 5. Add 3 μL of 2× SDS loading buffer to each sample and mix. 6. Heat the FA sample to 60°C and the DSS sample to 90°C, for 5 min in a water bath. 7. Cool the samples and load on a gel. 8. After migration, wash the gel in water and stain/destain the gel using staining/destaining buffers, respectively (see Fig. 3) or perform a Western-blot transfer (see Subheading 3.3.5). 3.3.4. Dot Blot

1. Deposit 2 μL of each fraction of purified protein directly onto a PVDF membrane. 2. Allow the protein solution to air-dry. 3. Prepare the TBS-Tween buffers 1 and 2. 4. Prepare the monocolonal antibody solution. 5. When the protein spots are completely dry, incubate the membrane in the TBS-Tween buffer 2 for 30 min. 6. Wash the membrane twice for 5 min with the TBS-Tween buffer 1.

102

A. Abdine et al.

7. Incubate the membrane with the monoclonal antibody solution for 1 h. 8. Wash the membrane twice for 5 min with the TBS-Tween buffer 1. 9. Prepare the diaminobenzidine solution. 10. Dry the membrane on a tissue. 11. Incubate the membrane with the diaminobenzidine solution for a few seconds, until the spots are colored. 12. Dry the membrane on a tissue. The color remains on the spots containing the histidine-tagged protein, while it vanishes on the other spots. 3.3.5. Semidry Western-Blot

1. Prepare the Towbin buffer. 2. Prepare the transfer sandwich by superimposing two filter papers, the gel, the nitrocellulose membrane and another two filter papers in the Towbin buffer. 3. Place the transfer sandwich on the anode plate and clip the cathode plate on top. 4. Clamp the potential at 25 V and blot for 20 min. 5. Prepare TBS-Tween buffers 1 and 2, mouse anti-histidine antibody, and anti-mouse peroxidase-coupled antibody solutions. 6. Assess the sample transfer and relative concentration by staining the membrane with Ponceau S solution. This is a reversible stain that can be removed by washing in TBS-Tween buffer 1. 7. After protein transfer and destaining, saturate the nitrocellulose membrane by incubating in TBS-Tween buffer 2 for 30 min. 8. Rinse the membrane quickly with TBS-Tween buffer 1. 9. Incubate for 45 min at room temperature with the mouse antihistidine antibodies solution. 10. Wash the membrane for 4 × 5 min with TBS-Tween buffer 1. 11. Incubate for 30 min at room temperature with the anti-mouse peroxidase-coupled antibody solution. 12. Wash the membrane for 5 × 5 min with TBS-Tween buffer 1. 13. Incubate the membrane for 1 min with 1 mL of each of the chemiluminescence reagents (H2O2 and luminol). 14. In a light tight box, expose X-ray films to the membrane for varying times, usually between 30 s and 2 min, and develop the films in the photo developer.

6 3.3.6. Protein Quantification in Detergents

Cell-Free Membrane Protein Expression for Solid-State NMR

103

Protein concentration is measured using the BCA method (26, 27), using the test tube procedure of the Micro BCA Protein Assay Kit: 1. Prepare eight BSA standards by diluting the stock solution with the solubilization buffer (the final BSA concentration should fall between 0.5 and 20 μg/mL). Make three replicates of each dilution. Introduce 500 μL of each into 1.5-mL tubes (24 tubes in total) and store at room temperature. 2. Dilute the unknown protein sample with the solubilization buffer to three different concentrations that are expected to be between 1 and 20 μg/mL. Make three replicates of each dilution. Transfer 500 μL of each sample into 1.5-mL tubes (nine tubes in total). 3. Add 500 μL of Micro BCA Working Reagent to each of the 1.5-mL tubes (24 BSA standards and nine unknown proteins) and incubate for 60 min at 60°C. Cool to room temperature. 4. Measure the absorbance at 562 nm. 5. Plot a standard curve based on the absorbance of the BSA samples. 6. Deduce the protein concentration of each unknown protein sample.

3.3.7. Protein Quantification in Proteoliposomes

Lipids need to be removed from the sample by precipitating the proteins using cold acetone, followed by a centrifugation to separate the protein pellet from the supernatant containing the lipids: 1. Prepare the 24 BSA standard tubes (Subheading 3.3.6, step 1). 2. Dilute the unknown protein sample with the solubilization buffer to three different concentrations that are expected to be between 1 and 20 μg/mL. Make three replicates of each dilution. Transfer 500 μL of each sample into 1.5-mL tubes (nine tubes in total). 3. Add 1 mL of cold acetone to each tube of unknown protein. Vortex the tubes and incubate for 60 min at −20°C. Centrifuge at 10,000 × g for 10 min at room temperature. Discard the supernatant. Incubate the tubes for 30 min at room temperature to allow for acetone evaporation. Add 500 μL of the solubilization buffer, and vortex the tubes again. 4. Add 500 μL of Micro BCA Working Reagent to each of the 1.5-mL tubes (24 BSA standards and nine unknown proteins) and incubate for 60 min at 60°C. Cool to room temperature. 5. Measure the absorbance at 562 nm. 6. Plot a standard curve based on the absorbance of the BSA samples. 7. Deduce the protein concentration of each unknown protein sample.

104

A. Abdine et al.

3.3.8. Lipid Quantification in Proteoliposomes

The phospholipid content in the proteoliposome sample is assessed by the Rouser method, which measures the phosphate concentration in the sample (28). Full eye, face, and skin protection is required for this method. 1. Set a heating block at 180°C. 2. Wash the glass tubes in nitric acid solution before use and dry them in an oven. 3. Prepare five phosphate standard samples by diluting the phosphate stock solution into five samples containing 1–5 μg of KH2PO4 per tube. Classically, 5 μg of KH2PO4 give an absorbance of 0.9 at 800 nm. Store at 4°C. 4. Prepare the ascorbic acid solution (Subheading 2.3). 5. Collect three proteoliposomes volumes containing approximately 1–5 μg of lipids. Transfer the samples into clean glass tubes. 6. Add 0.65 mL of the perchloric acid solution and place the tubes in the heated block at 180°C for 30 min of digestion, until the yellow color has disappeared. 7. Add 0.65 mL of the perchloric acid solution per phosphate standard tube (digestion is not necessary). 8. Put all the tubes on ice, and set the heated block at 100°C (alternatively, a boiling water bath can be used). 9. Once cooled, add to the tubes the following: 3.3 mL of water, 0.5 mL of the ammonium molybdate solution, and 0.5 mL of the ascorbic acid solution. Agitate on a vortex after each addition. 10. Put the tubes in the heated block at 100°C for 5 min. 11. Put the tubes on ice for 5 min. 12. Read the absorbance of the cooled samples at 800 nm. 13. Plot a standard curve based on the absorbance of the phosphate standard samples. The exact concentration in the phospholipid samples is calculated based on the standard curve.

3.3.9. Proteoliposome Density Characterization

A discontinuous sucrose flotation gradient analysis (21, 29) can be performed to check that the membrane protein is correctly reconstituted into proteoliposomes. The proteoliposomes can be layered at the bottom (see below) or at the top (see Note 16) of the sucrose layers. 1. After MscL reconstitution, add 120 mg of sucrose to 0.2 mL of the proteoliposome suspension (containing approximately 96 μg of protein) and mix gently by pipetting, until the sucrose has completely dissolved. The final volume is approximately 0.265 mL and the final sucrose concentration is 45%. Adjust the final volume to 0.4 mL with the buffered 45% sucrose solution.

6

Cell-Free Membrane Protein Expression for Solid-State NMR

105

2. Transfer the resulting suspension to a 3-mL ultracentrifuge tube, and keep it vertical in an appropriate rack for the following steps. 3. Carefully deposit 0.7 mL of the buffered 20% sucrose solution, with minimal flow along the tube wall to avoid layers mixing. 4. Repeat with 0.7 mL of the buffered 10% sucrose solution. 5. Ultracentrifuge at 100,000 × g for 1.5 h in a swinging rotor at 18°C. 6. At the end of the centrifugation, carefully remove and transfer the tube into the rack, keeping it vertical. Carefully collect six fractions of 0.3 mL, from top to bottom, into 1.5-mL tubes, and adjust the final volume to 1 mL with water. 7. Wash the bottom of the 3 mL tube with 100 μL of 1× SDS loading buffer and remove a 15 μL aliquot for gel electrophoresis. 8. Centrifuge the 1.5-mL tubes at 16,000 × g for 15 min at room temperature, to pellet the proteoliposomes and remove the sucrose. 9. Resuspend each pellet in 100 μL of 1× SDS loading buffer and take a 15 μL aliquot of each for gel electrophoresis. 10. Analyze the aliquots by gel electrophoresis. For an expected ratio of protein to lipid of 1:4 (w/w), proteoliposomes appear at the interface between 20% and 10% sucrose. Protein-free liposomes lie at the top of the sucrose gradient, while lipid-free protein aggregates lie at the bottom of the sucrose gradient (see Note 17). 3.3.10. Water Quantification in the Sample

3.4. NMR Sample Preparation

Water content is estimated by weighing 10 mg of sample prior to and after 16 h in vacuum. 1. Weigh the empty rotor, cap, insert bottom, top, and screw (see Fig. 2). 2. Introduce the insert bottom into the rotor. 3. Introduce a couple of mg of the pelleted sample into the rotor on the tip of a spatula. Place the rotor into a 1.5-mL tube and centrifuge it at 10,000 × g for 1 min at room temperature. Repeat this until approximately 30 mg of sample has been introduced into the rotor. 4. Place the insert top into the rotor, without the top screw. Clean the upper part the rotor with a precision wiper before capping tightly. At this point, the rotor containing sample should be stored at −20°C until the NMR experiment is performed. 5. Introduce the rotor into the magic-angle spinning NMR probe and into the magnet of the NMR spectrometer. Spin the rotor at ~10 kHz for 15 min.

106

A. Abdine et al.

6. Extract the rotor and open it. Clean the upper part of the rotor again, in case a drop of water has come out. Place the top screw, tighten it, and then cap the rotor tightly. Weigh the full rotor to deduce the final sample mass. A typical sample consists of 3 mg proteins, 12 mg lipids and 15 mg water.

4. Notes 1. All solutions require Milli-Q water, but only solutions used inside the cell-free expression vessel need to be autoclaved and nuclease-free, to make sure that no ribonucleases are present that would damage the RNAs in the lysate. 2. Continuous-exchange cell-free protein expression with an RTS commercial kit from Roche/5Prime is advantageous for its convenience, reliability, for saving time and manpower in preparing the lysate, and for managing the stocks. The thermoregulated shaker, on the contrary, is not necessarily from Roche/5Prime. 3. Cell-free reactions should be performed on a small scale (RTS 100 or analog kits) for optimization studies, including detergents or lipids. Protein yield should be evaluated in a cell-free kit equipped with a continuous-exchange system (e.g., RTS 500), where the yield is usually higher. Once the protocol is optimized, the proteins can then be expressed on larger scales (e.g., RTS 9000). 4. Cell-free kits were stored at −80°C rather than −20°C, which increases the lifespan of the kits to over a year. Freeze–thaw cycles should be avoided. 5. Circular DNA was used for cell-free expression of MscL. Linear PCR templates can also be used, but they require larger amounts of DNA. DNA quality is important for cell-free expression. A plasmid purification procedure with a good yield (typically 100 μg of plasmid DNA per standard MIDI-prep), including an anion-exchange step, was found to be necessary. DNA should be dissolved in either nuclease-free water (our case) or Tris solution (typically 10 mM, at a pH between 8 and 8.5), but not in a buffer containing EDTA, as it would change the free magnesium ion concentration and reduce the protein expression yield. 6. Some amino acids are difficult to solubilize. Sonication or heating at 60°C can improve solubility. Met (M) and Cys (C) have to be supplemented with DTT (4 and 8 mM respectively). Tyr (Y), Trp (W), and Phe (F) have to be solubilized in 60 mM HEPES in nuclease-free water, adjusting the pH to 13, 1, or

6

Cell-Free Membrane Protein Expression for Solid-State NMR

107

7.5 respectively, using KOH or HCl. However, Trp (W), Asp (D), Asn (N), Cys (C), or Tyr (Y) will not dissolve completely and have to be used as suspensions. Alternatively, the Roche/5Prime RTS Amino acid Sampler provides appropriate stock solutions of each individual unlabeled amino acid. 7. Commercial mixtures of labeled amino-acids are also available (30). All labeled amino-acids can be specifically incorporated into the expressed proteins, except for Gln (Q) and Glu (E) when using the Roche/5Prime buffer, which contains large quantities of unlabeled glutamate. Some scrambling is observed with Ser (S), Asp (D), Asn (N), Gln (Q), and Glu (E). Some metabolic degradation can occur with Arg (R), Cys (C), Trp (W), Met (M), Asp (D), and Glu (E) during prolonged incubation. In such a case, an increased amount of amino acids can enhance the protein production. 8. Too vigorous mixing should be avoided while preparing the different lysates, as it may denature proteins or ribosomes in the extracts. 9. AEBSF is added after the expression is complete to prevent protease degradation of the target product. Proteases inhibitors can also be added to the reaction chamber, but they should be tested on a small scale beforehand. 10. If the chelating column is not precharged with Ni ions, before connecting it to the FPLC system, open it at both ends and wash it with 2 CV of distilled water, 1 CV of 100 mM EDTA, pH 8, 3 CV of distilled water, 2 CV of 0.1 M NiSO4 solution, and 5 CV of distilled water. Once the column is charged, an additional washing step with 5 CV of 20% ethanol is necessary before storage at 4°C. At the end of the protein purification run, wash the column with water (5 CV), 0.5 M NaOH (5 CV), water (5 CV) and store in 20% ethanol. 11. The total reaction run-on is expected to contain approximately 10 mg of protein but also detergent and impurities, which may bind to the column as well. Since the binding saturation of the column surface is on the order of 10 mg/mL, the reaction run-on is split in two so as to always remain below the saturation limit. 12. For MscL, the fractions containing the protein are eluted between 200 and 500 mM of imidazole, but this is highly protein dependent. Depending on the histidine-tag accessibility, the protein may elute at lower or higher imidazole concentrations. 13. Protein concentration is usually estimated using UV fluorescence of aromatic amino acids. If the protein is not sufficiently fluorescent, quantification can be performed using the commercial Micro BCA bicinchonic acid protein assay kit (Pierce). Care should be taken if lipids are present in the sample.

108

A. Abdine et al.

14. Instead of using polystyrene beads, detergent extraction can be performed using cyclodextrin inclusion compounds, as described by DeGrip et al. (24), or by dialysis (21). 15. Proteoliposomes can be dehydrated and rehydrated, but care should be taken, since the process may affect protein activity. 16. The discontinuous sucrose flotation gradient analysis to check that the membrane protein is correctly reconstituted can be performed with the proteoliposomes layered either at the top or at the bottom of the sucrose layers, or both (21, 29). If at the top, the proteoliposomes must be mixed with sucrose for a final sucrose concentration of 10%. The rest of the protocol is the same: After layering 0.7 mL of 45% sucrose, then 0.7 mL of 20% sucrose and then 0.7 mL of the proteoliposomes in 10% sucrose, the proteoliposomes usually appear at the interface between 20 and 10% sucrose. 17. If the amount of lipid-free protein aggregates is not negligible, the discontinuous sucrose flotation gradient can also be used as a purification technique by layering the entire reconstituted proteoliposome suspension.

Acknowledgments This work was supported by fellowships from the Ministère de l’Enseignement Supérieur et de la Recherche and the Fondation pour la Recherche Médicale (to A.A.), by the CNRS (UMR 7099 and 8619), the ANR (ANR-06-JCJC0014), the Univ Paris Diderot and the Université Paris-Sud 11. We thank Alexandre Ghazi, Catherine Berrier, Emmanuelle Billon-Denis, and Michiel A. Verhoeven for helping us optimize the protocols presented here. References 1. Lyford, L. K., and Rosenberg, R. L. (1999) Cell-free expression and functional reconstitution of Homo-oligomeric α7 Nicotinic Acetylcholine Receptors into Planar Lipid Bilayers. J. Biol. Chem. 274, 25675–25681. 2. Elbaz, Y., Steiner-Mordoch, S., Danieli, T., and Schuldiner, S. (2004) In vitro synthesis of fully functional EmrE, a multidrug transporter, and study of its oligomeric state. Proc. Natl. Acad. Sci. U.S.A. 101, 1519–1524. 3. Berrier, C., Park, K. H., Abes, S., Bibonne, A., Betton, J. M., and Ghazi, A. (2004) Cell-free synthesis of a functional ion channel in the absence of a membrane and in the presence of detergent. Biochemistry 43, 12585–12591.

4. Klammt, C., Lohr, F., Schafer, B., Haase, W., Dötsch, V., Ruterjans, H., Glaubitz, C., and Bernhard, F. (2004) High level cell-free expression and specific labeling of integral membrane proteins. Eur. J. Biochem. 271, 568–580. 5. Wagner, S., Bader, M. L., Drew, D., and de Gier, J. W. (2006) Rationalizing membrane protein overexpression. Trends Biotechnol. 24, 364–371. 6. Eshaghi, S. (2009) High-throughput expression and detergent screening of integral membrane proteins. Methods Mol. Biol. 498, 265–271. 7. Tate, C. G. (2010) Practical considerations of membrane protein instability during purification and crystallisation. Methods Mol. Biol. 601, 187–203.

6

Cell-Free Membrane Protein Expression for Solid-State NMR

8. Breyton, C., Pucci, B., and Popot, J.-L. (2010) Amphipols and fluorinated surfactants: Two alternatives to detergents for studying membrane proteins in vitro. Methods Mol. Biol. 601, 219–245. 9. White, S. H. (2010) Membrane Proteins of Known Structure. University of California at Irvine. http://blanco.biomol.uci.edu/ Membrane_Proteins_xtal.html Accessed 25 July 2011. 10. Warschawski, D. E. (2010) Membrane Proteins of Known Structure Determined by NMR. Drorlist. http://www.drorlist.com/nmr/MPNMR.html Accessed 25 July 2011. 11. Schneider, B., Junge, F., Shirokov, V. A., Durst, F., Schwarz, D., Dötsch, V., and Bernhard, F. (2010) Membrane protein expression in cellfree systems. Methods Mol. Biol. 601, 165–186. 12. Sobhanifar, S., Reckel, S., Junge, F., Schwarz, D., Kai, L., Karbyshev, M., Löhr, F., Bernhard, F., and Dötsch, V. (2010) Cell-free expression and stable isotope labelling strategies for membrane proteins. J. Biomol. NMR 46, 33–43. 13. Lehner, I., Basting, D., Meyer, B., Haase, W., Manolikas, T., Kaiser, C., Karas, M., and Glaubitz, C. (2008) The key residue for substrate transport (Glu14) in the EmrE dimer is asymmetric. J. Biol. Chem. 283, 3281–3288. 14. Abdine, A., Verhoeven, M. A., Park, K.-H., Ghazi, A., Guittet, E., Berrier, C., Van Heijenoort, C., and Warschawski, D. E. (2010) Structural study of the membrane protein MscL using cell-free expression and solid-state NMR. J. Magn. Reson. 204, 155–159. 15. Kalmbach, R., Chizhov, I., Schumacher, M. C., Friedrich, T., Bamberg, E., and Engelhard, M. (2007) Functional cell-free synthesis of a seven helix membrane protein: in situ insertion of bacteriorhodopsin into liposomes. J. Mol. Biol. 371, 639–648. 16. Marques, B., Liguori, L., Paclet, M. H., VillegasMendéz, A., Rothe, R., Morel, F., Lenormand, J.-L. (2007) Liposome-mediated cellular delivery of active gp91(phox). PLoS One 2, e856. 17. Katzen, F., Fletcher, J. E., Yang, J. P., Kang, D., Peterson, T. C., Cappuccio, J. A., Blanchette, C. D., Sulchek, T., Chromy, B. A., Hoeprich, P. D., Coleman, M. A., and Kudlicki, W. (2008) Insertion of membrane proteins into discoidal membranes using a cell-free protein expression approach. J. Proteome Res. 7, 3535–3542. 18. Berrier, C., Guilvout, I., Bayan, N., Park, K.-H., Mesneau, A., Chami, M., Pugsley, A. P., and Ghazi, A. (2011) Coupled cell-free synthesis and lipid vesicle insertion of a functional oligomeric channel MscL - MscL does not need the insertase YidC for insertion in vitro. Biochim. Biophys. Acta 1808, 41–46.

109

19. Abdine, A., Verhoeven, M. A., and Warschawski, D. E. (2011) Cell-free expression and labeling strategies for a new decade in solid-state NMR. New Biotechnol. 28, 272–276. 20. He, M. (2008) Cell-free protein synthesis: applications in proteomics and biotechnology. New Biotechnol. 25, 126–132. 21. Rigaud, J.-L., and Lévy, D. (2003) Reconstitution of Membrane Proteins into Liposomes. Methods Enzymol. 372, 65–86. 22. Triba, M. N., Zoonens, M., Popot, J.-L., Devaux, P. F., and Warschawski, D. E. (2006) Reconstitution and alignment by a magnetic field of a β-barrel membrane protein in bicelles. Eur. Biophys. J. 35, 268–275. 23. Leitz, A. J., Bayburt, T. H., Barnakov, A. N., Springer, B. A., and Sligar, S. G. (2006) Functional reconstitution of β2-adrenergic receptors utilizing self-assembling Nanodisc technology. BioTechniques 40, 601–612. 24. De Grip, W. J., Van Oostrum, J., and BoveeGeurts, P. H. M. (1998) Selective detergentextraction from mixed detergent/lipid/protein micelles, using cyclodextrin inclusion compounds: a novel generic approach for the preparation of proteoliposomes. Biochem. J. 330, 667–674. 25. Sukharev, S. I., Schroeder, M. J., and McCaslin, D. R. (1999) Stoichiometry of the Large Conductance Bacterial Mechanosensitive Channel of E. coli. A Biochemical Study. J. Membrane Biol. 171, 183–193. 26. Smith, P. K., Krohn, R. I., Hermanson, G. T., Mallia, A. K., Gartner, F. H., Provenzano, M. D., Fujimoto, E. K., Goeke, N. M., Olson, B. J., and Klenk, D. C. (1985) Measurement of protein using bicinchoninic acid. Anal. Biochem. 150, 76–85. 27. Wiechelman, K. J., Braun, R. D., and Fitzpatrick, J. D. (1988) Investigation of the bicinchoninic acid protein assay: identification of the groups responsible for color formation. Anal. Biochem. 175, 231–237. 28. Rouser, G., Fkeischer, S., and Yamamoto, A. (1970) Two dimensional thin layer chromatographic separation of polar lipids and determination of phospholipids by phosphorus analysis of spots. Lipids 5, 494–496. 29. Laird, D. M., Eble, K. S., and Cunningham, C. C. (1986) Reconstitution of mitochondrial F0.F1-ATPase with phosphatidylcholine using the nonionic detergent, octylglucoside. J. Biol. Chem. 261, 14844–14850. 30. Etezady-Esfarjani, T., Hiller, S., Villalba, C., and Wüthrich, K. (2007) Cell-free protein synthesis of perdeuterated proteins for NMR studies. J. Biomol. NMR 39, 229–238.

Chapter 7 Expression and Purification of Src-family Kinases for Solution NMR Studies Andrea Piserchio, David Cowburn, and Ranajeet Ghose Abstract NMR analyses of the structure, dynamics, and interactions of the Src family kinases (SFKs) have been hindered by the limited ability to obtain sufficient amounts of properly folded, soluble protein from bacterial expression systems, to allow these studies to be performed in an economically viable manner. In this chapter, we detail our attempts to overcome these difficulties using the catalytic domain (SrcCD) of c-Src, the prototypical SFK, as an illustrative example. We describe in detail two general methods to express and purify SrcCD from Escherichia coli expression systems in both fully active wild-type and kinase-deficient mutant forms, allowing the efficient and cost-effective labeling by NMR-active isotopes for solution NMR studies. Key words: Protein tyrosine kinases, Src-family kinases, Escherichia coli expression systems

1. Introduction Protein kinases in higher eukaryotes transfer the γ-phosphate of ATP to the hydroxyl groups of specific serine, threonine (serine/ threonine kinases) or tyrosine (tyrosine kinases) residues in substrate proteins. This covalent modification of the Ser/Thr/ Tyr–OH moiety is a fundamental mechanism of cellular regulation and intracellular signal transduction (1–3) and has a significant role in almost every aspect of cell growth, differentiation, maturation, motility, and regulated cell death. In vivo, the phosphorylation levels of kinase targets are tightly controlled by the opposing actions of kinases and their corresponding phosphatases (4, 5). A loss of this control due to mutations, changes in expression levels, alteration of protein–protein interactions including localization, and/or catalytic activity has been implicated in a variety of diseases

Alexander Shekhtman and David S. Burz (eds.), Protein NMR Techniques, Methods in Molecular Biology, vol. 831, DOI 10.1007/978-1-61779-480-3_7, © Springer Science+Business Media, LLC 2012

111

112

A. Piserchio et al.

including a multitude of human cancers (6–9), making protein kinases significant and established targets for anticancer therapies (10–13). Eukaryotic protein kinases contain a highly conserved catalytic domain (CD) (14, 15) that forms a dual-lobed structure (Fig. 1) with a smaller, mainly β-sheet N-terminal lobe (N-lobe), and a larger, mainly α-helical C-terminal lobe (C-lobe). The catalytic activity, while wholly contained within the CD, is often regulated by additional modular domains (16) or insertions within the CD itself (17). While crystallographic analyses of both serine/threonine (18) and tyrosine kinases (19) have provided valuable insights into the regulation of their catalytic activity, several questions remain. The current hypothesis is that most eukaryotic protein kinases function as simple two-state switches transitioning between well-defined “active” and “inactive” conformations based on the phosphorylation state of a small number of regulatory residues. Mounting evidence suggests that this may be an oversimplification and the boundary between fully active and fully inactive states may be not that well defined. Further, the simple phosphorylation-dependent switch scenario is complicated by mounting experimental evidence that hints towards the role played by longrange allosteric dynamic pathways in the regulation of kinase activity, where active-like conformations may be attained through remote interactions (20). Structural details on the nature of recognition of downstream substrates and of regulating kinases and phosphatases are still poorly defined for many of the subfamilies of

Fig. 1. (a) Domain arrangement in inactive full-length c-Src (PDB ID: 2SRC). The residues comprising the linker between the SH2 and the catalytic domain (CD), which engages the SH3 domain, is shown in ball-and-stick representation, as are the residues that comprise the C-terminal tail. The regulatory tyrosine residues in the activation loop (Y416, unphosphorylated in inactive c-Src) and the C-terminal tail (Y527, phosphorylated in inactive c-Src) are also shown in ball-and-stick representation. The ATP-analog, ANP (phosphoaminophosphonic acid-adenylate), bound to the ATP-binding site, is also shown in ball-and-stick representation. (b) Expanded view of SrcCD with the N-lobe and C-lobes indicated. Important regulatory elements that include the activation loop, Gly-rich loop, the HRD (that precedes the catalytic loop) and DFG motifs are also indicated. Y416 is shown in a ball-and-stick representation and the αC helix is indicated by a dashed ellipse. The ligand ANP is also shown bound to the ATP-binding pocket.

7

Expression and Purification of Src-family Kinases for Solution NMR Studies

113

the human kinome. This is not unexpected, given that some of these interactions may be weak or transient, and the resultant complexes may not be amenable to crystallographic analyses. Given this background, it would seem that solution NMR spectroscopy is tailor-made to resolve these unanswered issues about structure, dynamics, and interactions involving protein kinases. Targeting eukaryotic protein kinases by using solution NMR methodologies is especially important because of considerable interest in the pharmaceutical industry in designing small molecule inhibitors of enzymatic activity (21). In particular, the rational design of inhibitors targeting remote sites exerting allosteric control over catalytic activity in protein kinases (type III inhibitors) holds promise as a strategy for inhibiting kinase signaling in a highly selective fashion. These allosteric sites, which are more likely to be unique to specific protein kinases or kinase subfamilies, could be identified by NMR studies performed on a representative subgroup of the human kinome. These studies are expected to assume greater importance as the role of yet uncharacterized protein kinases in cancer becomes clearer by using a combination of RNAi technologies in tumor cell lines and genetic knockdown studies (22–25). Unfortunately, there have been only a handful of NMR studies reported on protein kinases. These include the receptor tyrosine kinase Eph (26), c-Abl (27), c-Src (28), ERK2 (29) protein kinase A (PKA) (30, 31), and Csk (32). A large number of these targets, including PKA and Eph, for which detailed NMR analyses are available, may be expressed and purified with relative ease and are, to a large extent, well behaved in solution, in contrast to a large majority of the human kinome. NMR studies on a broader class of protein kinases have been hindered by difficulties in bacterial expression and purification of sufficient quantities of soluble, properly folded protein for economically viable labeling with NMRactive isotopes. Indeed, the only comprehensive NMR study on a nonreceptor tyrosine kinase examined c-Abl expressed using the baculovirus Sf9 insect cell expression system (27). This system allows limited labeling options and is generally not an economically viable option in a noncommercial setting. Thus, to make these studies viable for a broader set of representative members of the kinome, the optimization of protocols for protein production in bacterial expression systems is required. The problems encountered in NMR studies of eukaryotic protein kinases are exemplified by the Src-family of nonreceptor tyrosine kinases (SFKs) (Fig. 1). c-Src is the prototypical representative of this nine member family (c-Src, Blk, Fgr, Fyn, Hck, Lck, Lyn, Yes, and Yrk) (16, 33, 34). SFKs have been found to be involved in a multitude of human cancers including those of the breast, colon, gastrointestinal tract, lung and in leukemias, lymphomas, and myelomas (35). In particular, the catalytic activity of

114

A. Piserchio et al.

c-Src is highly elevated in a majority of human colon cancers (36). SFKs in general and c-Src in particular are therefore key targets for the design of drugs to treat a wide range of human cancers (37). The catalytic activity of c-Src, and indeed of other SFKs, depends on the phosphorylation state of two conserved tyrosine residues (chicken c-Src numbering used throughout): Tyr416 (activation) and Tyr527 (suppression). Phosphorylation occurs at Tyr416 through an intermolecular mechanism, while Tyr527 is phosphorylated by another family of nonreceptor tyrosine kinases that is comprised of two members: C-terminal Src kinase (Csk) and Cskhomologous kinase (Chk) (38, 39). Several researchers have utilized a variety of systems, including insect cells (40), yeast (Schizosaccharomyces pombe) (41), and platelets (42) to express c-Src. Weijland et al. (41) used a S. pombe based strategy to express the catalytic domain of c-Src by coexpressing the catalytic domain of the tyrosine phosphatase PTP-PEST to reduce catalytic activity and the resultant cytotoxicity. As an extension of this strategy, Seeliger et al. (43) have reported the Escherichia coli expression of the catalytic domains of c-Src and c-Abl by coexpressing the downregulating YopH phosphatase (43). However, multiple issues, including poor sample homogeneity, multiple phosphorylation states (44), sample aggregation or degradation are often encountered when using these strategies. Escherichia coli, the system of choice for production of material amenable to analysis by solution NMR spectroscopy, is particularly affected by these drawbacks (28). The expression and purification of the catalytic domain of c-Src (SrcCD) from E. coli for NMR studies was hindered by several major complications that include the following: 1. Cytotoxicity resulting in poor cell growth after induction. 2. Protein misfolding leading to the accumulation of large amounts of insoluble material in inclusion bodies resulting in vanishing low yields of properly folded soluble protein. Attempts to refold these misfolded aggregates using chaperones in vitro have also not yielded promising results (Piserchio et al. unpubl. results). 3. The production of heterogeneously phosphorylated protein. Resolution of these issues is absolutely necessary for NMR, with its ability to provide both structural and dynamic information over a wide range of timescales, to play a role in studies of these key signaling molecules for drug discovery (45). Here, we describe our efforts toward alleviating these issues by optimal choice of constructs, the use of solubility-enhancing tags, coexpression with chaperones, and carefully optimizing overexpression conditions (28). These methods should provide a useful guide for the bacterial expression and purification of similar protein kinases for NMR studies.

7

Expression and Purification of Src-family Kinases for Solution NMR Studies

2. Expression of the Catalytic Domain of c-Src in Escherichia coli

115

The first strategy that we utilized for the bacterial expression of c-Src involved (1) the addition of an N-terminal MBP fusion tag to enhance both solubility and folding, (2) the coexpression of GroEL and GroES chaperones to further facilitate the accumulation of properly folded protein, an idea originally proposed by Cole (46), and (3) a decrease in the growth temperature after induction to 15°C, thus reducing the rapid accumulation of large quantities of proteins and macromolecular crowding effects that tend to facilitate aggregation. MBP is widely used to enhance the production of fused proteins through a mechanism that is far from clear. Several possible mechanisms have been proposed including the formation of large vesicular aggregates with the misfolded or unfolded proteins contained inside (47) or a general chaperoning effect (48). Our initial attempts to produce the catalytic domain of the so-called “kinasedead mutant” bearing a Lys → Met mutation (SrcCDK295M) (49) in M9 minimal medium showed that in fact, just the introduction of a N-terminal MBP fusion, while not eliminating the presence of protein in the insoluble inclusion bodies, drastically increased the amount of soluble material. However, a majority of the protein produced in this fashion found in solution consisted of soluble aggregates, presumably formed by misfolded material. Observation of similar phenomena has added credence to the speculation that MBP fusions can form micellar-like systems, formed by unfolded material in the core, and by the soluble MBP domain on the surface (47). Though a relatively small amount of properly folded SrcCDK295M was obtained by these preparations, it allowed us to collect the first 15N-HSQC spectra of SrcCDK295M. However, the quantity of soluble, properly folded SrcCD obtained using this procedure was not sufficient for detailed NMR analyses. This was because a large number of experiments, collected over long periods of time, are required for the resonance assignment process, necessitating multiple NMR samples produced using expensive labeling supplies. Nevertheless, the observation that material suitable for NMR experiments could be obtained from an E. coli expression system was indeed encouraging. The largest improvement in the yield of properly folded SrcCDK295M came when the N-terminal MBP fusion system was transformed into E. coli cells that overexpress GroEL and GroES chaperones. Cole and coworkers have shown that overexpression of these bacterial chaperones allow the purification of small amounts of SrcCDK295M from E. coli (49). We found that a more substantial improvement, also achievable in the minimal medium used for NMR sample preparation, can be obtained by integrating the MBP fusion and the GroELS systems. Furthermore, the same approach

116

A. Piserchio et al.

Fig. 2. Production and purification of SrcCDK295M. (a) SDS-PAGE gel (20%) showing different steps in the purification process: lane 1 – GroEL and GroES (expressed alone), lane 2 – cell lysate supernatant, lane 3 – cell lysate pellet, lane 4 – metal affinity column flow through, lane 5 – metal affinity column eluate, lane 6 – product after thrombin cleavage (MBP – higher molecular weight and SrcCDK295M – lower molecular weight), lane 7 – SrcCDK295M after Q-column purification. (b) Effects of controlled phosphorylation and dephosphorylation of SrcCDK295M on the Q column elution profiles. Dashed line: sample preincubated with catalytic amounts of commercial wild-type c-Src and Mg+2/ATP. Dark solid line: sample preincubated with alkaline phosphatase (nonspecific phosphatase). Light solid line: sample preincubated with Lyp phosphatase (specific for phosphorylated Y416). The identity of pSrcCDK295M was confirmed by immunoblots using anti-pTyr antibodies and MS/MS, following trypsin digestion.

could also be used to grow wild-type SrcCD (as opposed to SrcCDK295M), albeit with lower yields, with minimal alteration of the overall protocol. The introduction of the chaperones, however, did not completely eliminate the formation of soluble aggregates mentioned above. However, we noticed that, while these undesired aggregates bound to the amylose columns that are routinely employed to purify MBP fusion proteins, they were unable to bind Co+2 affinity columns despite the presence of an N-terminal His6tag, most likely because the His6-tag was occluded in the misfolded species. This fortuitous scenario allowed us to use metal affinity chromatography to efficiently separate the soluble aggregates from properly folded SrcCD. When a target protein is expressed as an MBP fusion, a critical step prior to NMR analyses is the removal of the bulky MBP tag from the fusion. SrcCD can be efficiently separated from the MBP tag by inserting a thrombin cleavage site between the two components of the fusion construct, although some care has to be taken to avoid protein degradation in the presence of thrombin. We found that resins that bind the N-terminal end of the cleaved fusion (metal-chelating for His6 or amylose for the MBP tag) cannot be efficiently used to isolate SrcCD, given the tendency of the kinase to bind nonspecifically to these columns. We then found that separation of MBP and SrcCD can be carried out using ion exchange chromatography (Fig. 2). The catalytically compromised SrcCDK295M could be efficiently separated from cleaved MBP in this fashion. Interestingly, during purification we found that two forms of SrcCDK295M (one selectively phosphorylated at the activating

7

Expression and Purification of Src-family Kinases for Solution NMR Studies

117

Tyr416 and the other not; confirmed by Western blot against anti-pTyr antibodies and MS/MS analysis) (28) were obtained as two distinct peaks in the elution chromatogram (Fig. 2). This last observation was surprising to some extent, because Q columns do not usually have sufficient resolution to separate identical proteins with unit changes in charge states. Crystallographic and computational analyses (e.g., refs. 50, 51) have revealed that activating phosphorylation alters the overall conformation of the catalytic domain, thus the protein surface that interacts with the column is also likely altered. This feature, we believe, allows the separation of the phosphorylated and nonphosphorylated species using ion-exchange chromatography. We provide below a step-by-step protocol for the purification of SrcCD as an MBP fusion in Subheading 4.1. The MBP/GroELS based protocol, though complex in the nature of its purification scheme, results in high yields of SrcCD purified to homogeneity (up to 700 nmol/L of culture). However, the biggest disadvantage lies in the fact that the catalytic domain of c-Src can only be purified either as a kinase-deficient mutant (SrcCDK295M) or in wild-type form in complex with a high-affinity inhibitor (see Subheading 4.1.2), using this strategy. This prevents access to the full range of catalytically relevant conformations and motional modes, in spite of having the ability to purify both constructs (mutant and wild-type) with the activating Tyr416 in the unphosphorylated and selectively phosphorylated forms. Thus, we developed an alternative strategy to purify wild-type SrcCD that displays full catalytic activity in solution. Our experiences seem to indicate that proper folding appears to be the main hindrance to SrcCD production in bacterial expression systems. In eukaryotes, the folding of several protein kinases including c-Src, occurs with the assistance of the Hsp90 chaperone machinery (52–54). This process is further assisted by an essential kinasespecific cochaperone Cdc37/p50, that can independently bind both the kinase target and the Hsp90 chaperone, thus approximating the kinase and the chaperone machinery, and stabilizing the transient ternary complex (55, 56). Indeed, ternary complexes of Hsp90, v-Src (a transforming viral form of c-Src that contains a tail truncation and lacks Tyr527) and Cdc37 (57) have been identified. We found that coexpressing the Cdc37 cochaperone (the plasmid encoding Cdc37 was a kind gift from Dr. Avrom Caplan, CCNY) with wild-type SrcCD leads to a properly folded soluble kinase domain even in the absence of the N-terminal MBP fusion protein (Fig. 3). Consequently, the His6-tagged kinase can be purified easily using metal affinity chromatography followed by size exclusion chromatography. Thrombin cleavage and ion-exchange chromatography, steps that can both lead to substantial losses in material, become unnecessary. Although the yields are somewhat lower, the methodology is less time consuming, and the wild-type kinase domain can be isolated without the use of an inhibitor, allowing

118

A. Piserchio et al.

Fig. 3. (a) SDS-PAGE gel (20%) showing the various steps in the purification of wild-type SrcCD (His6-tagged): lane 1 – cell lysate supernatant, lane 2 – cell lysate pellet, lane 3 – metal affinity column flow-through (unbound material), lane 4 – metal affinity column eluate, lane 5 – cSrc after purification using gel filtration. The positions of Cdc37, MBP-Lyp and SrcCD in lanes 3–5, respectively, are highlighted by boxes. Samples in lanes 4 and 5 were concentrated by a factor of 10. (b) Size exclusion column profile of the eluate from the metal affinity column purification. (c) Immunoblots of phosphorylated wild-type SrcCD (preincubated with 2 mM ATP and 10 mM Mg+2) with anti-pTyr antibody. All immunoblots were visualized using the Bio-Rad ChemiDoc system. Lane 1 – phosphorylated SrcCD, lane 2 – phosphorylated SrcCD after incubation with alkaline phosphatase.

access to the full complement of structural and dynamics changes that accompany activation. The detailed protocols used are described at length in Subheading 4.2.

3. Materials Most of the materials utilized in the experiments described below are routinely used in NMR laboratories for protein expression and purification. 3.1. Expression Vectors and Competent Cells

1. H-MBP-3C vector (58): a kind gift from Dr. Kaushik Dutta, New York Structural Biology Center (see Note 1). 2. pACYCDuet vector: for simultaneous expression of two proteins (see Note 2): SrcCD and Cdc37 should be cloned into this vector following the protocol described below. 3. 0.1-cm gap sterile electroporation cuvette for bacteria. 4. Electroporator (see Note 3). 5. pREP4-groELS (see Note 4).

electrocompetent

BL21

(DE3)

cells

6. BL21 (DE3) T1 competent cells. 3.2. Media for Bacterial Growth

1. LB (Luria Bertani) Miller medium: Dissolve 20 g of LB powder in 1 L of water, and sterilized by autoclaving. Store at room temperature.

7

Expression and Purification of Src-family Kinases for Solution NMR Studies

119

2. M9 salts: 47.9 mM anhydrous Na2HPO4 (6.8 g/L), 22.0 mM anhydrous KH2PO4 (3 g/L), and 8.6 mM NaCl (0.5 g/L), sterilized by autoclaving. Store at room temperature. 3. Ampicillin stock (1,000×): 100 mg/mL Ampicillin in water. Store at 4°C. 4. Kanamycin stock (1,000×): 50 mg/mL Kanamycin in water. Store at 4°C. 5. Chloramphenicol stock (1,000×): 35 mg/mL Chloramphenicol in Isopropanol. Store at 4°C. 6. LB plates: Mix 40 g/L of LB Miller agar and autoclave the suspension, let the material cool to ~55°C and add the desired antibiotic (100 mg/L Ampicillin, or 50 mg/L Kanamycin, or 35 mg/L Chloramphenicol) or combination of antibiotics (100 mg/L Ampicillin and 50 mg/L Kanamycin, or 100 mg/L Ampicillin and 35 mg/L Chloramphenicol). 7. 1M Isopropyl β-D-1-thiogalactopyranoside (IPTG): Prepare in water and store at 4°C. 8. Metal Solution I (1,000×): 1 mM MnCl2·4H2O, 0.2 mM CoCl2·6H2O, 2 mM ZnCl2, 1 mM CuSO4·5H2O, 50 mM H3BO4, and 0.001 mM Na2MoO4. The compounds are added in the order listed above, dissolving each compound completely before adding the following one. 9. Metal Solution II (1,000×): 1 mM FeSO4, filter-sterilize and store at 4°C. 10. MgSO4 stock (500×): 1M MgSO4, autoclave and store at room temperature. 11. CaCl2 stock (1,000×): 0.1M CaCl2, autoclave and store at room temperature. 12. Vitamin solution (100×): Prepare 10 mL aliquots in 15-mL centrifuge tubes, and store at −20°C (see Note 5). 13.

15

NH4Cl: The material is weighed and carefully dissolved in the minimum amount of preparation medium (40 mL for 0.5 g of NH4Cl), sterile-filtered, and added immediately to the medium.

14. [13C6]-Glucose: The material is weighed and carefully dissolved in the minimum amount of preparation medium (40 mL for 2.5 g of glucose), sterile-filtered, and added immediately to the medium. 15. [U-15N,13C,2H] CELTONE® powder (see Note 6). The material is weighed and carefully dissolved in the minimum amount of preparation medium, sterile-filtered, and added immediately to the medium. Store at room temperature. 16. Anhydrous dextrose.

120

A. Piserchio et al.

17.

14

N ammonium chloride.

18. 0.22-μm Filters. 3.3. Protein Purification

1. Human α-thrombin, store at −80°C. 2. Empty 20-mL bed volume chromatography columns. 3. Preequilibrated cobalt resin: Resuspend the cobalt-based beads, (see Note 7) used for metal-affinity purification of His6-tagged proteins, by gently inverting the bottle. Typically 4 mL of bed volume is used for every 0.5 L of culture (2 mL of slurry provides 1 mL of bed volume). Once resuspended, pipette the slurry into a 20 mL empty column until all the methanol is washed away and the resin settles. Gently wash the column with three bed volumes of deionized H2O followed by three bed volumes of lysis buffer. Resuspend the resin in 1–2 mL of lysis buffer (by gently inverting the closed column) and add to the cell lysate. 4. Lysis buffer: 20 mM Tris–HCl, pH 7.8, 0.5M NaCl, 1% Triton X-100, 0.1% β-mercaptoethanol. The buffer is prepared without β-mercaptoethanol and Triton-X, and stored at room temperature with these two chemicals added immediately before usage. 5. Q columns (5 mL): for ion-exchange chromatography (see Note 8). 6. Ion-exchange chromatography buffer A: 20 mM Tris–HCl, pH 7.4, 10% glycerol. Store at room temperature. 7. Ion-exchange chromatography buffer B: 20 mM Tris–HCl, pH 7.4, 1M NaCl, 10% glycerol. Store at room temperature. 8. Dialysis buffer (before size-exclusion chromatography): 20 mM Tris, pH 7.5 (titrated with 12M HCl), 150 mM NaCl, 1 mM EDTA, and 1 mM DTT (Dithiothreitol), prepared immediately before use. 9. Size exclusion chromatography buffer: 50 mM sodium phosphate, pH 6.5, 150 mM NaCl, 1 mM DTT. 4.82 g of NaH2PO4, and 2.14 g of Na2HPO4 are added to 1 L of water, the pH is adjusted if necessary using 12M HCl or 1M NaOH, The DTT is added immediately before use and the rest stored at room temperature. 10. Gel filtration column (see Note 9): Preequilibrate the column with 1.5 volumes of size exclusion chromatography buffer. The typical flow rate is 0.5 mL/min. 11. AP23464: 10 mM in DMSO. The high-affinity kinase inhibitor (59) is a kind gift from Dr. David Dalgarno, Ariad Pharmaceuticals. Dissolve 4.7 mg in 1 mL of dimethyl sulofoxide. 12. Imidazole buffer: 20 mM Tris, 0.5M NaCl, 1M Imidazole. Dissolve 1.2 g Tris, 14.61 g NaCl, and 34.04 g of Imidazole in ~ 450 mL H2O. Adjust the pH to 7.8 using 12M HCl, bring the solution to 0.5 L volume using a graduated cylinder, and

7

Expression and Purification of Src-family Kinases for Solution NMR Studies

121

readjust the pH to 7.8 using 12M HCl or 1M NaOH. Cover the bottle with aluminum foil and store at room temperature. 13. Cleavage buffer: 20 mM Tris–HCl, pH 7.8, 150 mM NaCl. Add β-mercaptoethanol to a final concentration of 4 mM just before the cleavage reaction. 14. 1M sodium hydroxide: dissolve 40 g of NaOH in ~ 950 mL of water and bring to 1 L volume using a graduated cylinder. 15. EDTA solution: 20 mM Tris, 0.5M EDTA. Dissolve 1.2 g of Tris and 73 g of ethylenediaminetetraacetic acid in 450 mL of water, adjust the pH to 8.0 using 1M NaOH, bring to volume with a graduated cylinder, and adjust the pH to 8.0 using 12M HCl or 1M NaOH. 16. AEBSF: 41.7 mM solution, 1 mL aliquots. Dissolve 10 mg of 4-(2-aminoethyl) benzenesulfonyl fluoride hydrochloride in 1 mL of water and store at −20°C. 17. French press. 18. 12M HCl.

4. Methods 4.1. Expression and Purification of the Catalytic Domain of c-Src as an N-terminal MBP Fusion 4.1.1. Overexpression of SrcCD

The detailed protocol provided below is for the bacterial expression and purification of the catalytic domain of the kinase deficient construct (SrcCDK295M) or the wild-type domain in the presence of a high-affinity inhibitor.

1. Thaw a tube containing the pREP4-groELS electrocompetent BL21 (DE3) cells, on ice (see Note 4). 2. Add ~20 ng of H-MBP-3C vector, encoding SrcCD, to the electrocompetent cells. Transfer the contents into a prechilled 0.1 cm gap electroporation cuvette and incubate for 2 min on ice. 3. Transform the cells with a 6 ms pulse at 1,730 V, and add 960 μL of LB medium, prewarmed to 37°C. Transfer into a sterile 15-mL conical centrifuge tube and shake at 250 rpm for 1 h in an incubator set to 37°C. 4. Plate ~100 μL of transformed cells on a prewarmed (37°C) LB plate containing both Kanamycin and Ampicillin (50 and 100 mg/L, respectively). Incubate overnight at 37°C. 5. Transfer the plate to 4°C for storage. 6. Autoclave 0.5 L of M9 salts in a 2-L flask, transfer 20 mL of this solution into a 50-mL conical tube. From the stock solutions, add 20 μL of 0.1M CaCl2 (final concentration: 0.1 mM),

122

A. Piserchio et al.

40 μL of 1M MgSO4 (final concentration: 2.0 mM), 20 μL of Metal Solutions I and II, 200 μL of Vitamins solution, 20 μL each of Ampicillin and Kanamycin (final concentration: 100 and 50 mg/L, respectively). Finally, add 0.1 g (5 g/L) of anhydrous dextrose and 20 mg (1 g/L) of ammonium chloride. Dissolve by agitation or vortexing, and filter the M9 medium into a preautoclaved 125-mL flask using a sterile 50-mL syringe and a sterile 0.22-μm filter. 7. Scoop a few colonies from the transformed plate using a sterile loop, transfer into the M9 medium and incubate overnight in a shaker (250 rpm) at 37°C. 8. Using a sterile culture pipette, transfer ~20 mL of the M9 salt solution into a sterile 50-mL centrifuge tube. Use this solution to dissolve 2.5 g of dextrose and 0.5 g of ammonium chloride (final concentration: 5 and 1 g/L, respectively). Then, add 5 mL of vitamin solution, dissolve by shaking or vortexing, and filter the entire contents into the original 2-L flask. Add 500 μL of 0.1M CaCl2 1 mL of 1M MgCl2 stock solutions (final concentration: 0.1 and 2.0 mM, respectively), 500 μL each of Metal Solutions I and II, 500 μL of Ampicillin and Kanamycin (final concentration: 100 and 50 mg/L, respectively) to the flask. 9. Add the 20 mL overnight culture directly into the 0.5 L of M9 medium. Incubate at 250 rpm at 37°C. 10. After 1 h, check the OD600 every 30 min; when the OD600 reaches ~0.8, add 100 μL of 1M IPTG (final concentration: 200 μM, see Note 10). Keep the flask at 37°C for about 30 min (see Note 11). 11. Transfer the flask to a second shaker with the temperature set to 15°C, and incubate at 250 rpm overnight (see Note 12). 12. Centrifuge the cells at 4,900 g for 20 min at 4°C. Store the cell pellet at −80°C. 13. For 15N,13C enrichment, replace 14NH4Cl and dextrose in steps 6 and 8 described above, with 15NH4Cl and [U-13C]-glucose, respectively. 14. Under normal circumstances, for uniform 15N,13C,2H enrichment, grow the cells in M9 medium in which H2O is replaced by D2O. However, in the case of SrcCD, we found that the introduction of 2H2O dramatically reduces the yields. Therefore, we follow the approach developed by Fiaux et al. (60), where a pool of 15N,13C,2H-labeled amino acids were used to supplement M9 medium prepared in H2O. In this case: In step 6 above, replace 14NH4Cl with 15NH4Cl; in step 8, above, replace 14 NH4Cl with 15NH4Cl, and use 0.5 g of dextrose (1 g/L instead of 5 g/L) and add 0.75–1.0 g of 15N,13C,2H-labeled algal lysate medium (e.g., CELTONE® 1.5–2.0 g/L).

7

Expression and Purification of Src-family Kinases for Solution NMR Studies

4.1.2. Protocol for the Purification of SrcCD from the MBP Fusion Protein

123

1. Lyse cells using a French Press (1,100 psi), followed by centrifugation at 16,000 g for 20 min at 4°C. Transfer the supernatant to a centrifuge tube. 2. Check the pH of the supernatant (see Note 13), if it is below 7.5, adjust by adding small amounts of solid tris-hydroxymethyl-aminomethane (Tris). 3. Transfer preequilibrated cobalt-resin into the centrifuge tube containing the lysate supernatant, and incubate for about 1 h under rotation or slight agitation at 4°C. 4. Pour the lysate and beads into an empty column. Elute the lysate and wash the column with four bed volumes of lysis buffer and two bed volumes of lysis buffer containing 3 mM imidazole (3 μL Imidazole buffer/mL of lysis buffer). 5. Elute the protein with four bed volumes of the lysis buffer containing 300 mM imidazole (combine three volumes of Imidazole buffer with seven volumes of lysis buffer). 6. Add 20 μL of EDTA solution/mL of eluate (final concentration: 10 mM; see Note 14) to the eluate prior to dialysis and then dialyze against cleavage buffer at 4°C. 7. Determine the concentration of the fusion protein by measuring the OD280 using 119,100 M-1-cm-1 as the extinction coefficient. 8. Add 5 units of α-thrombin/mg of fusion protein to the solution, immediately add β-mercaptoethanol to a final concentration of 4 mM. Cleave overnight at 4°C. 9. Check for completion of the cleavage reaction by running an SDS–PAGE gel. Add more thrombin if necessary. When the cleavage is complete, add solid DTT and the protease inhibitor, AEBSF (4-(2-aminoethyl) benzenesulfonyl fluoride) to final concentrations of 5 mM and 100 μM, respectively. Add AP23464, a high-affinity kinase inhibitor, to the cleavage buffer to a final concentration of 100 μM for wild-type SrcCD. 10. Dilute the completed reaction fivefold into ion exchange chromatography buffer A, and load the resulting sample onto a Q column (ion-exchange) at a flow-rate of 4 mL/min (the volume at this stage is likely to be large, possibly more than 100 mL; see Note 15). 11. Elute the cleaved kinase by running a salt gradient from 0 to 300 mM NaCl, using ion exchange chromatography buffer B, in 120 min using a flow-rate of 1 mL/min. Unphosphorylated SrcCDK295M appears at around 15.4 mS/cm and SrcCDK295M phosphorylated at Tyr416 appears at around 16.9 mS/cm. Wildtype SrcCD bound with the kinase inhibitor, AP23464, appears at around 20.4 mS/cm. MBP elutes at around 8–10 mS/cm, multiple peaks may appear due to nonspecific phosphorylation (see Note 16).

124

A. Piserchio et al.

4.2. Expression and Purification of the Fully Active, Wild-Type Catalytic Domain of c-Src

We provide below a step-by-step protocol for the expression and purification of wild-type SrcCD without the use of an N-terminal MBP tag. The SrcCD obtained using this protocol is fully active and does not require a kinase inhibitor to allow purification to homogeneity. To implement this protocol, first SrcCD and Cdc37 have to be cloned into a pACYCDuet vector (Chloramphenicol resistant) using standard procedures. SrcCD should be inserted in the first cloning site using the EcoRI and SalI restriction enzymes (resulting in a N-terminal His6-tag for SrcCD, see Note 17), and Cdc37 in the second site using XhoI and NdeI (see Note 18). SrcCD expression was also tested with and without the concomitant coexpression of a third protein, the tyrosine phosphatase, Lyp (61) (a kind gift from Dr. Ronald Seidel, Albert Einstein College of Medicine). Lyp selectively dephosphorylates the activating Tyr residue in SFKs (specifically Tyr394 in Lck), and was obtained in a H-MBP-3C Ampicillin resistant vector (see Note 1). 1. If using electrocompetent cells, transfer ~20 ng of pACYCDuet vector into the Eppendorf tube containing BL21(DE3) T1 electrocompetent cells. Transfer the contents of the Eppendorf tube into a prechilled 0.1 cm gap electroporation cuvette and incubate for 2 min on ice (see Note 19). If also overexpressing the phosphatase, then transfer both vectors (pACYC and H-MBP-3C, 20 ng each) into the cuvette at the same time (see Note 20). 2. Follow steps 4–14 of Subheading 4.1.1 but use Chloramphenicol (35 mg/L) and Ampicillin only if overexpressing the phosphatase. This applies to step 4 (Chloramphenicol, or Ampicillin and Chloramphenicol double resistant plates) and step 6 (Chloramphenicol, or Ampicillin and Chloramphenicol added to the M9 medium). 3. Purify the protein following steps 1–6 of Subheading 4.1.2. Note that the final purified protein retains the His6-tag. If the tag needs to be removed, a thrombin cleavage site could be inserted at step 1 in the SrcCD forward primer (described in Note 10). Then steps 7–9 from Subheading 4.1.2. would also apply. Note that, in our construct, the MBP-phosphatase fusion protein has an N-terminal His6 tag and coelutes with the kinase. 4. After elution, dialyze the protein(s) against dialysis buffer. 5. Concentrate the protein(s) down to 2 mL and inject it onto a size exclusion column equilibrated with size exclusion chromatography buffer. Note that the phosphatase elutes in the void (presumably due to some level of self-association) while the kinase elutes consistently at the volume expected for its molecular weight, and are both readily isolated.

7

Expression and Purification of Src-family Kinases for Solution NMR Studies

125

Both methods of expression (with and without coexpression with Lyp) lead to the isolation of active, wild-type SrcCD (confirmed by analysis of kinetics, see Note 21). However, a higher final OD600 and overall yield of purified protein was observed when the kinase was coexpressed with Lyp phosphatase (~190 nmol/L of culture).

5. Conclusions We have described how careful optimization of overexpression protocols allows the expression and purification of the catalytic domain of c-Src both in mutant, as well as active wild-type forms using E. coli expression systems. In particular, we developed two related strategies, the first employs both an N-terminal MBP fusion and coexpression with the GroEL and GroES chaperones, while the second, by taking advantage of the higher specificity of the Cdc37 cochaperone for the nascent c-Src chain, does not require the production of SrcCD as an MBP fusion protein. Both methods present advantages and disadvantages. In particular, the first method leads to higher yields, but it is more efficient in producing the catalytically compromised (K295M) mutant compared to the wild-type protein. The wild-type protein can only be purified in the presence of a high-affinity kinase inhibitor when using this method. The second method produces lower yields; however it also less labor intensive and more importantly, allows the production of the catalytic domain in active, wild-type form. Given the higher yields, the first method is probably more useful when SrcCD is required in large amounts for studies such as those to determine specific protein–protein interactions that do not require the intact catalytic properties of the kinase. However, given that the MBPdependent protocol also produces a considerable amount of soluble misfolded aggregates that have not been observed using the Cdc37-dependent protocol, we believe that the latter may be better suited for “in-cell” NMR studies in E. coli (62–64). Obviously, the latter protocol is required when wild-type, fully active SrcCD needs to be produced. While this manuscript was in production, Campos-Olivas et. al. (65) utilized some of the procedures described herein, to produce sufficient amounts of SrcCD, and obtain backbone resonance assignments in complex with the kinase inhibitor imatinib.

126

A. Piserchio et al.

6. Notes 1. The H-MBP-3C vector contains a thrombin cleavage site and features a maltose binding protein (MBP) tag to enhance solubility and a His6 sequence to facilitate protein purification using a metal affinity chromatography. The protease 3C site described in the original publication (58) has been replaced by a thrombin site in the present case. 2. The pACYCDuet vector was purchased from Novagen (EMD Biosciences). 3. We used an Eppendorf 2510 electroporator. 4. We observed that the transformation of the pREP4-groESL vector was often inefficient, and therefore, we avoided the simultaneous double transformation of pREP4-groESL and H-MBP-3C vector encoding SrcCD, by preparing a stock of electrocompetent cells pretransformed with the former. Electrocompetent cells were preferred over chemically competent cells, due to their ease of preparation and high efficiency of transformation. These cells were prepared by streaking out commercial BL21 (DE3) competent cells on an LB plate and incubating the plate overnight at 37°C. From this, electrocompetent cells were prepared using standard protocols. 5. We used Kao and Michaylchuk vitamin solution (100×) purchased from Sigma. 6. CELTONE® is a rich growth medium for bacteria derived from algal extracts. It may be purchased from Cambridge Isotopes Laboratories. 7. We used TALON beads purchased from Clontech. 8. We used HiTrap Q columns from GE Healthcare Biosciences Corp. 9. We used a HiPrep 16/60 Sephacryl S-100 column from GE Healthcare Biosciences Corp. All protein purification was carried out using an Akta Explorer 100 from GE Healthcare Biosciences Corp. 10. Lower IPTG concentrations are recommended at low temperature to avoid toxicity. 11. This helps to increase the final yield by boosting mRNA production, while most protein is synthesized during the following lower temperature growth. In the first hour after induction, bacteria mainly activate mRNA transcription, while actual translation is still minimal.

7

Expression and Purification of Src-family Kinases for Solution NMR Studies

127

12. All bacterial growth postinduction (after the initial half an hour at 37°C, see Note 11) was carried out at a low temperature (15°C). It is well known that low temperatures decrease the degradation rates of unstable proteins; in the case of SrcCD, we observed that low temperature growth is essential for the production of soluble, properly folded SrcCD. We speculate that SrcCD folding is a slow process that is in competition with the formation of aggregates both soluble and insoluble. In our protocol both the N-terminal MBP fusion and the GroEL/ GroES chaperones (or the Cdc37 cochaperone) provide assistance in stabilizing the nascent chain and protect it from aggregation. Lower temperatures likely assist this process by reducing the rates of protein synthesis and aggregation. 13. We noticed that after long induction times at low temperature, E. coli cells release significant amounts of acid molecules into the lysis buffer. This is, in some instances, sufficient to overcome the buffering power of the solution leading to a pH below 7.0, compromising efficient binding of the His6 tag to the metal affinity column. 14. EDTA is introduced to chelate Co2+ cations. Metal leaks from the resin can be caused by mechanical stress and by high Imidazole concentrations. If not chelated by EDTA, Co2+ can be difficult to eliminate by dialysis, since it can bind the His6-tag as well as the metal binding pockets in SrcCD if left in the sample. Co2+ forms a reddish precipitate once a substantial amount of reducing agent has been added (interfering with the remaining purification process), while the Co2+ remaining in solution, being paramagnetic, degrades the quality of the NMR spectra. 15. We normally do not load more than 15 mg of fusion protein into the ion exchange column at a given time. 16. Unlike the catalytically compromised mutant, wild-type SrcCD does not elute once loaded on the Q column. Presumably the wild-type protein is less stable under conditions of low ionic strength used for sample loading, and precipitates on the column. We found however, that this construct could also be successfully loaded and eluted using the protocol described above by adding a high-affinity kinase inhibitor (59) after cleaving the fusion protein. 17. For SrcCD (human), we used the following primers: Forward: GGTGGTGGAATTCGTCCAAGCCGCAGA CTCAGGG. Reverse: GGTGGTGGTCGACCTAGAGGTTCTCCCC GGGCTGGTA.

128

A. Piserchio et al.

18. For Cdc37 we used the following primers: Forward: GGTGGTGCATATGGTGGACTACAGCG TGTGGGAC. Reverse: GGTGGTGCTCGAGTCACACACTGACATC CTTCTCATCGCC. 19. The electrocompetent cells were homemade and derived from the commercial BL21 (DE3) T1 expression system (chemical competent). We expect that transformation directly into the commercial electrocompetent cells would produce equivalent results. 20. In contrast to what was observed for the pREP4-GroESL vectors, cotransformation of both the H-MBP-3C Lyp vector and the pACYCDuet hosting SrcCD and Cdc37 does not present any difficulties, and can be performed following standard protocols with chemically competent BL21 (DE3) cells. 21. The kcat for the phosphorylation of a substrate peptide polyE4Y by SrcCD was measured using a photometric assay based on the recycling of ADP and the concomitant consumption of NADH catalyzed by L-lactate-dehydrogenase (LDH) and pyruvate kinase (PK). PK regenerates ATP from ADP (a byproduct of the SrcCD phosphorylation process) by converting phosphoenolpyruvate into pyruvate. LDH converts then the pyruvate into lactate by oxidizing NADH. Velocities are obtained following the time-dependent decrease of the NADH fluorescence. This method yielded a kcat of 240 ± 10/min, a value comparable to that obtained previously (41).

Acknowledgments The following grants from the National Institutes of Health, GM084278 (to R. G.), GM047021 (to D. C.) and RR03060 (toward partial support of the NMR facilities at CCNY), are acknowledged. The authors are members of the New York Structural Biology Center, a STAR center supported by the New York State Office for Science, Technology and Academic Research. The authors thank Dr. Avrom Caplan (CCNY), Dr. Ronald Siedel (Albert Einstein College of Medicine), and Dr. Kaushik Dutta (NYSBC) for gifts of materials, and members of the Ghose and Cowburn labs for useful discussions.

7

Expression and Purification of Src-family Kinases for Solution NMR Studies

129

References 1. Hunter, T. (2000) Signaling – 2000 and beyond. Cell 100, 113–127. 2. Johnson, L. N. (2009) The regulation of protein phosphorylation. Biochem. Soc. Trans. 37, 627–641. 3. Tarrant, M. K., and Cole, P. A. (2009) The chemical biology of protein phosphorylation. Annu. Rev. Biochem. 78, 797–825. 4. Ostman, A., and Bohmer, F. D. (2001) Regulation of receptor tyrosine kinase signaling by protein tyrosine phosphatases. Trends Cell Biol. 11, 258–266. 5. Shi, Y. (2009) Serine/threonine phosphatases: mechanism through structure. Cell 139, 468–484. 6. Kolibaba, K. S., and Druker, B. J. (1997) Protein tyrosine kinases and cancer. Biochim. Biophys. Acta 1333, F217–248. 7. Blume-Jensen, P., and Hunter, T. (2001) Oncogenic kinase signalling. Nature 411, 355–365. 8. Chong, P. K., Lee, H., Kong, J. W., Loh, M. C., Wong, C. H., and Lim, Y. P. (2008) Phosphoproteomics, oncogenic signaling and cancer research. Proteomics 8, 4370–4382. 9. Pawson, T., and Kofler, M. (2009) Kinome signaling through regulated protein–protein interactions in normal and cancer cells. Curr. Opin. Cell Biol. 21, 147–153. 10. Johnson, L. N. (2009) Protein kinase inhibitors: contributions from structure to clinical compounds. Q. Rev. Biophys. 42, 1–40. 11. Noble, M. E., Endicott, J. A., and Johnson, L. N. (2004) Protein kinase inhibitors: insights into drug design from structure. Science 303, 1800–1805. 12. Nichols, G. L. (2003) Tyrosine kinase inhibitors as cancer therapy. Cancer Invest. 21, 758–771. 13. Shawver, L. K., Slamon, D., and Ullrich, A. (2002) Smart drugs: tyrosine kinase inhibitors in cancer therapy. Cancer Cell 1, 117–123. 14. Nolen, B., Taylor, S., and Ghosh, G. (2004) Regulation of protein kinases; controlling activity through activation segment conformation. Mol. Cell 15, 661–675. 15. Kornev, A. P., and Taylor, S. S. (2010) Defining the conserved internal architecture of a protein kinase. Biochim. Biophys. Acta 1804, 440–444. 16. Parsons, S. J., and Parsons, J. T. (2004) Src family kinases, key regulators of signal transduction. Oncogene 23, 7906–7909.

17. Roux, P. P., and Blenis, J. (2004) ERK and p38 MAPK-activated protein kinases: a family of protein kinases with diverse biological functions. Microbiol. Mol. Biol. Rev. 68, 320–344. 18. Chen, Z., Gibson, T. B., Robinson, F., Silvestro, L., Pearson, G., Xu, B., Wright, A., Vanderbilt, C., and Cobb, M. H. (2001) MAP kinases. Chem. Rev. 101, 2449–2476. 19. Hubbard, S. R., and Till, J. H. (2000) Protein tyrosine kinase structure and function. Annu. Rev. Biochem. 69, 373–398. 20. Shi, Z., Resing, K. A., and Ahn, N. G. (2006) Networks for the allosteric control of protein kinases. Curr. Opin. Struct. Biol. 16, 686–692. 21. Fedorov, O., Muller, S., and Knapp, S. (2010) The (un)targeted cancer kinome. Nature Chem. Biol. 6, 166–169. 22. Grueneberg, D. A., Degot, S., Pearlberg, J., Li, W., Davies, J. E., Baldwin, A., Endege, W., Doench, J., Sawyer, J., Hu, Y., Boyce, F., Xian, J., Munger, K., and Harlow, E. (2008) Kinase requirements in human cells: I. Comparing kinase requirements across various cell types. Proc. Natl. Acad. Sci. U.S.A. 105, 16472–16477. 23. Baldwin, A., Li, W., Grace, M., Pearlberg, J., Harlow, E., Munger, K., and Grueneberg, D. A. (2008) Kinase requirements in human cells: II. Genetic interaction screens identify kinase requirements following HPV16 E7 expression in cancer cells. Proc. Natl. Acad. Sci. U.S.A. 105, 16478–16483. 24. Bommi-Reddy, A., Almeciga, I., Sawyer, J., Geisen, C., Li, W., Harlow, E., Kaelin, W. G., Jr., and Grueneberg, D. A. (2008) Kinase requirements in human cells: III. Altered kinase requirements in VHL−/− cancer cells detected in a pilot synthetic lethal screen. Proc. Natl. Acad. Sci. U.S.A. 105, 16484–16489. 25. Grueneberg, D. A., Li, W., Davies, J. E., Sawyer, J., Pearlberg, J., and Harlow, E. (2008) Kinase requirements in human cells: IV. Differential kinase requirements in cervical and renal human tumor cell lines. Proc. Natl. Acad. Sci. U.S.A. 105, 16490–16495. 26. Wiesner, S., Wybenga-Groot, L. E., Warner, N., Lin, H., Pawson, T., Forman-Kay, J. D., and Sicheri, F. (2006) A change in conformational dynamics underlies the activation of Eph receptor tyrosine kinases. EMBO J. 25, 4686–4696. 27. Vajpai, N., Strauss, A., Fendrich, G., CowanJacob, S. W., Manley, P. W., Grzesiek, S., and Jahnke, W. (2008) Solution conformations and

130

28.

29.

30.

31.

32.

33.

34. 35.

36.

37.

38.

39.

A. Piserchio et al. dynamics of Abl kinase-inhibitor complexes determined by NMR substantiate the different binding modes of imatinib/nilotinib and dasatinib. J. Biol. Chem. 283, 18292–18302. Piserchio, A., Ghose, R., and Cowburn, D. (2009) Optimized bacterial expression and purification of the c-Src catalytic domain for solution NMR studies. J. Biomol. NMR 44, 87–93. Piserchio, A., Warthaka, M., Devkota, A. K., Kaoud, T. S., Lee, S., Abramczyk, O., Ren, P., Dalby, K. N., and Ghose R. (2011) Solution NMR insights into docking interactions involving inactive ERK2. Biochemistry 50, 3660–3672. Masterson, L. R., Mascioni, A., Traaseth, N. J., Taylor, S. S., and Veglia, G. (2008) Allosteric cooperativity in protein kinase A. Proc. Natl. Acad. Sci. U.S.A. 105, 506–511. Masterson, L. R., Cheng, C., Yu, T., Tonelli, M., Kornev, A., Taylor, S. S., and Veglia, G. (2010) Dynamics connect substrate recognition to catalysis in protein kinase A. Nature Chem. Biol. 6, 821–828. Liu, D., Xu, R., and Cowburn, D. (2009) Segmental isotopic labeling of proteins for nuclear magnetic resonance. Meth. Enzymol. 462, 151–175. Boggon, T. J., and Eck, M. J. (2004) Structure and regulation of Src family kinases. Oncogene 23, 7918–7927. Martin, G. S. (2001) The hunting of the Src. Nature Rev. Mol. Cell Biol. 2, 467–475. Alvarez, R. H., Kantarjian, H. M., and Cortes, J. E. (2006) The role of Src in solid and hematologic malignancies: development of newgeneration Src inhibitors. Cancer 107, 1918–1929. Russello, S. V., and Shore, S. K. (2003) Src in human carcinogenesis. Front. Biosci. 8, s1068–1073. Trevino, J. G., Summy, J. M., and Gallick, G. E. (2006) Src inhibitors as potential therapeutic agents for human cancers. Mini Rev. Med. Chem. 6, 681–687. Grgurevich, S., Mikhael, A., and McVicar, D. W. (1999) The Csk homologous kinase, Chk, binds tyrosine phosphorylated paxillin in human blastic T cells. Biochem. Biophys. Res. Commun. 256, 668–675. Zrihan-Licht, S., Deng, B., Yarden, Y., McShan, G., Keydar, I., and Avraham, H. (1998) Csk homologous kinase, a novel signaling molecule, directly associates with the activated ErbB-2 receptor in breast cancer cells and inhibits their proliferation. J. Biol. Chem. 273, 4065–4072.

40. Sicheri, F., Moarefi, I., and Kuriyan, J. (1997) Crystal structure of the Src family tyrosine kinase Hck. Nature 385, 602–609. 41. Weijland, A., Neubauer, G., Courtneidge, S. A., Mann, M., Wierenga, R. K., and SupertiFurga, G. (1996) The purification and characterization of the catalytic domain of Src expressed in Schizosaccharomyces pombe. Comparison of unphosphorylated and tyrosine phosphorylated species. Eur. J. Biochem. 240, 756–764. 42. Feder, D., and Bishop, J. M. (1990) Purification and enzymatic characterization of pp60c-src from human platelets. J. Biol. Chem. 265, 8205–8211. 43. Seeliger, M. A., Young, M., Henderson, M. N., Pellicena, P., King, D. S., Falick, A. M., and Kuriyan, J. (2005) High yield bacterial expression of active c-Abl and c-Src tyrosine kinases. Protein Sci. 14, 3135–3139. 44. Stover, D. R., Liebetanz, J., and Lydon, N. B. (1994) Cdc2-mediated modulation of pp60csrc activity. J. Biol. Chem. 269, 26885–26889. 45. Pellecchia, M., Bertini, I., Cowburn, D., Dalvit, C., Giralt, E., Jahnke, W., James, T. L., Homans, S. W., Kessler, H., Luchinat, C., Meyer, B., Oschkinat, H., Peng, J., Schwalbe, H., and Siegal, G. (2008) Perspectives on NMR in drug discovery: a technique comes of age. Nature Rev. Drug Discov. 7, 738–745. 46. Cole, P. A. (1996) Chaperone-assisted protein expression. Structure 4, 239–242. 47. Nomine, Y., Ristriani, T., Laurent, C., Lefevre, J. F., Weiss, E., and Trave, G. (2001) Formation of soluble inclusion bodies by HPV E6 oncoprotein fused to maltose-binding protein. Protein Expr. Purif. 23, 22–32. 48. Kapust, R. B., and Waugh, D. S. (1999) Escherichia coli maltose-binding protein is uncommonly effective at promoting the solubility of polypeptides to which it is fused. Protein Sci. 8, 1668–1674. 49. Wang, D., Huang, X. Y., and Cole, P. A. (2001) Molecular determinants for Csk-catalyzed tyrosine phosphorylation of the Src tail. Biochemistry 40, 2004–2010. 50. Ozkirimli, E., and Post, C. B. (2006) Src kinase activation: a switched electrostatic network. Protein Sci. 15, 1051–1062. 51. Cowan-Jacob, S. W. (2006) Structural biology of protein tyrosine kinases. Cell Mol. Life Sci. 63, 2608–2625. 52. Welch, W. J., and Feramisco, J. R. (1982) Purification of the major mammalian heat shock proteins. J. Biol. Chem. 257, 14949–14959.

7

Expression and Purification of Src-family Kinases for Solution NMR Studies

53. Hahn, J. S. (2009) The Hsp90 chaperone machinery: from structure to drug development. BMB Rep. 42, 623–630. 54. Zuehlke, A., and Johnson, J. L. (2010) Hsp90 and co-chaperones twist the functions of diverse client proteins. Biopolymers 93, 211–217. 55. Caplan, A. J., Mandal, A. K., and Theodoraki, M. A. (2007) Molecular chaperones and protein kinase quality control. Trends Cell Biol. 17, 87–92. 56. Karnitz, L. M., and Felts, S. J. (2007) Cdc37 regulation of the kinome: when to hold ‘em and when to fold ‘em. Science STKE 2007, pe22. 57. Whitelaw, M. L., Hutchison, K., and Perdew, G. H. (1991) A 50-kDa cytosolic protein complexed with the 90-kDa heat shock protein (hsp90) is the same protein complexed with pp60v-Src hsp90 in cells transformed by the Rous sarcoma virus. J. Biol. Chem. 266, 16436–16440. 58. Alexandrov, A., Dutta, K., and Pascal, S. M. (2001) MBP fusion protein with a viral protease cleavage site: one-step cleavage/purification of insoluble proteins. Biotechniques 30, 1194–1198. 59. Dalgarno, D., Stehle, T., Narula, S., Schelling, P., van Schravendijk, M. R., Adams, S., Andrade, L., Keats, J., Ram, M., Jin, L., Grossman, T., MacNeil, I., Metcalf, C., 3rd, Shakespeare, W., Wang, Y., Keenan, T., Sundaramoorthi, R., Bohacek, R., Weigele, M., and Sawyer, T. (2006) Structural basis of Src tyrosine kinase inhibition with a new class of potent and

60.

61.

62.

63.

64.

65.

131

selective trisubstituted purine-based compounds. Chem. Biol. Drug Des. 67, 46–57. Fiaux, J., Bertelsen, E. B., Horwich, A. L., and Wuthrich, K. (2004) Uniform and residue-specific 15N-labeling of proteins on a highly deuterated background. J. Biomol. NMR 29, 289–297. Wu, J., Katrekar, A., Honigberg, L. A., Smith, A. M., Conn, M. T., Tang, J., Jeffery, D., Mortara, K., Sampang, J., Williams, S. R., Buggy, J., and Clark, J. M. (2006) Identification of substrates of human protein-tyrosine phosphatase PTPN22. J. Biol. Chem. 281, 11002–11010. Burz, D. S., Dutta, K., Cowburn, D., and Shekhtman, A. (2006) In-cell NMR for protein–protein interactions (STINT-NMR). Nature Protoc. 1, 146–152. Selenko, P., and Wagner, G. (2006) NMR mapping of protein interactions in living cells. Nature Meth. 3, 80–81. Sakakibara, D., Sasaki, A., Ikeya, T., Hamatsu, J., Hanashima, T., Mishima, M., Yoshimasu, M., Hayashi, N., Mikawa, T., Walchli, M., Smith, B. O., Shirakawa, M., Guntert, P., and Ito, Y. (2009) Protein structure determination in living cells by in-cell NMR spectroscopy. Nature 458, 102–105. Campos-Olivas, R., Marenchino, M., Scapozza, L. and Gervasio, F. L. (2011) Backbone assignment of the tyrosine kinase Src catalytic domain in complex with imatinib. Biomol. NMR Asgn. 5, 221–224.

Chapter 8 NMR Studies of Large Protein Systems Shiou-Ru Tzeng, Ming-Tao Pai, and Charalampos G. Kalodimos Abstract Over the recent years, there has been increased interest in applying NMR spectroscopy for the characterization of proteins and protein complexes of large molecular weight. The combination of multidimensional NMR, novel pulse sequences allowing for the selection of slowly relaxing coherence pathways, and the development of a range of labeling techniques has enabled high-resolution NMR analyses of supramolecular systems of even megadalton size. Here, we describe how NMR can be used to obtain structural information in large systems by using as an example the recent structure determination of SecA ATPase (204 kDa) in complex with a signal peptide. Key words: Paramagnetic resonance enhancement, Macromolecular complex, Molecular machinery, Stable isotope labeling

1. Introduction Application of NMR spectroscopy to supramolecular systems (>200 kDa) has been revolutionized by specific labeling of methyl groups (1). The labeling protocol, pioneered by the group of Lewis Kay, is very simple and robust (2, 3). The approach exploits some very favorable properties of methyl groups in proteins: (a) they occur frequently in the hydrophobic cores of proteins and at the interfaces of biomolecular complexes, and are thus excellent reporters of structure and dynamics; (b) the three protons of the methyl group all contribute to the intensity of the same signal, and therefore methyl probes are significantly more sensitive than other candidates; and (c) methyl groups are intrinsically optimized for use in TROSY

Alexander Shekhtman and David S. Burz (eds.), Protein NMR Techniques, Methods in Molecular Biology, vol. 831, DOI 10.1007/978-1-61779-480-3_8, © Springer Science+Business Media, LLC 2012

133

134

S.-R. Tzeng et al.

spectroscopy and the simple 1H-13C HMQC experiment can be used to select for pathways with favorable relaxation properties (4). Currently, the methyl groups of five different amino acids can be labeled in a highly specific and scramble-free manner: Ala, Ile (δ1), Leu, Met, and Val. These five residues are highly abundant, typically accounting for 35–45% of the total number of residues in a protein and are distributed throughout the protein, thus providing almost complete coverage of the protein space. The methyl-labeling approach combined with methyl-TROSY currently provides the method of choice for NMR characterization of large protein systems. Although this approach has proven to be very robust for recording spectra of large proteins with high sensitivity and resolution, a major hurdle in obtaining site-specific information remains the difficulty in obtaining assignments. While the traditional approach of assigning the backbone and subsequently linking the methyl side chains to the backbone has worked efficiently for smaller proteins, it is not applicable to larger systems. The only approach currently is to “disassemble” the supramolecular system. For higher order oligomeric systems, such as the proteasome (5), this means preparing the subunit in its monomeric form and for large, single-chain proteins, such as the SecA (6), preparing isolated domains or fragments. In principle, determining solution structures of supramolecular protein–ligand complexes by NMR should be feasible, provided that the crystal structures of the free partners are previously known. Because usually only methyl groups can be robustly and unambiguously detected for supramolecular systems, in cases where complex interactions are mediated by hydrophobic contacts involving methyl-bearing residues it is likely that intermolecular NOEs can be detected, thereby enabling the reliable docking of the complex. Unfavorable motions commonly observed at protein interfaces, however, may result in line broadening and render NOE detection unfeasible. Although the NOE has served as the gold standard for protein structure determination by NMR, the old, but recently resurrected, paramagnetic relaxation enhancement (PRE) (7, 8) technique holds great promise for obtaining both structural and dynamic information in supramolecular protein complexes (6). By combining transferred NOESY, line broadening, and PRE experiments, the structure of the 204 kDa SecA ATPase in complex with a secretory signal peptide was recently determined (6). Using this system as an example, we describe strategies to (a) obtain samples optimally labeled for methyl detection, (b) assign the methyl resonances of the large protein system, and (c) obtain intermolecular distance restraints for the structure determination of large protein–ligand complexes.

8

NMR Studies of Large Protein Systems

135

2. Materials 1. Frozen, transformed Escherichia coli BL21(DE3) cells to overexpress protein of interest. 2. M9 medium: 6 g/L Na2HPO4, 3 g/L KH2PO4, pH 7.0–7.4., 0.5 g/L NaCl, 1.0 g/L NH4Cl or 15NH4Cl. Autoclave, let the medium cool down, and then add 0.1 mL/L of 1M CaCl2, 1 mL/L of 1M MgSO4, and 2 g/L D-[2H,12C]-glucose or D-[2H,13C]-glucose. 3. 1M CaCl2 stock: Dissolve 11.0 g of CaCl2 in 100 mL of D2O; filter sterilize. 4. 1M MgSO4 stock: Dissolve 12.04 g of MgSO4 in 100 mL of D2O; filter sterilize. 5. D2O (Cambridge Isotope Laboratories, CIL). 6. BIOEXPRESS (CIL). 7. ISOGRO (Isotec). 8.

13

CH3-2H-alanine.

9. α-Ketobutyrate (CIL or Isotec). 10. α-Ketoisovalerate (CIL or Isotec). 11.

13

CH3-methionine (CIL).

12. IPTG. 13. AMICON stir cell (Millipore). 14. L-broth: 10 g tryptone/L, 5 g yeast extract/L, 5 g NaCl/L, adjusted to pH 7.4 with NaOH; autoclave. 15. (2,2,5,5-Tetramethyl-2,5-dihydro-1H-pyrrol-3-yl)methyl methanesulfonothioate (MTSL) (Toronto Research Chemicals Inc.): Dissolve in acetonitirile. 16. Ascorbic acid.

3. Methods 3.1. Protein Labeling for Methyl Detection

1. Pick a freshly transformed colony (see Note 1) of BL21(DE3) cells and inoculate a 1–2 mL culture of L-broth in D2O containing 0.1% of glucose at 37°C until cells reach an OD600 of ~0.7–0.8. 2. Centrifuge the cells at 1,200 × g for 20 min at room temperature and resuspend them in 5–10 mL of sterile M9 medium in D2O (M9/D2O) in a sterile flask to a starting OD600 of ~0.05. Incubate the culture in a shaking incubator (220–250 rpm) at 37°C until it reaches an OD600 of ~0.6.

136

S.-R. Tzeng et al.

3. Centrifuge as in step 2, and resuspend the cells in 50–100 mL of M9/D2O prepared with either D-[2H,13C]glucose (see Note 2) or D-[2H, 12C]glucose (see Note 3) and 15NH4Cl containing 2% of BIOEXPRESS (CIL) or ISOGRO (Sigma) (see Note 4). The starting OD600 should be ~0.1. Incubate the culture in a shaking incubator (220–250 rpm) at 37°C until the OD600 is ~0.6. 4. Centrifuge as in step 2, resuspend the cells in 1 L of M9/D2O, and grow until the OD600 is ~0.25. 5. At this point, amino acid precursors for methyl labeling can be added. (a) Add precursors (see Note 5) or amino acids 30–60 min prior to IPTG induction; add the following quantities (final concentration): 100 mg/L of 13CH3-2Hα-alanine (see Note 6) for Ala labeling; 45–50 mg/L of α-ketobutyrate for isoleucine labeling; 85–100 mg/L of α-ketoisovalerate for leucine and valine labeling (see Note 7); 250 mg/L of [13CH3]-methionine for Met labeling. The methyl groups of all five residues can be labeled in one sample in a scramblefree manner (see Note 8) (Fig. 1a). (b) Continue incubating the culture for approximately 1 h. The OD600 should reach a value of ~0.3–0.4. 6. Add IPTG to 0.5 mM to induce protein overexpression. 7. Continue postinduction growth for 6–8 h (see Note 9). 8. Harvest the cells by centrifugation at 5,000 × g for 15 min at 4°C. 9. Freeze the wet cell pack at −80°C. a

b 12

20

Met

16 18

Leu, Val 20

(p.p.m.)

21 Ala

13C

13C

(p.p.m.)

14

Ile

22 23 24

22 25 24 26 26 27 2.0

1.6

1.2 0.8 1H (p.p.m.)

0.4

0.0

1.2 1.0

0.8 0.6

0.4

0.2

0.0

Fig. 1. (a) 1H-13C HMQC of [U-2H,12C], Ala-, Leu-, Met-, Val-, Ile-δ1-[13CH3] labeled protein. The methyl groups of all five amino acids can be labeled with no scrambling. (b) 1H-13C HMQC of the same protein as in (a, black) but prepared using 10% BIOEXPRESS (green). The Leu methyl groups are completely suppressed, whereas the Val methyl groups are only minimally affected.

8

3.2. NMR Assignment

NMR Studies of Large Protein Systems

137

To assign a large protein, such as SecA (204 kDa; 901 residues per subunit), a domain-parsing strategy is followed. 1. Isolate and characterize, by using NMR, virtually all domains of the full-length protein and a number of fragments comprising contiguous domains (see Note 10). The size of the isolated domains and fragments should be such that backbone and side-chain assignment is feasible using standard approaches. 2. Prepare Ala, Ile, Met, Leu, and Val methyl-labeled samples for the full-length protein and the domains thereof (see Note 11). Record methyl-TROSY for all of the samples, overlay, and compare the spectra of the individual domains against the spectra of the longer fragments and full-length protein. If good resonance correspondence among domains, fragments, and the full-length protein can be demonstrated, then assignment is in principle transferable. 3. Record standard triple-resonance NMR experiments for isolated domains and obtain backbone assignments. Record the threedimensional spectra required for the assignment of the methyl groups (see Note 12). 4. Transfer the methyl assignments obtained for the isolated domains to the larger fragments and finally to the full-length protein by visually inspecting the methyl-TROSY spectra. Only the assignment of the obvious and well-dispersed resonances can be safely transferred this way. 5. Record 13C HMQC-NOESY-HMQC spectra (see Note 13) for the methyl-labeled samples. Use the NOE patterns to confirm and extend the assignment transfer from the domains to the full-length protein. If a crystal structure is available, it can be used to determine the distances between the methyl groups and assist with the assignment. 6. Prepare site-directed mutations to assign ambiguous resonances and further extend and confirm the assignments (see Note 14).

3.3. Paramagnetic Relaxation Enhancement Measurements

1. To prepare nitroxide spin label (MTSL)-derivatized ligand via cysteine-specific modification of engineered ligand derivatives containing single-solvent-accessible cysteine residues at sites of interest (see Note 15), add MTSL from a concentrated stock in acetonitrile to the ligand solution (free from any reducing agent) at a tenfold molar excess over the ligand and allow the reaction to proceed at 4°C for ~12 h. If available, confirm the completion of the reaction by mass spectrometry. 2. Remove excess MTSL by extensive dialysis using an Amicon stirred cell. 3. Determine PRE-derived distances from 1H-13C HMQC spectra by measuring peak intensities before (paramagnetic) and after

138

S.-R. Tzeng et al.

(diamagnetic) reduction of the nitroxide spin label by the addition of 5 mM ascorbic acid (see Note 16). 4. Convert PRE values to distances by using a modified SolomonBloembergen equation for transverse relaxation (7). 5. Incorporate distance (intermolecular) restraints into the structure calculation protocol of the complex. 6. Restrain resonances strongly affected by the presence of the spin label in the ligand (Ipara /Idia < 0.15) and whose resonances broaden beyond detection in the paramagnetic spectrum with only an upper bound distance estimated from the noise of the spectrum plus 4 Å. 7. Restrain resonances that appear in the paramagnetic spectra (Ipara /Idia < 0.85) as the calculated distance with ±4 Å upper/ lower bounds. 3.4. Structure Determination

1. Determine the interface between the ligand and the large protein using differential line broadening (9). The residues affected by complex formation can be used as ambiguous restraints. 2. If the ligand is a flexible peptide, use transferred NOESY (10) to determine the structure of the peptide in the complex. Determine the structure of the complex by using a CNS-based software, such as HADDOCK (11) or Xplor-NIH (12). Use the crystal structure of the large protein to define the starting conformation, and both unambiguous and ambiguous restraints obtained from NOE, PRE, line broadening, and chemical shift perturbation experiments.

4. Notes 1. Freshly transformed colonies always give better protein yield. 2. [U-2H,13C]-glucose should be used when uniform 13C labeling is desired or when all side-chain carbons of the methyl-bearing residues are to be 13C labeled for magnetization transfer from methyls to the backbone. In this case, the uniformly 13C-labeled ketoacid precursor must be used. 3. [U-2H,12C]-glucose should be used to produce an NMR sample in which all carbons are 12C labeled, except the methyl carbons of interest. 1H-13C HMQC spectra of such samples are recorded without the use of the constant time version and typically provide the best resolution. Such a sample can also be used for relaxation experiments (1). 4. Up to ~2.5% of a rich labeling medium can be used to increase the protein yield with no effect on the specific labeling of the methyl groups.

8

NMR Studies of Large Protein Systems

139

5. Precursors can be purchased in protonated form and dissolved in D2O for exchange to take place: at pH 12.5 (45°C), 2–3 h for α-ketoisovalerate, and at pH 10.5 (45°C), 12–14 h for α-ketobutyrate; the pH values are optimized for exchange and prevent the generation of dimers through condensation of two ketoacid molecules. 6.

CH3-2Hα-alanine can be prepared by using the tryptophan synthase enzyme to catalyze the proton-to-deuterium exchange of the α hydrogen, as described by Matthews and coworkers (13). 13

7. Incorporation of 13CH3/12CD3 isotope labels into the isopropyl moieties of Val and Leu residues should be used for very large proteins since the inter-methyl dipolar relaxation is significantly reduced. The methyl-TROSY spectra show significant gains in resolution with practically no losses in sensitivity despite the twofold dilution of the NMR-active methyls in such samples. Precursors have also become available that allow any of the methyl isotopomers (13CHD2, 13CH2D, and 13CH3) to be incorporated into the protein (1). The different isotopomers can be used for relaxation experiments. 8. In this case, addition of ~2% of a rich labeling medium (e.g., BIOEXPRESS or ISOGRO) is required to suppress scrambling associated with the addition of the alanine amino acid. Interestingly, further increase of the rich labeling medium (~10%) suppresses completely the methyl labeling of Leu while having a minimal effect on the methyl labeling of Val (Fig. 1b). Since the methyl groups of these two residues often overlap, this labeling scheme can be used to differentiate between the two. 9. Critical step: Excessively prolonged growth after induction should be avoided to prevent generation of methyl groups with undesired isotopomers. 10. The design of domains and fragments thereof that would retain their fold and are soluble in isolation can be quite tricky. In this respect, the availability of a crystal structure can be of tremendous help. 11. When the methyl residues of all five residues are labeled in the same sample of a very large protein, the signal may be significantly compromised due to enhanced inter-methyl relaxation. The preparation of multiple samples each containing a single amino acid labeled may be desirable in such a case. 12. An arsenal of pulse sequences are available for methyl assignment (14). 13. The highly deuterated background suppresses spin diffusion, and thus the mixing time for the NOESY experiments can be set as high as 500 ms allowing for NOEs to be observed between methyl groups as far as ~8 Å.

140

S.-R. Tzeng et al.

14. Amino acids should be typically substituted by an isosteric amino acid to prevent significant changes in the local environment and protein packing, which could introduce significant chemical shift effects. 15. Nonreactive Cys residues can be judged by the Elman’s test. Sites for MTSL incorporation should be selected so that they cause no or minimal effect on protein structure. This can be assessed by NMR. 16. PRE rates should typically be measured using several MTSLderivatized ligands, each containing a single MTSL at a different site. Because PRE rates provide long-range distance information, in the absence of available NOE data a large number of PREs are required to properly determine the structure of a protein– ligand complex. The complex between SecA and the signal peptide was determined using 160 PRE-derived intermolecular restraints. References 1. Ruschak, A. M., and Kay, L. E. (2010) Methyl groups as probes of supra-molecular structure, dynamics and function. J. Biomol. NMR 46, 75–87. 2. Goto, N., Gardner, K., Mueller, G., Willis, R., and Kay, L. (1999) A robust and cost-effective method for the production of Val, Leu, Ile (delta 1) methyl-protonated 15N-, 13C-, 2H-labeled proteins. J. Biomol. NMR 13, 369–374. 3. Tugarinov, V., Kanelis, V., and Kay, L. E. (2006) Isotope labeling strategies for the study of highmolecular-weight proteins by solution NMR spectroscopy. Nat. Protoc. 1, 749–754. 4. Tugarinov, V., Hwang, P., Ollerenshaw, J., and Kay, L. (2003) Cross-correlated relaxation enhanced 1H-13C NMR spectroscopy of methyl groups in very high molecular weight proteins and protein complexes. J. Am. Chem. Soc. 125, 10420–10428. 5. Sprangers, R., and Kay, L. E. (2007) Quantitative dynamics and binding studies of the 20S proteasome by NMR. Nature 445, 618–622. 6. Gelis, I., Bonvin, A., Keramisanou, D., Koukaki, M., Gouridis, G., Karamanou, S., Economou, A., and Kalodimos, C. G. (2007) Structural basis for signal-sequence recognition by the translocase motor SecA as determined by NMR. Cell 131, 756–769. 7. Battiste, J., and Wagner, G. (2000) Utilization of site-directed spin labeling and high-resolution heteronuclear nuclear magnetic resonance for global fold determination of large proteins with

limited nuclear overhauser effect data. Biochemistry 39, 5355–5365. 8. Tang, C., Schwieters, C., and Clore, G. (2007) Open-to-closed transition in apo maltose-binding protein observed by paramagnetic NMR. Nature 449, 1078–1082. 9. Takeuchi, K., and Wagner, G. (2006) NMR studies of protein interactions. Curr. Opin. Struct. Biol. 16, 109–117. 10. Post, C. (2003) Exchange-transferred NOE spectroscopy and bound ligand structure determination. Curr. Opin. Struct. Biol. 13, 581–588. 11. de Vries, S. J., van Dijk, M., and Bonvin, A. M. (2010) The HADDOCK web server for data-driven biomolecular docking. Nat. Protoc. 5, 883–897. 12. Schwieters, C. D., Kuszewski, J. J., Tjandra, N., and Clore, G. M. (2003) The Xplor-NIH NMR molecular structure determination package. J. Magn. Reson. 160, 65–73. 13. Isaacson, R., Simpson, P., Liu, M., Cota, E., Zhang, X., Freemont, P., and Matthews, S. (2007) A new labeling method for methyl transverse relaxation-optimized spectroscopy NMR spectra of alanine residues. J. Am. Chem. Soc. 129, 15428–15429. 14. Tugarinov, V., and Kay, L. (2003) Ile, Leu, and Val methyl assignments of the 723-residue malate synthase G using a new labeling strategy and novel NMR methods. J. Am. Chem. Soc. 125, 13868–13878.

Chapter 9 Protein Dynamics by 15N Nuclear Magnetic Relaxation Fabien Ferrage Abstract Nitrogen-15 relaxation is the most ubiquitous source of information about protein (backbone) dynamics used by NMR spectroscopists. It provides the general characteristics of hydrodynamics as well as internal motions on subnanosecond, micro- and millisecond timescales of a biomolecule. Here, we present a full protocol to perform and analyze a series of experiments to measure the 15N longitudinal relaxation rate, the 15N transverse relaxation rate under an echo train or a single echo, the 15N–1H dipolar cross-relaxation rate, as well as the longitudinal and transverse cross-relaxation rates due to the cross-correlation of the nitrogen-15 chemical shift anisotropy and the dipolar coupling with the adjacent proton. These rates can be employed to carry out model-free analyses and can be used to quantify accurately the contribution of chemical exchange to transverse relaxation. Key words: Nuclear magnetic resonance, Protein dynamics, Relaxation rates, Nitrogen-15, Longitudinal relaxation, Transverse relaxation, Cross-relaxation, Cross-correlated relaxation, Chemical exchange

1. Introduction Nuclear magnetic resonance is a fantastic tool to investigate the dynamics of biomolecules and, in particular, proteins. Among the numerous techniques available, which provide access to internal motions over a wide range of timescales, measurements of backbone nitrogen-15 relaxation rates have proven to be by far the most popular method to sample and quantify protein dynamics (1–3). Nitrogen-15 relaxation rates can be analyzed to determine the hydrodynamic properties of proteins (4), internal motions faster than overall motions (subnanosecond) as well as slower motions on microsecond–millisecond timescales that give rise to chemical exchange-induced relaxation. Fast subnanosecond motions are closely related to the local atomic density of a protein, and they

Alexander Shekhtman and David S. Burz (eds.), Protein NMR Techniques, Methods in Molecular Biology, vol. 831, DOI 10.1007/978-1-61779-480-3_9, © Springer Science+Business Media, LLC 2012

141

142

F. Ferrage

contribute to conformational entropy (5, 6) and are a good indication of the local malleability of the protein structure. Hydrodynamic properties can be used for structural refinement of a single-domain (7) or tight multidomain protein or protein complex (8), interdomain motions may also be analyzed (9). Microsecond and millisecond motions and chemical reactions are essential to protein function. In most cases, the kinetics of these events can be determined by nitrogen-15 relaxation methods (10, 11), while, in favorable cases, both thermodynamics and structural changes between exchanging states can be characterized (12). The canonical set of nitrogen-15 relaxation experiments comprises the measurements of longitudinal, R1, and transverse, R2, auto-relaxation rates, as well as the 15N–{1H} nuclear Overhauser effect, from which the dipolar cross-relaxation rate between the nitrogen-15 nucleus and its neighboring proton, sNH, can be extracted (13). In this review, I present basic protocols to set up and analyze these experiments, including the latest developments. In addition, we provide protocols for two additional experiments for measuring longitudinal and transverse cross-relaxation rates due to the cross-correlation of the nitrogen-15 chemical shift anisotropy (CSA) and the dipole–dipole (DD) coupling with its attached proton. These rates can be analyzed alongside the canonical set in order to evaluate the contribution of chemical exchange effects to transverse relaxation (14, 15). 1.1. Theory

We provide here the minimal theoretical background that is necessary to define the various terms that are used in this review. For a detailed understanding of relaxation theory, the reader should refer to some of the many reviews that have been published on the subject (16–19). Relaxation is the irreversible process through which a spin system evolves toward a steady state. Different elements of the density operator evolve with different auto-relaxation rates, and they convert into one another with well-defined cross-relaxation rates. The molecular processes underlying relaxation are the fluctuations of high-amplitude orientation-dependent interactions, such as dipole–dipole couplings and the anisotropy of the chemical shift. The amplitude of these interactions is known so that one can extract the characteristics of motions from relaxation rates. Relaxation rates do not depend directly on the correlation function of interaction Hamiltonians but on their Fourier transform: the spectral density function J(w). Here, we describe a protocol to measure the following relaxation rates for 15N nuclei in a 15N–1H pair. The longitudinal auto-relaxation rate, R1: R1 = D (6(1 + a 2 ) J (w N ) + 2 J (w H - w N ) + 12 J (w H + w N )). (1) The transverse auto-relaxation rate, R2:

9

Protein Dynamics by 15N Nuclear Magnetic Relaxation

143

R2 = D (4(1 + a 2 ) J (0) + 3(1 + a 2 ) J (w N ) + J (w H - w N ) + 6 J (w H ) + 6 J (w H + w N )).

(2)

The 1H–15N dipolar cross-relaxation rate, sNH: s NH = D (-2 J (w H - w N ) + 12 J (w H + w N )).

(3)

The CSA/DD transverse cross-correlated cross-relaxation rate, dtN: (4)

dtN = D a(8 J (0) + 6 J (w N )).

The CSA/DD longitudinal cross-correlated cross-relaxation rate, dlN: dlN = 12D aJ (w N ),

(5) 3 rNH

2

2 2 2 with D = 1 æ m0 ö  g H g N and a = - 2 B0 (s / / - s^ ) 6 20 çè 4p ÷ø rNH 3g H

; gH and gN

are the gyromagnetic ratios of the proton and nitrogen-15 nuclei, respectively; wH and wN are the Larmor angular frequencies of the proton and nitrogen-15, respectively (see Note 1); rNH is the internuclear distance; m0 is the permeability of free space; ħ is the Planck constant divided by 2p; B0 is the static magnetic field and s// and s ^ are the axial and perpendicular components of the anisotropic chemical shift tensor of the nitrogen-15 nucleus (which we consider to be axially symmetric). The 1H–15N dipolar cross-relaxation rate is measured indirectly from nuclear Overhauser effects. In this case, signal intensities are measured under two conditions: at the steady state under effective proton saturation Iss and at equilibrium Ieq. The ratio of these intensities is: (6) I ss / I eq = 1 + (g H s NH ) / (g N R1 ) In addition, we measure the transverse relaxation rate R2 under two different conditions, the more typical CPMG (Carr-PurcellMeiboom-Gill) train of echoes and by a single echo combined with continuous 1H composite pulse decoupling. The later rate includes, in most cases, the full contribution of chemical exchange to transverse relaxation, Rex, which is a probe of chemical reactions and motions on micro- to millisecond timescales (20, 21), a particularly relevant range for biological processes. In the case of fast exchange between two sites A and B (with a timescale tex and a difference of chemical shift between the two sites Dw), we have: Rex = p A pB Dw 2 t ex ,

(7)

where pA and pB are the populations of the two exchanging sites A and B respectively, pA + pB = 1. The focus of this review is not to describe how to determine the various parameters of the exchange process, which is well

144

F. Ferrage

presented in the following reviews (10, 22). Nevertheless, we describe a protocol to identify the presence of chemical exchange.

2. Materials 1. Appropriately labeled protein, preferably (2H, 15N) or (15N) [while studies of (2H, 15N, 13C) and (15N, 13C) labeled proteins are also possible] sample at ~1 mM concentration (see Note 2). 2. NMR spectrometer with a magnetic field strength of ³11.7 T (see Note 3). 3. nmrPipe: software to process, display and analyze 2D spectra (http://spin.niddk.nih.gov/NMRPipe/). 4. Curvefit: software to fit the relaxation decays and build-up curves (http://biochemistry.hs.columbia.edu/labs/palmer/ software/curvefit.html). 5. Grace: software to display the results of the Curvefit analysis (http://plasma-gate.weizmann.ac.il/Grace/).

3. Methods 3.1. Preliminary Set-up

A few preliminary procedures should be followed before starting the series of experiments. First, when all experiments are run on the same spectrometer, one should ensure the temperature calibration is very recent, in any case, it is safer to calibrate the temperature prior to recording all experiments. When data are collected by using two or more spectrometers, the exact match of temperatures should be verified directly on the same standard methanol sample on each spectrometer. It is more important to run all experiments at the exact same temperature than to have a small error on the nominal temperature of the experiments. 1. When running experiments on the first spectrometer, introduce a sample of perdeuterated methanol into the spectrometer. Set the temperature at the desired value according to the latest calibration. Let the sample equilibrate for 5 min. Match and tune the probe, and shim the magnet. 2. Run a simple 1D proton experiment with a small angle excitation (typically with a 1 ms 1H pulse at high power). Measure the difference in chemical shifts between the hydroxyl and methyl protons, Dd. The temperature is (23): T = -16.7467 Dd 2 - 52.5130 Dd + 419.1381.

(8)

9

Protein Dynamics by 15N Nuclear Magnetic Relaxation

145

3. Change the nominal temperature on the spectrometer and repeat step 2 until the desired temperature is reached. Write down the value of Dd. 4. When running experiments on another spectrometer, directly match the value of Dd. Second, pulses should be calibrated for each set of measurements collected on each spectrometer. The typical values obtained by using a standard sample should be used as a guess to start calibration but not directly to run experiments. 1. Introduce the protein sample in the spectrometer, let it equilibrate for a few minutes. Shim, tune, and match each channel that is going to be used. Set the carrier on the water signal and run a 1D proton spectrum with a short (1 ms) excitation at high power (see Note 4). Phase and use this spectrum as a reference. 2. Set the pulse duration to the expected value for a 360° flip angle. Collect a spectrum. Optimize the value of the pulse duration to get a null signal. Write down the duration of the corresponding 90° pulse. 3. Prepare a 1D version of a 15N–1H Heteronuclear Single Quantum Coherence (HSQC) experiment, preferably with a heteronuclear gradient echo. Set the calibrated duration for proton pulses and the expected value for 15N pulses. Make sure the carrier on the 15N channel is set at 117 ppm. Set the number of scans to 16. Run a 1D reference spectrum. If the signalto-noise is very high, reduce the number of scans. If it is low, increase accordingly. 4. Now, set the duration of the first 90° pulse on 15N to the expected duration for a 180° pulse. Run the experiment. Optimize to get a null signal. Write down the corresponding value for the calibrated 90° pulse. 5. If the sample is carbon-13 labeled, follow steps 3 and 4 with 13 C instead of 15N. The 13C carrier should be set at 35 ppm and the proton signal should be null between 2 and 3 ppm when the flip angle is 180°. 6. Alternatively, or, when using a carbon-13 and perdeuterated sample, run steps 3 and 4 with a 1D version of an HNCO experiment, where the first 13C pulse is adjusted to 180°. The carrier on the 13C channel should be set to 174 ppm (see Note 5). Third, the carrier, spectral width, and number of time points should be optimized in the 15N dimension (this should have been done during the assignment experiments). In particular, cross-correlated relaxation experiments on small proteins can have a long duration because of long phase cycles,

146

F. Ferrage

but are not signal-to-noise limited. Such optimization is an important time saver. 1. Set up a 2D HSQC experiment with a wide spectral width (larger than 30 ppm) so that only the arginine NeHe signals are folded. 2. Process the spectrum and adjust the spectral width and carrier. Folded peaks should not overlap with nonfolded peaks, and it is advised to keep about 1 ppm on each side of the spectrum with no peak. This process may require a few iterations. In the end, a spectral width between 16 and 24 ppm should be obtained for most small- or medium-size proteins. 3. This last step should be carried each time one runs experiments at a new B0 field. Record a final spectrum with the optimal carrier and spectral width as well as a large number of points in the indirect dimension. Process it with the lowest number of points that provides good resolution of all peaks of interest. Use this value for all experiments. Be careful not to underestimate the number of points, it may save 10% of the time but you may also have to repeat the full series of experiments. 3.2. Auto-Relaxation Rates Measurements 3.2.1. Longitudinal Relaxation Rate

The sequences presented in Fig. 1a, b should be employed for this experiment. There is no fully satisfactory combination of water-flip back schemes and suppression of CSA/DD cross-correlated crossrelaxation pathways during the relaxation delay. One solution is to saturate the water resonance in each scan and use a long recovery delay between scans. To saturate the water resonance, drop the water-flip back pulse at the end of the first INEPT in Fig. 1a and use a strong gradient G2. In that case, the two shaped pulses in Fig. 1b can be substituted either by composite pulse decoupling (see Fig. 1c) or a series of proton 180° pulses (see Fig. 1d). Here, we use a scheme that can be used when no amide resonance is lying too close to the water resonance. It is one of the standard sequences available on Bruker spectrometers. 1. Set pulse durations and amplitudes according to the spectrometer specifications, the calibration and the description of the pulse sequence in Fig. 1a, b. Set delays according to the pulse sequence. Calibrate proton shape pulses according to the spectrometer-specific protocol. 2. Set the recovery delay between scans to a large value, usually 2 s; 3 s can be used if spectrometer time is not a problem. 3. For a well-behaved protein, start with eight scans (16–32 for low-concentration samples). Set the relaxation delay to a low value (we use 20 ms). Run the first 1D experiment. Phase and save this spectrum. 4. Set the relaxation delay to a value close to the expected average T1. At 300 K, this is 500 ms for a globular 100-residue protein and 700 ms for a 150-residue protein. Acquire a new spectrum.

9

Protein Dynamics by 15N Nuclear Magnetic Relaxation

147

Fig. 1. Pulse sequences used for measuring 15N auto-relaxation rates. (a) General scheme. (b) Relaxation sequence for measuring the longitudinal relaxation rate R1; (c) relaxation sequence for measuring the transverse relaxation rate under a single echo R2echo; (d) relaxation sequence for the measurement of the transverse relaxation rate under a CPMG echo train R2CPMG. All narrow (filled) and (wide ) open rectangles represent 90° and 180° pulses, respectively. Pulse phases are along the x-axis of the rotating frame unless otherwise mentioned. Proton composite pulse decoupling during the delay t1 was performed with a GARP scheme and a radio frequency field (rf) amplitude of 1 kHz. The 13C channel pulse was a 500 ms smoothed CHIRP pulse (37), with a sweep of 60 kHz on a 600 MHz spectrometer; the carrier at the center of the pulse was 110 ppm. Composite-pulse decoupling during acquisition was performed on the 15N channel with a GARP scheme (38) and an rf amplitude of 1,090 Hz. The delay ta is 2.56 ms; the delay tb can be adjusted around 5 ms. The phase cycles were: f1 = {x, −x }; f3 = {x, x, −x, −x }; f4 = {x, x, −x, −x }; f5 = {−y, −y, y, y }; facq = {x, −x, −x, x }. When relaxation block (b) is used, f2 = { y, y, y, y, −y, −y, −y, −y } and facq = {x, −x, −x, x, −x, x, x, −x }. The amplitude profile of the pulsed field gradient was a sine bell shape. Their durations and peak amplitudes over the x, y, and z orientations (when triple axis gradients are available) were, respectively: G1; 600 ms, 9.5 G/cm, 9.5 G/cm, 0; G2; 1 ms, 0, 0, 30 G/cm; G3; 600 ms, 15 G/cm, –15 G/cm, 0; G4; 1 ms, 0, 0, 40 G/cm; G5; 1 ms, 0, 0, 8.1 G/cm. Coherence selection was achieved by inverting the amplitude of the gradient G4 and phase f1. (b) The carrier is placed at 8.2 ppm during the relaxation block; gray bell-shaped pulses are 1.6 ms Q3 Gaussian cascade pulses at 600 MHz (39) (see Note 9). (c) WALTZ-16 decoupling should be used for 1H decoupling during the relaxation block (40). See text for how to choose the relaxation delays. (d) Gray rectangles are 180° pulses, depending on the spectrometer and probe, they should be either at high power or less, but should not be longer than 100 ms. t should be set to 500 ms.

148

F. Ferrage

5. Compare the intensities and repeat with adjusted delays until the ratio of intensities is about 0.3. We call this delay tmax + 20 ms. 6. The best sampling of the decay is achieved with an even decay of intensities between each time point. If n different relaxation delays are to be acquired, the intensity of the j-th experiment should be: I j = I 1 (1 - 0.7( j - 1) / (n - 1)).

(9)

We expect a mono-exponential decay with an average rate R1av and we know that:

(

)

exp -R1av t max = 0.3 Û R1av = - ln 0.3 / t max

(

)

(10)

and exp -R1avt j = I j .

(11)

t j = (t max / ln0.3)ln(1 - 0.7( j - 1) / (n - 1)).

(12)

So that

7. A typical number of relaxation delays is 8. This number can be reduced to 6, even 4 in cases where each spectrum takes a very long time to acquire. Record all experiments in an interleaved manner, changing the relaxation time once each full 1D spectrum is acquired. This requires a small modification of standard pulse sequence programs. When using standard pulse sequences, the sequence of delays should be: t1, tn, t2, tn−1, t3, tn−2, etc., so that the decay of intensities would not appear mono-exponential if the quality of the spectra was to decay, because of a bubble for instance, which would otherwise be reflected by a higher, erroneous rate. Similarly, at least t1 and, if possible t3 should be repeated to identify a possible decay in the quality of spectra. If this decay is larger than the error bars, the dataset should be discarded. 8. Before starting the series of experiments, set the number of “dummy scans” in the first experiment to a very high value (at least 256, preferably 512). This leaves enough time for the temperature control system of the spectrometer to reach equilibrium. 9. Import the spectra into nmrPipe (24). Process them with typical parameters, the window function should be a sine-bell function, the power should be set to 2 and the shift parameter between 0.5 and 0.35 in each dimension (the lower the value, the more resolution). Truncation artifacts in the 2D spectrum should be avoided at all cost, as they may contaminate the intensity of neighboring peaks. 10. Peak pick the first spectrum, or export a peak list from the assignment software, save the peak list as peakX_0.tab, where X is the number of the spectrum. To extract the intensity of the peaks with the best possible accuracy, use the nonlinear line

9

Protein Dynamics by 15N Nuclear Magnetic Relaxation

149

shape analysis tool nlinLS, which is provided with nmrPipe. First, measure the full width, in points, of the peaks in each dimension: WH and WN. Then, run the following line: nlinLS -in peakX_0.tab -out peakX_1.tab -data spectrumX.ft2 -w WH/6 WN/6 -delta X_AXIS WH/10 Y_AXIS WN/10. This supposes that the proton dimension is the X dimension of the spectrum. The option w defines the region, where the peak is fitted (the current command specifies a square of dimensions WH and WN); the option delta adds a constraint on the following parameters, here the position of each peak. 11. It is likely that nlinLS will initially provide a series of error messages, most often indicating bad convergence of the fit. In spite of these error messages, the output file will be of better quality than the input file and should be used as an input for a second round of analysis with nlinLS. The process is iterative and may require up to five iterations for convergence and a proper fit with no error message. If the width of a peak is diverging from typical values in the output file, an additional constraint can be added to the peak width parameters, WX and WY. Peaks may also be added or excluded from clusters to improve convergence. Use the final, error-free list obtained from the first spectrum as the first input for all other fits. 12. Run this iterative procedure for each spectrum. 13. Collect the intensities and peak numbers with the getCols tool, which is provided with nmrPipe. nlinLS provides both peak intensities and volumes. Only the intensities should be used. Although the precision should be better than the spectral noise, this turns out to be a good estimate of the error in the intensities. Use this value as the error for all the peaks of each spectrum. 14. Using a short script or a spreadsheet editor, prepare a table with peak numbers in the first column, then the intensities of peaks in the series of spectra in columns 2, 4, 6, etc., and the respective error in columns 3, 5, 7, etc. 15. Run a short script to create a series of Curvefit input files with names: N.in, where N is the peak number (see Note 6). 16. Run the command: batch_curve in. The fit for each peak number appears (see Fig. 2). It is very important to check every single fit and identify errors that may come from improper intensity fits or other potential errors. Write down the peaks and spectra that lead to apparently wrong intensities and inspect the spectra for potential errors. There may be no error. 3.2.2. Transverse Relaxation Rates with a Single Echo

The sequence presented in Fig. 1b should be employed for this experiment. Only relaxation delays that lead to full cycles of the proton composite pulse decoupling (CPD) should be used, which modifies slightly the protocol from the measurement of longitudinal relaxation rates.

150

F. Ferrage

Fig. 2. Example of a longitudinal relaxation decay curve as shown by Grace during the Curvefit procedure.

1. Most parameters are identical to those used in longitudinal relaxation measurements. To cover the entire amide region the carrier for the proton CPD is set to 8.2 ppm and the amplitude to about 2 × x Hz; where x is the proton Larmor frequency of the spectrometer in MHz. Set the CPD scheme to WALTZ-16 (see Note 7). The half relaxation delay T should be set to multiples of 96 × tcpd, where tcpd is the duration of a 90° pulse at the decoupling power. 2. Contrary to other sequences, proton CPD with a carrier on the amides prevents a proper control of the polarization of water. Set the radiofrequency amplitude to zero for the water flip back pulse following the first INEPT and use a long recovery delay (at least 2 s) in between scans. 3. Follow steps 2–7 of the protocol for longitudinal relaxation (Subheading 3.2.1) with the following changes. Start with the shortest relaxation delay, i.e., zero. The fact that the total relaxation delay has to be a multiple of 192 × tcpd, severely limits the choice of possible delays, use all accessible delays between zero and tmax. 4. To obtain the transverse relaxation rate under a single echo R2echo, the analyses of spectra are exactly the same as that described in steps 8–15 of the preceding protocol (Subheading 3.2.1).

9 3.2.3. Transverse Relaxation Rates with a CPMG Scheme

Protein Dynamics by 15N Nuclear Magnetic Relaxation

151

For the sake of consistency, this protocol is described here. However, this is the most challenging experiment as heating from high radiofrequency fields may lead to bubble formation and sample degradation. It is strongly advised to run this experiment as the last one of the series. 1. Most parameters are identical to those used in longitudinal relaxation measurements. Set the power level for 15N 180° pulses during the CPMG echo train. This experiment is one of the most demanding that can be run on a high-resolution probe. The maximum power (i.e., shortest nitrogen-15 pulses) that can be employed during a CPMG echo train is usually provided to users for each probe on each spectrometer. If no value is recommended, ask the person in charge of the spectrometer maintenance. If you are in charge of the spectrometer maintenance, ask the manufacturer. Often, particularly on cryogenic probes, these pulses should not be applied at full power. For proper accuracy of the experiment, 180° pulses should be no longer than 100 ms. 2. In addition to Fig. 1d, include a temperature-compensation loop at the beginning of the sequence. At the end of the recycling delay, include a series of far off-resonance (200 kHz works fine) 15N 180° pulses at the amplitude of the CPMG echo train. Do not include 1H 180° pulses as these would alter the 1 H longitudinal polarization. The duration of this train should be such that the total number of 15N 180° pulses is constant whatever the relaxation delay is. An additional delay, equal to the relaxation delay, should also be added so that the total duration for the recovery of proton polarization is constant. 3. Follow steps 2–7 of the protocol for longitudinal relaxation (Subheading 3.2.1). Zero can be used as the shortest relaxation delay. 4. Use a long recovery delay (preferably 2 s) and a large number of dummy scans (512) to let the temperature control system reach equilibrium before the first experiment. 5. Follow steps 9–16 of the R1 analysis (Subheading 3.2.1) to obtain transverse relaxation rates under a CPMG train, R2CPMG.

3.3. 15N–{ 1H} Nuclear Overhauser Effect Measurements

The sequence presented in Fig. 3 should be used for this experiment. It displays a series of improvements introduced recently (25–27). The two experiments (under effective proton saturation and at equilibrium) have to be run in an interleaved manner. 1. Set up the saturation scheme (Fig. 3b). The saturation element is symmetric, 1H pulses have a 180° flip angle, the carrier is placed in the amide region (8.2 ppm), the interpulse delay tNOE should be a multiple of 1/JNH, where JNH is the 1H–15N one-bond scalar coupling constant. In globular proteins at low pH, tNOE = 22 ms

152

F. Ferrage

Fig. 3. Pulse sequence used for recording steady-state 15N–{1H} nuclear Overhauser effects. For each measurement, reference and steady-state experiments have to be recorded in an interleaved manner. In the reference experiment, one should run the part of the pulse program displayed in box (a). At the end of the recovery delay TNOE = 10 s (or more), the proton carrier is placed on resonance with the water signal and a very selective water-flip back pulse is applied (3 ms sinc shaped or longer). To record steady-state experiments, the boxed sequence in (a) is substituted by the scheme shown in (b) for the effective saturation of amide proton resonances. After an optional delay T¢NOE = 2 s for stable detection of the lock signal, the proton carrier is placed in the center of the amide region (at 8.2 ppm) as shown by the arrow labeled by N. The motif [delay tNOE/2 – 180° pulse – delay tNOE/2] is repeated nNOE times. The interpulse delay, tNOE, is typically 22 ms (11 ms may also be used, see text). The rf amplitude for the pulses should be 7.5 kHz at a 500 MHz Larmor frequency and 9 kHz at 600 MHz. A gradient G1 is applied at the end of the last tNOE/2 delay to suppress all transverse components of the proton polarization. The carrier was moved on-resonance with the water signal as indicated by the W arrow. The number of cycles, nNOE, was set so that the total duration for effective saturation was 4 s. All narrow (filled ) and wide (open) rectangles represent 90° and 180° pulses, respectively. Pulse phases are along the x-axis of the rotating frame unless otherwise mentioned. Proton composite pulse decoupling during the delay t1 was performed with a GARP scheme and an rf amplitude of 1 kHz. Composite-pulse decoupling during acquisition was performed on the 15N channel with a GARP scheme (38) and an rf amplitude of 1,090 Hz. The delay ta is 2.56 ms. The phase cycles were: f1 = {y, –y}; f2 = {x, x, –x, –x}; f3 = {x, x, –x, –x }; f4 = {−y, –y, y, y }; facq = {x, –x, –x, x }. The amplitude profile of the pulsed field gradient was a sine bell shape. The durations and peak amplitudes over the x, y, and z orientations were, respectively: G1; 600 ms, 15 G/cm, 15 G/cm, 0; G2; 1 ms, 0, 0, 25 G/cm; G3; 1 ms, 0, 0, 40 G/cm; G4; 1 ms, 0, 0, 8.1 G/cm. Coherence selection was achieved by inverting the amplitude of the gradient G3 and phase f4.

is a safe value, but tNOE = 11 ms should be used in disordered proteins or on high pH samples, where proton exchange with the solvent is faster. The amplitude of these radiofrequency pulses should be 1.5 × X kHz, where X is the 1H Larmor frequency in MHz divided by 100 (e.g., 9 kHz on a 600 MHz spectrometer). 2. Set up the duration of the saturation scheme. The total duration of the saturation is linked to the 15N R1 rates, but not to the 1H relaxation rates, since the present scheme ensures an effective saturation of protons after each saturation element. If the longitudinal relaxation experiments have been fully analyzed, identify the lowest value of all 15N R1 rates. Set the value of nNOE so that the total duration of the saturation scheme is 4/R1. If only an estimate of the average longitudinal relaxation rates R1av is available from a comparison of 1D spectra (steps 3–5 of the protocol for the measurement of R1 rates; Subheading 3.2.1),

9

Protein Dynamics by 15N Nuclear Magnetic Relaxation

153

set the total duration of saturation to at least 6/R1av and preferably 8/R1av, if time permits. 3. Set up the reference experiment (Fig. 3a). The soft water-flip back pulse should be long (at least a 3 ms sinc-shaped pulse, and longer for spectrometers with a Larmor frequency under 700 MHz) as it should not touch amide resonances closest to the water resonance. The total duration of the recovery delay, TNOE, should be long. In this case, the spin system should be at equilibrium, including the protons. The use of a water-flip back scheme permits the use of significantly shorter recovery delays. In general, these delays should be no shorter than 10 s. It is strongly advisable to spend up to an hour to compare the intensity of the first 1D spectrum (obtained with t1 = 0) for various values of TNOE and identify for which value of TNOE the signal has reached its maximum value. Since the signal-to-noise ratio will be too low to properly evaluate the difference between 97% and 100% of the signal, add at least 2 s to the value of TNOE where the signal appears to have reached its maximum value. 4. The total number of scans should be at least 24 and up to 64. The experiment should last between 12 and 24 h. If longer acquisition times are necessary, reduce the number of scans for each experiment and run a series of experiments no longer than 24 h. The number of dummy scans should not be greater than 64. 5. Follow steps 9–14 of the protocol for longitudinal relaxation (Subheading 3.2.1) to obtain intensities. 6. Compute the NOE ratio (Eq. 6). Although it may not be necessary for the following analysis, derive the dipolar crossrelaxation rate from the NOE ratio using Eq. 6. 3.4. Measurements of CSA/DD CrossCorrelated CrossRelaxation Rates

These experiments are less commonly run in spite of their great utility. The set up turns out to be straightforward after more typical relaxation experiments have been run. 1. Set pulse durations and amplitudes according to the spectrometer specifications, the calibration and the description of the pulse sequence, which is shown in Fig. 4. Set delays according to the pulse sequence. Calibrate proton shape pulses according to the spectrometer-specific protocol. Most of this part has already been set up in the preceding experiments. 2. For both experiments, the minimum number of scans imposed by the phase cycle is higher, leading to very long experimental times when high resolution data needs to be collected (large number of t1 points). On the other hand, shorter recycle delays can be used, typically between 1 and 1.5 s with little loss of sensitivity and no loss of accuracy. 3. These experiments use symmetrical reconversion (15, 28). This means that all four relaxation pathways in a two-operator space are detected. These four experiments have to be run in an

154

F. Ferrage

Fig. 4. Pulses sequences for measuring CSA/DD cross-correlated cross-relaxation rates. (a) General scheme; (b) relaxation block for measuring the transverse cross-relaxation rate; (c) relaxation block for measuring the longitudinal cross-relaxation rate. Only specific elements are detailed here. The delay d is equal to the shortest value of t1 so that the first effective value of t1 is zero. The phase cycles were: f1 = { y, –y }; f2 = {x, x, x, x, –x, –x, –x, –x}; f3 = {x, x, x, x, x, x, x, x, –x, –x, –x, –x, –x, –x, –x, –x}; f4 = {x, x, x, x, x, x, x, x, x, x, x, x, x, x, x, x, –x, –x, –x, –x, –x, –x, –x, –x, –x, –x, –x, –x, –x, –x, –x, –x}; f5 = {x}; f6 = {x, x, –x, –x}; f¢6 = {x, x, y, y}; facq = {x, –x, –x, x, –x, x, x, –x, –x, x, x, –x, x, –x, –x, x, –x, x, x, –x, x, –x, –x, x, x, –x, –x, x, –x, x, x, –x}. Gradient durations and peak amplitudes over the x, y, and z orientations were, respectively: G1; 600 ms, 15 G/ cm, 15 G/cm, 0; G2; 600 ms, 6.5 G/cm, 0, 0; G3; 2 ms, 0, 0, 40 G/cm; G4; 1 ms, –9.5 G/cm, 9.5 G/cm, 0 G/cm; G5; 600 ms, 3.5 G/cm, 0 G/cm, 14.5 G/cm; G6; 1 ms, –35 G/cm, –35 G/cm, –35 G/cm; G7; 1 ms, 35 G/cm, 35 G/cm, 35 G/cm. Frequency sign discrimination was performed using States-TPPI.

9

Protein Dynamics by 15N Nuclear Magnetic Relaxation

155

interleaved manner (see Note 8). Remember to set the number of points in the indirect dimension to the proper value (i.e., eight times the number of complex time points). 3.4.1. Transverse Cross-Correlated Cross-Relaxation Rates

1. Set up the relaxation delay(s). The limiting factor for sensitivity is the intensity in the cross-relaxation experiments (pathways II and III in Fig. 4a). This intensity is maximum when the relaxation delay is equal to the auto-relaxation time. The optimal relaxation delay typically lies between 30 and 80 ms. Explore this interval in 20 ms steps. The intensity versus time curve is usually flat around the maximum. When two relaxation delays appear to give the same maximum intensity, first check a relaxation delay at the midpoint between the two. If the intensity is better, keep this delay. If the intensity is the same, choose the shortest delay so the intensity in the auto-relaxation experiments will be better. 2. This experiment can be run with a single relaxation delay. If time permits, repeat the optimal relaxation delay and run another delay with sufficient intensity on the cross-relaxation experiments. 3. Follow steps 9–14 of the longitudinal relaxation protocol (Subheading 3.2.1) to obtain the intensities of the four interleaved experiments. The following quantity should be computed: S (4T ) = I II (4T )I III (4T ) / I I (4T )I IV (4T ).

(

)

S (4T ) = tanh dNt 4T .

(13) (14)

4. If only one relaxation delay is recorded, simply invert Eq. 14. If several time points are recorded, compile the values of S and the corresponding errors in a text file and follow steps 15 and 16 of the longitudinal relaxation protocol (Subheading 3.2.1). The header of each N.in file should set the hyperbolic tangent mode for the fit to evaluate dtN . 3.4.2. Longitudinal Cross-Correlated Cross-Relaxation Rates

1. Set up the relaxation delay(s). The constraints are similar to those for the transverse cross-relaxation rate (Subheading 3.4.1). A difference is that the cross-relaxation experiments are now number I and IV because of the conversion between longitudinal polarization and two-spin order in the middle of the crossrelaxation delay. The optimal delay should be found between 80 and 200 ms. However, particularly in nondeuterated proteins, there is a strong advantage to using shorter delays. If not, one has to calculate a correction for the rates that takes into account the effects of proton–proton cross-relaxation (15). In that case, at least two well-separated time points should be recorded. 2. This experiment can be run with a single relaxation delay if it is short and/or the protein is deuterated. To make sure that the correction is small, it is advisable to run two experiments with different delays.

156

F. Ferrage

3. Follow steps 3–4 for the transverse cross-relaxation protocol (Subheading 3.4.1 ). Figure 5 shows the build-up of the symmetrical reconversion observable for longitudinal crosscorrelated cross-relaxation. These data were recorded with a small deuterated protein, thus allowing the use of long crossrelaxation delays. 4. If two or more relaxation delays are recorded on a nondeuterated sample, evaluate the correction factor, D, for each time point, and compute the ratio: 2

(15)

I IV / I I = C exp(-D 4T ) / (1 + 32d1N DT 3 / 3.

From these2 ratios, fit the value of D. In most cases, evaluate the factor 32d1N DT 3 / 3 , it should be much smaller than 1 so that D can be obtained from a simple exponential fit. 5. Compute for each time point: dlN T = atanh

(II

I IV

)(

)

/ I II I III / 1 + 2D 2T 2 / 3 .

(16)

Proceed to a linear fit to obtain dlN . 3.5. Analysis and Interpretation of 15N Relaxation Rates

The analysis of this dataset closely follows the one presented by Kroenke et al. (14). The use of more accurate measurements of nuclear Overhauser effects and longitudinal CSA/DD cross-correlated

Fig. 5. Example of a longitudinal cross-correlated cross-relaxation build-up curve as shown by Grace during the Curvefit procedure.

9

Protein Dynamics by 15N Nuclear Magnetic Relaxation

157

cross-relaxation rates as well as the comparison of transverse relaxation rates measured under a CPMG echo train and a single echo are the main improvements. 3.5.1. Identification of Chemical Exchange

1. Compute the transverse relaxation rate expected in the absence of exchange R20, for each backbone 15N nucleus: R20 = (R1 - 1.25s NH )dtN / dlN - 1.08s NH .

(17)

2. Derive the exchange contribution to the transverse relaxation rate measured with a CPMG sequence: Rexecho = R2echo - R20 .

(18)

3. Derive the exchange contribution to the transverse relaxation rate measured with a single echo: RexCPMG = R2CPMG - R20 .

(19)

4. Figure 6 shows, as an example, a plot of RexCPMG for Calbindin D9k at 296 K. All but one residue shows contributions of exchange to transverse relaxation rates during CPMG that are significantly larger than 1/s. This confirms that motions of the backbone of Calbindin D9k are very limited on timescales in the 10 ms/ms range. Only L6 and T45 show small but significant contributions of chemical exchange to transverse relaxation. RexCPMG should never be negative, so the presence of a few values around −0.7/s (with an expected error smaller than 0.2/s) indicates that the accuracy of the method is not as good as its precision. In this particular case, the use of a carbon-13 labeled sample makes the derivation of R20 less accurate since contributions of 15 N–13C dipolar interactions to 15N auto-relaxation also have to be taken into account and predicted. 5. The comparison of RexCPMG and Rexecho provides insights on the order of magnitude of the timescales of the exchange processes. If a nonzero value of Rexecho is determined while RexCPMG is zero, the exchange process is slower than 1 ms. If both values are identical, the exchange process is faster than 1 ms. If RexCPMG is smaller than Rexecho but significantly larger than zero, the timescale of the exchange process is close to 1 ms or two exchange processes take place, one faster and one slower than 1 ms. Note that RexCPMG should never be larger than Rexecho. 3.5.2. A Note on the Model-Free Analysis of 15N Relaxation Rates

The ensemble of rates can be used as the input for a model-free analysis (29). A series of software are available: modelfree (30); fast modelfree (31); tensor2 (32); and dynamics (33) are good examples. I do not describe here the use of such software. The only point that discussed is the choice of the ensemble of relaxation rates that should be employed. The above-mentioned softwares are designed to use the longitudinal relaxation rate R1, the transverse relaxation

158

F. Ferrage

Fig. 6. Contribution of chemical exchange processes to transverse relaxation in a single echo experiment RexCPMG in Calbindin D9k at 296 K.

rate R2CPMG, and the NOE ratio as inputs. In most cases, these rates should be used. One of the most difficult tasks of the analysis is the detection of an exchange contribution to transverse relaxation rates R2CPMG. When a nonzero contribution RexCPMG is identified, the rates are at best fit with a simpler model for fast motions. When a protein shows significant contributions of chemical exchange to the relaxation of many 15N nuclei, this can be detrimental to the overall quality of the analysis of hydrodynamic properties and fast local motions. The analysis described herein provides exchangefree transverse relaxation rates R20. If, and only if, the ensemble of RexCPMG rates shows a very flat baseline around zero (see for instance ref. 15), the R20 rates can be employed instead of the R2CPMG rates in the model-free analysis.

4. Notes 1. Note that since the gyromagnetic ratio of 15N is negative, these two frequencies have opposite signs. Keeping this in mind helps understand the analysis of relaxation rates. 2. Particular care should be given to the sample conditions for relaxation studies. The stability of the sample is very important since data are collected as series of experiments in several days (or possibly more) and will be used together for the analysis.

9

Protein Dynamics by 15N Nuclear Magnetic Relaxation

159

We advise against running such a series of experiments on samples with very short (2 days or less) lifetimes unless high reproducibility can be achieved in sample preparation and several samples can be used to collect the ensemble of data discussed here. In order to ensure the quality of the data and a safe interpretation of hydrodynamic properties and chemical exchange, it is advised to verify the concentration dependence of relaxation rates which can be affected by aggregation and oligomerization. The transverse relaxation rate measured with a single echo is very sensitive to such processes and can be used to find the optimal concentration (34). One of the common problems that can arise during the weeklong data collection is the appearance of bubbles, which can degrade the field homogeneity and decrease the signal intensity, especially when a Shigemi tube is being used (as bubbles will be trapped in the detection volume). It is advisable that samples should be degassed before recording long relaxation experiments. 3. The series of experiments described herein is most easily adapted for small and middle-size proteins (less than 200 amino acids, monomeric). In most cases, the data are easily recorded on any spectrometer (B0 ³ 11.7 T) when a concentration around ~1 mM can be used. When the samples are concentration limited, the use of spectrometers equipped with cryogenic probes should be preferred, particularly at low fields (B0 £ 14.1 T). When using cryogenic probes, optimal homogeneity of the magnetic field B0 should be reached in order to ensure proper suppression of the water signal. Probes equipped with triple-axis gradients enhance the water suppression (35). When only z-axis gradients are available, the sign of gradients should be chosen such that the phase of the water transverse polarization builds up (36). For instance, two pulsed field gradients separated in the pulse sequence by a single 180° pulse on the proton channel should have opposite signs in the manner of bipolar gradients. 4. Using a small angle to start helps reduce the effects of radiation damping. It is also practical, since the signal is directly proportional to the flip angle (modulo 360°), one can use the intensity in this first experiment as a meter for the deviations of following pulses from 360°. For example, if the intensity in the first attempt at a 360° pulse is positive and about twice as large as that of the 1 ms pulse experiment, it is likely that a pulse 2 ms shorter will be an exact 360° pulse. 5. If, as described here, only adiabatic pulses are used on the 13C channel, pulse calibration on this channel is optional, since the quality of the inversion by the adiabatic pulse will not be affected by a small deviation of the rf amplitude from its nominal value. 6. Sample script for an exponential decay:

160

F. Ferrage

9

Protein Dynamics by 15N Nuclear Magnetic Relaxation

161

The input file is a text file made of 25 columns; the first one is the peak/residue number while columns 2–25 are the intensity and error for the 12 longitudinal relaxation experiments. 7. WALTZ-16 is not as good a decoupling scheme as GARP. However, its basis element is much shorter. Therefore, when very low amplitude decoupling is performed, or when short durations are favored (like in the present case) WALTZ-16 should be used instead of GARP. 8. Since the four experiments for each relaxation delay are run in an interleaved manner, there is no need to record different relaxation delays in series. If doubts arise about the stability of the sample, run one set for a given value of the relaxation delay. Additional datasets for other relaxation delays (or repeats of the same) can be acquired after the full series of experiments. 9. When working on a spectrometer with a different Larmor frequency, all pulse durations should be scaled with the inverse of the field (e.g., 1.6 ms at 600 MHz corresponds to 1.2 ms at 800 MHz). There are a few exceptions: high-power, 15N CPMG, and 15N decoupling radiofrequency amplitudes usually do not scale up with B0.

Acknowledgments I am grateful to Geoffrey Bodenhausen, David Cowburn, Ranajeet Ghose, Arthur G. Palmer, and Philippe Pelupessy for their many contributions to my training, from hands-on practice to many insightful discussions. I thank Mikael Akke for the sample of Calbindin D9k and Kaushik Dutta for carefully reading this manuscript. References 1. Mittermaier, A., and Kay, L. E. (2006) Review – New tools provide new insights in NMR studies of protein dynamics. Science 312, 224–228. 2. Palmer, A. G. (2004) NMR characterization of the dynamics of biomacromolecules. Chem. Rev. 104, 3623–3640. 3. Massi, F., Wang, C. Y., and Palmer, A. G. (2006) Solution NMR and computer simulation studies of active site loop motion in triosephosphate isomerase. Biochemistry 45, 10787–10794. 4. Tjandra, N., Feller, S. E., Pastor, R. W., and Bax, A. (1995) Rotational Diffusion Anisotropy of Human Ubiquitin from N-15 NMR Relaxation. J. Am. Chem. Soc. 117, 12562–12566. 5. Akke, M., Brüschweiler, R., and Palmer III, A. G. (1993) NMR Order Parameters and Free Energy:

An Analytical Approach and Its Application to Cooperative Ca2+ Binding by Calbindin Dgk. J. Am. Chem. Soc. 115, 9832–9833. 6. Frederick, K. K., Marlow, M. S., Valentine, K. G., and Wand, A. J. (2007) Conformational entropy in molecular recognition by proteins. Nature 448, 325–329. 7. Kuszewski, J., Gronenborn, A. M., and Clore, G. M. (1999) Improving the packing and accuracy of NMR structures with a pseudopotential for the radius of gyration. J. Am. Chem. Soc. 121, 2337–2338. 8. Ryabov, Y., and Fushman, D. (2007) Structural Assembly of Multidomain Proteins and Protein Complexes Guided by the Overall Rotational Diffusion Tensor. J. Am. Chem. Soc. 129, 7894–7902.

162

F. Ferrage

9. Ryabov, Y. E., and Fushman, D. (2007) A model of interdomain mobility in a multidomain protein. J. Am. Chem. Soc. 129, 3315–3327. 10. Palmer, A. G., and Massi, F. (2006) Characterization of the dynamics of biomacromolecules using rotating-frame spin relaxation NMR spectroscopy. Chem. Rev. 106, 1700–1719. 11. Palmer, A. G. (2004) NMR characterization of the dynamics of biomacromolecules. Chem. Rev. 104, 3623–3640. 12. Vallurupalli, P., Hansen, D. F., and Kay, L. E. (2008) Structures of invisible, excited protein states by relaxation dispersion NMR spectroscopy. Proc. Natl. Acad. Sci. U.S.A. 105, 11766–11771. 13. Kay, L. E., Torchia, D. A., and Bax, A. (1989) Backbone Dynamics of Proteins as Studied by N-15 Inverse Detected Heteronuclear NMRSpectroscopy – Application to Staphylococcal Nuclease. Biochemistry 28, 8972–8979. 14. Kroenke, C. D., Loria, J. P., Lee, L. K., Rance, M., and Palmer III, A. G. (1998) Longitudinal and Transverse H-1-N-15 Dipolar N-15 Chemical Shift Anisotropy Relaxation Interference: Unambiguous Determination of Rotational Diffusion Tensors and Chemical Exchange Effects in Biological Macromolecules. J. Am. Chem. Soc. 120, 7905–7915. 15. Pelupessy, P., Ferrage, F., and Bodenhausen, G. (2007) Accurate Measurement of Longitudinal Cross-Relaxation Rates in Nuclear Magnetic Resonance. J. Chem. Phys. 126, 134508. 16. Korzhnev, D. M., Billeter, M., Arseniev, A. S., and Orekhov, V. Y. (2001) NMR Studies of Brownian Tumbling and Internal Motions in Proteins. Prog. Nucl. Magn. Reson. Spectrosc. 38, 197–266. 17. Luginbuhl, P., and Wuthrich, K. (2002) Semiclassical nuclear spin relaxation theory revisited for use with biological macromolecules. Prog. Nucl. Magn. Reson. Spectrosc. 40, 199–247. 18. Nicholas, M. P., Eryilmaz, E., Ferrage, F., Cowburn, D., and Ghose, R. (2010) Nuclear spin relaxation in isotropic and anisotropic media, Prog. Nucl. Magn. Reson. Spectrosc. 57, 111–158. 19. Cavanagh, J., Fairbrother, W. J., Palmer III, A. G., Rance, M., and Skelton, N. J. (2006) Protein NMR Spectroscopy: Principles and practice, Academic Press, San Diego. 20. Wang, L. C., Pang, Y. X., Holder, T., Brender, J. R., Kurochkin, A. V., and Zuiderweg, E. R. P. (2001) Functional dynamics in the active site of the ribonuclease binase. Proc. Natl. Acad. Sci. U.S.A. 98, 7684–7689. 21. Wang, C. Y., and Palmer, A. G. (2003) Solution NMR methods for quantitative identification of

chemical exchange in N-15-labeled proteins. Magn. Reson. Chem. 41, 866–876. 22. Kempf, J. G., and Loria, J. P. (2004) Measurement of Intermediate Exchange Phenomena. Meth. Mol. Biol. 278, 185–231. 23. Findeisen, M., Brand, T., and Berger, S. (2007) A H-1-NMR thermometer suitable for cryoprobes. Magn. Reson. Chem. 45, 175–178. 24. Delaglio, F., Grzesiek, S., Vuister, G. W., Zhu, G., Pfeifer, J., and Bax, A. (1995) NMRPipe: a Multidimensional Spectral Processing System Based on UNIX Pipes. J. Biomol. NMR 6, 277–293. 25. Ferrage, F., Piserchio, A., Cowburn, D., and Ghose, R. (2008) On the measurement of N-15-{H-1} nuclear Overhauser effects. J. Magn. Reson. 192, 302–313. 26. Ferrage, F., Cowburn, D., and Ghose, R. (2009) Accurate Sampling of High-Frequency Motions in Proteins by Steady-State 15N-{1H} Nuclear Overhauser Effect Measurements in the Presence of Cross-Correlated Relaxation. J. Am. Chem. Soc. 131, 6048–6049. 27. Ferrage, F., Reichel, A., Battacharya, S., Cowburn, D., and Ghose, R. (2010) On the measurement of N-15-{H-1} nuclear Overhauser effects. 2. Effects of the saturation scheme and water signal suppression. J. Magn. Reson. 207, 294–303. 28. Pelupessy, P., Espallargas, G. M., and Bodenhausen, G. (2003) Symmetrical reconversion: measuring cross-correlation rates with enhanced accuracy. J. Magn. Reson. 161, 258–264. 29. Lipari, G., and Szabo, A. (1982) Model-Free Approach to the Interpretation of Nuclear Magnetic Resonance Relaxation In Macromolecules 1. Theory and Range of Validity. J. Am. Chem. Soc. 104, 4546–4559. 30. Mandel, A. M., Akke, M., and Palmer III, A. G. (1995) Backbone Dynamics of Escherichia coli Ribonuclease HI : Correlations with Structure and Function in an Active Enzyme. J. Mol. Biol. 246, 144–163. 31. Cole, R., and Loria, J. P. (2003) FAST-Modelfree: A program for rapid automated analysis of solution NMR spin-relaxation data. J. Biomol. NMR 26, 203–13. 32. Dosset, P., Hus, J. C., Blackledge, M., and Marion, D. (2000) Efficient analysis of macromolecular rotational diffusion from heteronuclear relaxation data. J. Biomol. NMR 16, 23–28. 33. Fushman, D., Cahill, S., and Cowburn, D. (1997) The main chain dynamics of the dynamin pleckstrin homology (PH) domain in solution: analysis of 15N relaxation with

9

Protein Dynamics by 15N Nuclear Magnetic Relaxation

monomer/dimer equilibration. J. Mol. Biol. 266, 173–194. 34. Butterwick, J. A., Loria, J. P., Astrof, N. S., Kroenke, C. D., Cole, R., Rance, M., and Palmer, A. G. (2004) Multiple time scale backbone dynamics of homologous thermophilic and mesophilic ribonuclease HI enzymes. J. Mol. Biol. 339, 855–871. 35. Sarkar, R., Moskau, D., Ferrage, F., Vasos, P. R., and Bodenhausen, G. (2008) Single or triple gradients? J. Magn. Reson. 193, 110–118. 36. Muhandiram, D. R., Yamazaki, T., Sykes, B. D., and Kay, L. E. (1995) Measurement of 2H T1ro Relaxation Times in Uniformly 13 C-Labeled and Fractionally 2H-Labeled Proteins in Solution. J. Am. Chem. Soc. 117, 11536–11544.

163

37. Bohlen, J. M., and Bodenhausen, G. (1993) Experimental Aspects of Chirp NMRSpectroscopy. J. Magn. Reson. A 102, 293–301. 38. Shaka, A. J., Barker, P. B., and Freeman, R. (1985) Computer-Optimized Decoupling Scheme for Wideband Applications and LowLevel Operation. J. Magn. Reson. 64, 547–552. 39. Emsley, L., and Bodenhausen, G. (1990) Gaussian Pulse Cascades-New Analytical Functions for Rectangular Selective Inversion and In-phase Excitation in NMR. Chem. Phys. Lett. 165, 469–476. 40. Shaka, A. J., Keeler, J., Frenkiel, T., and Freeman, R. (1983) An Improved Sequence for Broad Band Decoupling – WALTZ-16. J. Magn. Reson. 52, 335–38.

Chapter 10 Bacterial Production and Solution NMR Studies of a Viral Membrane Ion Channel Jolyon K. Claridge and Jason R. Schnell Abstract Advances in solution nuclear magnetic resonance (NMR) methodology that enable studies of very large proteins have also paved the way for studies of membrane proteins that behave like large proteins due to the added weight of surfactants. Solution NMR has been used to determine the high-resolution structures of several small, membrane proteins dissolved in detergent micelles and small bicelles. However, the usual difficulties with membrane proteins in producing, purifying, and stabilizing the proteins away from native membranes remain, requiring intensive screening efforts. Low levels of heterologous expression can be the most detrimental aspect to studying membrane proteins. This is exacerbated for NMR studies because of the costs of isotopically enriched media. Thus, solution NMR studies have tended to focus on relatively small, membrane proteins that can be expressed into inclusion bodies and refolded. Here, we describe the methods used to produce, purify, and refold the proton channel M2 into detergent micelles, and the procedures used to determine chemical shift assignments and the atomic level structure of the closed form of the homotetrameric channel. Key words: Membrane proteins, Ion channel, NMR, M2

1. Introduction Several difficulties arise in applying solution nuclear magnetic resonance (NMR) to the study of membrane proteins. Foremost among the difficulties are obtaining sufficient quantities of pure and natively folded membrane proteins at a reasonable cost, adverse effects of increased relaxation rates from complexes of protein and detergent micelles or detergent and lipid bicelles, limited lifetime of natively folded membrane proteins outside of lipid bilayers, and cost and availability of deuterated detergents. The requirement that the native structure be stable for days or months remains a general problem

Alexander Shekhtman and David S. Burz (eds.), Protein NMR Techniques, Methods in Molecular Biology, vol. 831, DOI 10.1007/978-1-61779-480-3_10, © Springer Science+Business Media, LLC 2012

165

166

J.K. Claridge and J.R. Schnell

for structural studies of membrane proteins and largely relies on trial and error to overcome. Access to relatively inexpensive deuterated detergents facilitates collection of information-rich methyl-based experiments, and in some cases it is necessary also to prevent rapid spin diffusion in backbone amide-based NOESY experiments. Increases in transverse relaxation rates with increasing protein size often mean that structure determinations to high resolution by solution NMR becomes very difficult for proteins with rotational correlation times of less than ~20 ns, unless extraordinary efforts are made. In general, this limit corresponds to protein sizes of ~35 kDa for water-soluble proteins at 30°C. However, studies of membrane proteins become challenging at much lower sizes because of the additional mass of the detergent micelles, which might double or triple the effective tumbling times. The introduction of TROSY-based pulse sequences (1, 2) and methyl-based approaches (3) that have been used extensively for much larger water-soluble proteins greatly benefit studies of small- and mediumsized membrane proteins (4, 5) as well. A common difficulty in studying membrane proteins by solution NMR (or, indeed, by any other method) arises from the inability to produce the proteins in suitable quantities. For NMR, the high unit cost of culture media enriched in the rare isotopes needed for multidimensional NMR studies prohibits the large-scale cultures (>10 L) common for crystallographic studies. Although isotopic labeling in eukaryotic systems (6, 7) or via cell-free methods (8) have been reported, the predominant method by which isotopically labeled proteins are produced remains by expression in bacteria. Using such methods, both the number and size of membrane proteins that have been solved by solution NMR have grown rapidly in the past few years (9, 10). Several of these studies expressed the proteins into bacterial membranes (e.g., refs. 3–5, 11); however, a larger trend has been to use solution NMR to study relatively small-membrane proteins that can be refolded from bacterial inclusion bodies. Membrane proteins containing a small number of transmembrane helices, which are more likely to fold in vitro, appear to be abundant in genomes (12, 13). Expression into inclusion bodies may happen spontaneously or by fusing the target protein to one that is efficiently “packaged” into inclusion bodies. One such fusion protein is “trpLE” (14), which has been used to express and purify several hydrophobic peptides: Vpu from HIV (15), transmembrane regions of GPCRs (16), caveolins (17), and several others (18–20) (see Note 1). TrpLE contains the leader sequence of the trp operon of Escherichia coli fused to a sequence of 97 residues found near the C-terminus of the anthranilate synthase gene (21, 22). Fusion to trpLE directs the protein to inclusion bodies, with the advantages of reduced

10

Bacterial Production and Solution NMR Studies of a Viral…

167

toxicity and protease resistance (23). In the trpLE construct, cysteines have been mutated to alanines to prevent disulfide bond formation, and methionines have been mutated to leucine to allow for the introduction of a single CNBr cleavage site at the fusion junction (14). In this chapter, we outline the approach used to produce, purify, and determine the structure of the M2 proton channel from influenza A in detergent micelles (24). M2 is one of the smallest ion channels known, being assembled from a homotetramer of a single-pass membrane protein. However, it is surprisingly sophisticated, being pH gated and highly proton selective. Past circulating strains of M2 are the target of a class of antiviral drugs, the adamantanes, and understanding drug resistance may help in developing drugs that are again effective. We describe how M2 is expressed at high yield into bacterial inclusion bodies as a trpLE fusion, chemically cleaved, purified, and refolded into detergent micelles to provide high-resolution NMR spectra of the native homotetramer (24) (Fig. 1).

Fig. 1. An overview of the homotetrameric structure of the high pH, closed M2 proton channel. Alpha helices are shown as cylinders. An idealized schematic of the DHPC micelle is shown as a single layer of detergent molecules with the hydrocarbon chains coating the transmembrane domain. The positioning of the micelle is based on the absence of cross peaks in amide backbone strips at the resonance frequency of water in an 15N-edited NOESY (24).

168

J.K. Claridge and J.R. Schnell

2. Materials 2.1. Expression of the TrpLE-M2 Fusion Protein

2.2. Fusion Protein Purification, Cleavage, and Preparation of Pure M2

●

Expression strain of E. coli: BL21(DE3) pLysS cells (Novagen).

●

LB agar plates: 10 mg/mL tryptone, 5 mg/mL yeast extract, 10 mg/mL NaCl, and 16 mg/mL agar. Adjust pH to 7.0, if necessary, with NaOH or HCl and autoclave. Add 100 mM kanamycin and 100 mM chloramphenicol just before solidification begins, and pour into plates.

●

LB medium: 10 mg/mL tryptone, 5 mg/mL yeast extract, and 10 mg/mL NaCl. Adjust pH to 7.0, if necessary, with NaOH or HCl and autoclave.

●

M9 medium: 42 mM Na2HPO4, 22 mM KH2PO4, 20 mM NH4Cl, 10 mM NaCl, 22 mM glucose, 2 mM MgSO4, 0.1 mM CaCl2, and 1× MEM vitamin solution (10 mL of a 100× stock purchased from Sigma–Aldrich), 100 mM kanamycin, and 100 mM chloramphenicol. The salts (Na2HPO4, KH2PO4, NH4Cl, and NaCl) are first dissolved into 990 mL of distilled and 0.22-mm filtered water and autoclaved, followed by addition of the remaining components. For production of 15N-labeled protein, uniformly 15N-labeled ammonium chloride is substituted. For 13C-labeled protein, 11 mM of uniformly 13C-labeled glucose is substituted. For partial deuteration, the M9 solutes are dissolved in 99.8% 2H2O and sterile filtered through 0.2-mm filter flasks, rather than autoclaved.

●

1 M DTT: Prepare stock solution in distilled, filtered water.

●

10% SDS: Prepare by adding 1 g of SDS to 10 mL of distilled, filtered water.

●

NuPAGE Novex 12% Bis–Tris Gel (Invitrogen).

●

4× NuPAGE LDS Sample Buffer (Invitrogen).

●

NuPAGE® MES SDS Running Buffer (Invitrogen).

●

1 M IPTG: Prepare stock solution in distilled, filtered water.

●

8 M Urea: Prepare stock solution in distilled, filtered water.

●

Lysis buffer: 50 mM Tris–HCl, pH 8.0, 200 mM NaCl.

●

Guanidine buffer: 20 mM Tris–HCl, pH 8.0, 200 mM NaCl, 6 M guanidine HCl, and 15 mM imidazole.

●

Elution buffer: 20 mM Tris–HCl, pH 7.0, 200 mM NaCl, 6 M guanidine HCl, and 400 mM imidazole.

●

Reverse phase C4 column for HPLC was purchased from GraceVydac (214TP C4, 300 Å silica, 5 mm beads, 2.1 × 150 mm).

●

Hexafluoroisopropanol: ³98% purity.

●

Formic acid: ³98% purity.

●

Syringe filter: 0.2 mm polytetrafluoroethylene.

2.3. Reconstitution of M2 into Detergent Micelles

2.4. NMR Chemical Shift Assignments and Restraint Measurements

2.5. Structure Calculation and Measurement of Tryptophan Gate Dynamics

10

Bacterial Production and Solution NMR Studies of a Viral…

169

●

HPLC buffer A: 5% isopropanol, 95% water, 0.1% trifluoroacetic acid.

●

HPLC buffer B: 57% isopropanol, 38% acetonitrile, 5% water, 0.1% trifluoroacetic acid.

●

Ni-NTA agarose beads.

●

Cyanogen bromide: Solid, ³98.5 % purity.

●

Dialysis tubing: 10 kDa MWCO, 22-mm internal diameter.

●

Dialysis cassette with 3.5 kDa MWCO.

●

Reconstitution buffer: 50 mM sodium phosphate, pH 7.5, 6 M guanidine HCl, 0.3 M dihexanoyl-sn-glycerol-3-phosphocholine (DHPC) detergent, 30 mM sodium glutamate. Adjust pH with NaOH.

●

NMR buffer: 50 mM sodium phosphate, pH 7.5, 30 mM sodium glutamate. Adjust pH with NaOH.

●

Deuterium oxide: 98% purity.

●

Rimantadine: 0.3 M stock prepared in 80 mM DHPC.

●

Mini dialysis cups for protein refolding: 3.5 kDa MWCO, 10–100 mL capacity (Thermo Scientific).

●

Centrifugal concentrator: 5 kDa MWCO.

●

All NMR experiments are conducted on a 14.1 T spectrometer equipped with a cryogenic probe.

●

Processing and preliminary analysis of data, peak fitting, and extraction of intensities for determining scalar bond couplings are performed using NMRPipe (25).

●

Resonance assignments and quantitation of NOE cross-peak intensities are performed using CARA (26).

●

Prediction of backbone dihedral angles from chemical shifts is performed using TALOS (27).

●

Cylindrically shaped polyacrylamide gel: A solution containing 4.5% acrylamide concentration with an acrylamide/bisacrylamide molar ratio of 40 is cast in a 6-mm cylindrical vessel.

●

A gel press kit (New Era Enterprises, Inc.) is used to push the cylindrical gel (6 mm in diameter) into an open-ended 4.2-mm inner diameter NMR tube.

●

Structure calculations were performed using the program XPLOR-NIH (28).

●

The program PALES (29) was used to evaluate the fit of residual dipolar couplings (RDCs) to structures.

●

Fitting of relaxation dispersion data was accomplished using the CPMGfit software (Dr. Art Palmer).

170

J.K. Claridge and J.R. Schnell

3. Methods The following protocols describe in detail our process for expressing, purifying, reconstituting the M2 proton channel into detergent micelles, assigning chemical shifts, and performing structural analyses. However, our lab has used the same protocol for several membrane proteins containing one or two transmembrane domains. A generalized flowchart of the procedure is provided in Fig. 2. The gene of interest was inserted between the HindIII (5¢) and BamHI (3¢) sites in the pMMHb plasmid (kindly provided by Stephen Blacklow, Brigham and Women’s Hospital, Boston), which contains a gene conferring kanamycin resistance. Cyanogen bromide is used to cleave the expressed construct on the C-terminal side of a unique methionine, resulting in release of the N-terminal (His)9-tagged trpLE, and the C-terminal M2 construct with no extra residues. Purified peptide of different isotopic composition is gently refolded from denaturing conditions into buffered detergent solution suitable for solution NMR experiments. Samples are typically stable for up to 3 weeks. Resonance assignments are obtained

Fig. 2. A generalized flowchart showing the production of tetrameric M2 samples from trpLE fusions expressed in Escherichia coli.

10

Bacterial Production and Solution NMR Studies of a Viral…

171

from a combination of triple-resonance NMR experiments, an 15 N-separated NOESY, and a 13C-separated NOESY collected on 15 N, 13C, 2H-labeled, 15N-labeled, and 15N, 13C-labeled peptide, respectively. Structure calculations are performed by an iterative process that incorporates local experimental restraints first, followed by longrange restraints. RDCs are incorporated last with a low-temperature refinement to avoid local minima that arise from the degeneracy of RDC magnitudes. 3.1. Expression of the TrpLE-M2 Fusion Protein

1. Transform the plasmid into E. coli BL21(DE3) pLysS cells, plate on LB agar containing kanamycin and chloramphenicol, and incubate overnight at 37°C. 2. Inoculate cultures of 200–500 mL Luria–Bertani (LB) media with individual colonies of freshly transformed bacteria and grow overnight at 37°C with moderate shaking (150 rpm). 3. Centrifuge the cultures at 2,000 × g for 25 min at 4°C and resuspend into 40 mL of chilled M9 medium. Add the resuspended cells to the large-scale M9 cultures (2–4 L) such that the OD600 is 0.2–0.3 relative to water. Grow each liter of culture in a 2.5-L baffled flask at 37°C with moderate shaking (150 rpm). 4. When the OD600 reaches 0.6–0.7, induce expression of the trpLE-M2 fusion by adding IPTG from a stock of 1 M IPTG to a final concentration of 1 mM. Grow overnight. The final OD600 is typically between 1.2 and 1.4 (see Note 2). 5. Analyze protein expression levels by using SDS-PAGE (12% Bis–Tris gel). Spin down cell quantities equivalent to 250 mL of OD 600 = 0.6 cell culture at 5,000 × g for 5 min at room temperature and redissolve the pellet into 40 mL of 8 M urea, 20 mL of 4× LDS sample buffer, 5 mL of 1 M DTT, and 5 mL of 10% SDS. Load 20 mL of sample per lane and run the gel using MES SDS running buffer.

3.2. Fusion Protein Purification, Cleavage, and Preparation of Pure M2

1. Harvest the cells by centrifugation at 5,000 × g for 30 min at 4°C. Resuspend the cell pellets in lysis buffer using a Dounce homogenizer, and sonicate on ice for a total of 3 min with a 20% duty cycle. Spin down the inclusion bodies and cell debris at 10,000 × g for 25 min at 4°C. Solubilize the pellets in 40 mL of lysis buffer and spin down at 10,000 × g for 25 min at 4°C to purify away additional water-soluble contaminants (see Note 3). 2. Dissolve the water-insoluble matter in 50 mL (per 1 L culture) of guanidine buffer using a Dounce homogenizer. Pellet undissolved matter, which includes nucleic acids, by centrifugation at 100,000 × g for 1.5 h at 4°C. Add the supernatant to 2 mL of Ni-NTA agarose beads (see Note 4) preequilibrated with guanidine buffer. After a 1-h incubation at 4°C with gentle rotation, pour the slurry into a gravity column and wash with 80 mL of guanidine buffer. Elute bound trpLE-M2 fusion

172

J.K. Claridge and J.R. Schnell

protein from the column by adding elution buffer in three stages of 5 mL each. 3. Dialyze the elution containing the trpLE-M2 fusion in 10 kDa molecular weight cutoff (MWCO) dialysis tubing against 4 L of H2O, with several exchanges over 4 h. The trpLE-M2 fusion precipitates as white flakes. Centrifuge in a swinging-bucket rotor at 1,500 × g for 30 min at 4°C. 4. Chemically cleave the M2 peptide from trpLE by dissolving the pellet into 5 mL of 70% formic acid containing 1 g of CNBr (see Note 5). Cover the reaction vessel with aluminum foil and allow the reaction to proceed for 2 h under a low-pressure stream of nitrogen gas. Load the sample via syringe into a dialysis cassette with a 3.5 kDa MWCO and dialyze against 4 L of water for 1 h, snap frozen in liquid nitrogen, and then lyophilize. 5. In preparation for reverse-phase HPLC, dissolve the lyophilized sample in 1.5 mL of hexafluoroisopropanol, which clarifies after 15 min at 40°C. Subsequently, add 0.5 mL of formic acid and 2 × 1 mL of water. Draw this solution into a 5-mL syringe, degas using a vacuum hose, and load onto a C4 reverse-phase column through a syringe filter. Collect fractions over a linear gradient from 0 to 100% buffer B (100–0% buffer A). Cleaved trpLE elutes first (~55% buffer B), followed by the uncut fusion (~65% buffer B), and the cleaved M2 peptide (~75% buffer B). Cleavage efficiencies for the trpLE-M2 fusion are routinely between 70 and 80%. Lyophilize the pooled fractions. For quantitation and aliquoting of peptide, dissolve the dried samples in 50% acetonitrile containing 0.1% TFA, quantitate by absorbance at 280 nm based on amino acid composition, and relyophilize (see Note 6). 3.3. Reconstitution of M2 into Detergent Micelles

1. Dissolve purified peptide (1.2 mg) into reconstitution buffer (see Note 7) at a concentration of 250 mM (final volume is 960 mL), split into three 3.5 kDa MWCO dialysis cups, and dialyze for 12 h against 2 L of NMR buffer with slow stirring and one buffer change at 10 h. 2. Concentrate the sample to ~0.7 mM monomer using a centrifugal concentrator with a 5 kDa MWCO. 3. Add rimantadine to 10 mM. Rimantadine binds at four equivalent sites near the gate on the lipid-facing side of the channel and stabilizes the closed conformation of the pore. Because of the poor water solubility of rimantadine, it is added from a 0.3 M stock that contains 80 mM DHPC (Subheading 2.3).

3.4. NMR Chemical Shift Assignments and Restraint Measurements

4. Add deuterium oxide for the magnetic field lock to a concentration of 5% in three steps. 1. To achieve nearly complete sequence-specific backbone chemical shift assignment of 1HN, 15N, 13C¢, 13Ca, and 13Cb, use

10

Bacterial Production and Solution NMR Studies of a Viral…

173

TROSY versions of the HNCA, HNCACB, and HNCO experiments (30, 31) on fully 15N-, 13C-, and 85% 2H-labeled protein (see Note 8). Process spectra in NMRPipe (25), and assign resonances and quantitate NOE cross-peak intensities in CARA (26). 2. Once backbone chemical shifts are known, use TALOS (27) to predict regions of secondary structure. For M2, this indicates that the transmembrane domain and the C-terminal juxtamembrane region are alpha-helical. This is confirmed by the characteristic local NOE patterns, and is used to aid assignment of most 1Ha and 1Hb intraresidue and sequential NOEs in a 3D 15 N-edited NOESY spectrum (110 ms; see Note 9 and Fig. 3). 3. Collect a methyl-based 3D 13C-edited NOESY (150 ms mixing time) on fully 15N-, 13C-labeled protein; this is particularly helpful in the methyl-rich transmembrane segment to confirm backbone proton assignments, extend the side-chain proton assignments, and identify intermolecular contacts (Fig. 4; see Note 8). Obtain stereospecific assignments of gamma methyls of valine and delta methyls of leucine from 10% 13C-labeled protein by recording a constant time 1H–13C HSQC with 28-ms carbon evolution, which allows discrimination between coupled and uncoupled methyls based on the sign of the cross peak (32). 4. Complete assignment of side-chain proton resonances is facilitated by determining a large number of c1 and c2 rotamers.

Fig. 3. Projection of the 1H,1H plane of 15N-separated NOESY spectra on an M2 sample containing (a) fully protonated or (b) fully deuterated DHPC detergent showing the loss of information due to spin diffusion when the hydrocarbon chain of DHPC is protonated (see Note 9). NOE mixing times were 90 and 110 ms, respectively.

174

J.K. Claridge and J.R. Schnell

Fig. 4. Final, lowest-energy structure of the high pH M2 channel showing the position of methyl groups throughout the transmembrane domain, including the helix–helix interfaces. Helices are shown as ribbons and methyl protons as filled circles. Structural elements in adjacent helices are shaded darker or lighter.

Determine the c1 of isoleucines, threonine, and valines from methyl-based measurements of the 3-bond scalar couplings 3 JNCg and 3JC¢C g (33, 34), and the c2 of leucine and isoleucine from 3JCaCd (35, 36). Determine the c1 of long-chain aliphatic (arginine, leucine, and lysine) and aromatic (histidine, phenylalanine, tryptophan, and tyrosine) side chains from 3JNCg and 3JC¢Cg values measured in an 1H–15N constant-time TROSY experiments on 15N-, 13C-, and 85% 2H-labeled protein (37, 38). 5. NOEs that cannot be explained by intramonomer distances based on the local secondary structure are identified as intermonomer NOEs. Carry out the assigning of intermonomer distance restraints and structure calculations iteratively until all NOE cross peaks in the NOESY spectra are self-consistent (see Subheading 3.5). First, identify protein-drug NOEs in the 15 N-edited and 13C-edited NOESYs described in step 3, and subsequently confirm them by acquiring an 15N-edited NOESY (500-ms mixing time) on a sample containing uniformly 15N- and 2 H-labeled protein, protonated rimantadine, and perdeuterated DHPC (see Note 10). 6. Weakly align the ion channel relative to the magnetic field by using a strained gel (39–41). Soak the protein and detergent solution into a cylindrically shaped polyacrylamide gel, initially 6 mm in diameter and 9 mm in length; squeeze this into the

10

Bacterial Production and Solution NMR Studies of a Viral…

175

4.2-mm inner diameter of an open-ended NMR tube. Obtain the RDCs by subtracting the scalar coupling (J), measured on an unaligned sample, from the measured coupling (J + RDC) of the aligned sample. In both cases, obtain couplings by interleaving a regular gradient-enhanced HSQC and a gradient-selected TROSY (42), both acquired with 80 ms of 15N evolution. 3.5. Structure Calculation and Measurement of Tryptophan Gate Dynamics

1. Begin structure calculations from a random coil structure generated in XPLOR-NIH that incorporates intramonomer NOEs, backbone dihedral angle restraints derived from chemical shifts, and side-chain c1 and c2 restraints. Enforce the intramonomer NOE restraints by using flat-well harmonic potentials, with the force constant fixed at 50 kcal-mol-1-Å-2. For fixed side-chain c1 and c2 angles, apply flat-well (±30°) harmonic potentials with a force constant of 30 kcal-mol-1-rad-2. During the simulation, ramp the van der Waals, improper angle, and bond angle force constants to 4.0 kcal-mol-1-Å-2, 1.0 kcal-mol-1-degree-2 and 1.0 kcal-mol-1-degree-2, respectively. Calculate a total of 20 monomer structures with a standard high-temperature simulated annealing protocol in which the bath temperature is cooled from 1,000 to 200 K. 2. To obtain an initial set of tetramer structures, replicate and uniquely translate the lowest-energy monomer structure four times. Perform another high-temperature simulated annealing run (1,000 to 200 K) using all previous restraints plus intermonomer NOEs, which are applied in a fourfold manner consistent with the C4 symmetry implied by a single set of resonances. Calculate a total of 100 tetramer structures. 3. Independently cross validate the 100 tetramer structures by 1 H–15N RDCs using singular value decomposition (SVD) as implemented in PALES. Assess the goodness of fit by the Pearson correlation coefficient (r) and quality factor (Q) (43). Select the 15 structures with the best agreement with RDCs (r ~ 0.91 and Q ~ 0.25). 4. Set the approximate initial values of the magnitude (Da) and rhombicity (Rh) of the alignment tensor to the average values of Da = 14.0 Hz and Rh = 0.20 that were obtained from the best SVD fits. Due to flexibility in the tetrameric assembly on the timescale of RDCs (up to microseconds (44)), the rhombicity is nonzero when the monomers are fit to a single alignment tensor. Thus, the monomers are restrained against independent alignment tensors during refinement. During this final refinement, cool the bath from 200 to 20 K. Fix the force constants for NOE and dihedral restraints at 100 kcal-mol-1-Å-2 and 40 kcal-mol-1-rad-2, respectively. In addition, ramp a weak database-derived “Rama” potential function from 0.02 to 0.20

176

J.K. Claridge and J.R. Schnell

(dimensionless force constant) for the general treatment of sidechain rotamers. The RDC restraint force constant was ramped from 0.010 to 0.125 kcal-mol-1-Hz-2, thereby supplementing, but not supplanting, the NOE restraints. Generate 10 RDCrefined structures for each of the 15 structures validated by RDCs, and add the structure with the lowest total energy to the final ensemble. Choose the structure with heavy atom conformation closest to the mean to represent the ensemble. 5. Measure the timescale of chemical shift exchange of the Trp41 side chain using an 15N relaxation dispersion CPMG experiment (45). Because the single tryptophan He1 of M2(18–60) is downfield from the backbone amides, measure relaxation dispersion to high precision using a 1D experiment with many scans (>1,000). 6. Fit the dependence of 15N relaxation due to chemical exchange on the frequency of refocusing (1/tcp) of chemical shift evolution to a two-site exchange model given by Rex ∝ 1 – (2tex/tcp) tanh (tcp/2tex), where Rex is the contribution to transverse relaxation due to chemical shift exchange and tex is the correlation time of the process that is generating the chemical shift exchange (46). Analyze the relaxation-compensated CPMG experiment (45) using the program CPMGfit.

4. Notes 1. Single-pass membrane proteins approaching 100 amino acids are robustly expressed into bacterial inclusion bodies with N-terminal trpLE fusions. We have observed that the expression yields of longer proteins or those containing additional TM helices can be increased by the addition of a second, C-terminal trpLE with an intervening methionine for cleavage. 2. Optimal temperature and IPTG conditions should be explored for each construct. In some cases, lower IPTG and lower temperature (100 mM IPTG and 18°C) or higher IPTG and higher temperature (1 mM IPTG and 42°C) result in higher expression levels. In some cases, better expression can be obtained using low volumes (200 mL) in 2.5-L baffled flasks and shaking at very high speeds (300 rpm) to increase aeration. 3. Beta-mercaptoethanol at 0.05% can be added to all cellprocessing buffers if cysteines are present in the target protein. 4. For maximum yields, it may be necessary to use fresh Ni-NTA resin for each preparation because of degradation that occurs in 6 M guanidine.

10

Bacterial Production and Solution NMR Studies of a Viral…

177

5. Use of CNBr for cleavage of the fusion protein requires a unique methionine at the fusion site. However, the side chain of the residue following the methionine can affect cleavage efficiencies. The hydroxyls of serine and threonine residues impair cleavage efficiency and should be avoided. In contrast, we have observed particularly high cleavage efficiencies when a glycine follows methionine. In addition, lower yields are observed for reactions with low concentrations of the fusion protein. Formylation of tryptophan and lysine side chains can occur when the reaction is allowed to proceed for longer than 3 h. CNBr is very toxic and corrosive, requiring a chemical hood and all safety precautions necessary to avoid direct contact. 6. A light, fluffy consistency of the lyophilized peptide prior to reconstitution correlated with best-quality samples for NMR. 7. The presence of glutamate is necessary to avoid nonspecific aggregation of M2 above ~200 mM (47). It is important to include glutamate in the 2H2O that is added to NMR samples to prevent localized aggregation. 8. Deuteration of the protein and the use of TROSY is critical for observing many of the cross peaks in the transmembrane domain because of the slow, rotational tumbling time of the tetrameric protein (~20 kDa) plus bound detergent. 9. The 3D 13C-edited NOESY spectrum was collected in the presence of perdeuterated detergent to prevent obfuscation of the aliphatic region. Collection of the 15N-edited NOESY spectrum also required deuterated detergent to prevent loss of magnetization due to rapid spin diffusion. Subsequent tests established that deuteration of only the acyl chains was sufficient to suppress rapid spin diffusion. The cost of perdeuterated DHPC (D35) is approximately tenfold more than that of DHPC deuterated only at the acyl chains (D22). 10. To ensure that essentially all nonexchangeable protein protons were replaced with deuterium, cells were grown in 99.9% 2H2O and perdeuterated glucose (Cambridge Isotope Laboratories).

Acknowledgments James J. Chou and the National Institutes of Health, USA (NIH) are acknowledged for supporting development of the approaches described. J.R.S. was supported by a Ruth Kirschstein Fellowship from the NIH. Matthew E. Call is acknowledged for many useful modifications and additions to the trpLE fusion expression and purification protocol.

178

J.K. Claridge and J.R. Schnell

References 1. LeMaster, D. M. (1994) Isotope labeling in solution protein assignment and structural analysis. Prog. Nucl. Magn. Reson. Spectrosc. 26, 371–419. 2. Pervushin, K., Riek, R., Wider, G., and Wuthrich, K. (1997) Attenuated T2 relaxation by mutual cancellation of dipole-dipole coupling and chemical shift anisotropy indicates an avenue to NMR structures of very large biological macromolecules in solution. Proc. Natl. Acad. Sci. U.S.A 94, 12366–12371. 3. Ruschak, A. M., and Kay, L. E. (2010) Methyl groups as probes of supra-molecular structure, dynamics and function. J. Biomol. NMR 46, 75–87. 4. Gautier, A., Mott, H. R., Bostock, M. J., Kirkpatrick, J. P., and Nietlispach, D. (2010) Structure determination of the seven-helix transmembrane receptor sensory rhodopsin II by solution NMR spectroscopy. Nat. Struct. Mol. Biol. 17, 768–774. 5. Imai, S., Osawa, M., Takeuchi, K., and Shimada, I. (2010) Structural basis underlying the dual gate properties of KcsA. Proc. Natl. Acad. Sci. U.S.A. 107, 6216–6221. 6. Bruggert, M., Rehm, T., Shanker, S., Georgescu, J., and Holak, T. A. (2003) A novel medium for expression of proteins selectively labeled with 15 N-amino acids in Spodoptera frugiperda (Sf9) insect cells. J. Biomol. NMR 25, 335–348. 7. Strauss, A., Bitsch, F., Cutting, B., Fendrich, G., Graff, P., Liebetanz, J., Zurini, M., and Jahnke, W. (2003) Amino-acid-type selective isotope labeling of proteins expressed in Baculovirusinfected insect cells useful for NMR studies. J. Biomol. NMR 26, 367–372. 8. Makino, S., Goren, M. A., Fox, B. G., and Markley, J. L. Cell-free protein synthesis technology in NMR high-throughput structure determination. Methods Mol. Biol. 607, 127–147. 9. Kim, H. J., Howell, S. C., Van Horn, W. D., Jeon, Y. H., and Sanders, C. R. (2009) Recent Advances in the Application of Solution NMR Spectroscopy to Multi-Span Integral Membrane Proteins. Prog. Nucl. Magn. Reson. Spectrosc. 55, 335–360. 10. Warchawski, D. (2010) Membrane proteins of known structure determined by NMR. http:// www.drorlist.com/nmr/MPNMR.html. 11. Chill, J. H., Louis, J. M., Miller, C., and Bax, A. (2006) NMR study of the tetrameric KcsA potassium channel in detergent micelles. Protein Sci. 15, 684–698. 12. Liu, Y., Engelman, D. M., and Gerstein, M. (2002) Genomic analysis of membrane protein families: abundance and conserved motifs. Genome Biol. 3, research0054.

13. Marsden, R. L., Lee, D., Maibaum, M., Yeats, C., and Orengo, C. A. (2006) Comprehensive genome analysis of 203 genomes provides structural genomics with new insights into protein family space. Nucleic Acids Res. 34, 1066–1080. 14. Staley, J. P., and Kim, P. S. (1994) Formation of a native-like subdomain in a partially folded intermediate of bovine pancreatic trypsin inhibitor. Protein Sci. 3, 1822–1832. 15. Ma, C., Marassi, F. M., Jones, D. H., Straus, S. K., Bour, S., Strebel, K., Schubert, U., OblattMontal, M., Montal, M., and Opella, S. J. (2002) Expression, purification, and activities of full-length and truncated versions of the integral membrane protein Vpu from HIV-1. Protein Sci. 11, 546–557. 16. Zheng, H., Zhao, J., Wang, S., Lin, C. M., Chen, T., Jones, D. H., Ma, C., Opella, S., and Xie, X. Q. (2005) Biosynthesis and purification of a hydrophobic peptide from transmembrane domains of G-protein-coupled CB2 receptor. J. Pept. Res. 65, 450–458. 17. Diefenderfer, C., Lee, J., Mlyanarski, S., Guo, Y., and Glover, K. J. (2009) Reliable expression and purification of highly insoluble transmembrane domains. Anal. Biochem. 384, 274–278. 18. Call, M. E., Schnell, J. R., Xu, C., Lutz, R. A., Chou, J. J., and Wucherpfennig, K. W. (2006) The structure of the zetazeta transmembrane dimer reveals features essential for its assembly with the T cell receptor. Cell 127, 355–368. 19. Chong, Y. H., Ball, J. M., Issel, C. J., Montelaro, R. C., and Rushlow, K. E. (1991) Analysis of equine humoral immune responses to the transmembrane envelope glycoprotein (gp45) of equine infectious anemia virus. J. Virol. 65, 1013–1018. 20. Smith, J. G., Mothes, W., Blacklow, S. C., and Cunningham, J. M. (2004) The mature avian leukosis virus subgroup A envelope glycoprotein is metastable, and refolding induced by the synergistic effects of receptor binding and low pH is coupled to infection. J. Virol. 78, 1403–1410. 21. Bertrand, K., Squires, C., and Yanofsky, C. (1976) Transcription termination in vivo in the leader region of the tryptophan operon of Escherichia coli. J. Mol. Biol. 103, 319–337. 22. Miozzari, G. F., and Yanofsky, C. (1978) Translation of the leader region of the Escherichia coli tryptophan operon. J. Bacteriol. 133, 1457–1466. 23. Kleid, D. G., Yansura, D., Small, B., Dowbenko, D., Moore, D. M., Grubman, M. J., McKercher, P. D., Morgan, D. O., Robertson, B. H., and Bachrach, H. L. (1981) Cloned viral protein vaccine for foot-and-mouth disease: responses in cattle and swine. Science 214, 1125–1129.

10

Bacterial Production and Solution NMR Studies of a Viral…

24. Schnell, J. R., and Chou, J. J. (2008) Structure and mechanism of the M2 proton channel of influenza A virus. Nature 451, 591–595. 25. Delaglio, F., Grzesiek, S., Vuister, G. W., Zhu, G., Pfeifer, J., and Bax, A. (1995) NMRPipe: a multidimensional spectral processing system based on UNIX pipes. J. Biomol. NMR 6, 277–293. 26. Keller, R. (2004) The Computer Aided Resonance Assignment Tutorial, First Edition ed., CantinaVerlag, Goldau, Switzerland. http://cara.nmrsoftware.org/downloads/3-85600-112-3.pdf. 27. Cornilescu, G., Delaglio, F., and Bax, A. (1999) Protein backbone angle restraints from searching a database for chemical shift and sequence homology. J. Biomol. NMR 13, 289–302. 28. Schwieters, C. D., Kuszewski, J., Tjandra, N., and Clore, G. M. (2002) The Xplor-NIH NMR molecular structure determination package. J. Magn. Reson. 160, 66–74. 29. Zweckstetter, M., Bax, A. (2000) Prediction of sterically induced alignment in a dilute liquid crystalline phase: aid to protein structure determination by NMR. J. Am. Chem. Soc. 122, 3791–3792. 30. Salzmann, M., Wider, G., Pervushin, K., and Wuthrich, K. (1999) Improved sensitivity and coherence selection for [N-15,H-1]-TROSY elements in triple resonance experiments. J. Biomol. NMR 15, 181–184. 31. Kay, L. E., Ikura, M., Tschudin, R., and Bax, A. (1990) Three-dimensional triple resonance NMR spectroscopy of isotopically enriched proteins. J. Magn. Reson. 89, 496–514. 32. Neri, D., Szyperski, T., Otting, G., Senn, H., and Wuthrich, K. (1989) Stereospecific nuclear magnetic resonance assignments of the methyl groups of valine and leucine in the DNAbinding domain of the 434 repressor by biosynthetically directed fractional 13C labeling. Biochemistry 28, 7510–7516. 33. Grzesiek, S., Vuister, G. W., and Bax, A. (1993) A simple and sensitive experiment for measurement of JCC couplings between backbone carbonyl and methyl carbons in isotopically enriched proteins. J. Biomol. NMR 3, 487–493. 34. Vuister, G. W., Wang, A. C., and Bax, A. (1993) Measurement of three-bond nitrogen-carbon J couplings in proteins uniformly enriched in 15N and 13C. J. Am. Chem. Soc. 115, 5334–5335. 35. Bax, A., Vuister, G. W., Grzesiek, S., Delaglio, F., Wang, A. C., Tschudin, R., and Zhu, G. (1994) Measurement of homo- and heteronuclear J couplings from quantitative J correlation. Methods Enzymol. 239, 79–105. 36. MacKenzie, K. R., Prestegard, J. H., and Engelman, D. M. (1996) Leucine side-chain rotamers in a glycophorin A transmembrane

179

peptide as revealed by three-bond carbon-carbon couplings and 13C chemical shifts. J. Biomol. NMR 7, 256–260. 37. Hu, J.-S., Grzesiek, S., and Bax, A. (1997) Chi1 angle information from a simple two-dimensional NMR experiment which identifies trans 3JNCg couplings in isotopically enriched proteins. J. Biomol. NMR 9, 323–328. 38. Hu, J.-S., Grzesiek, S., and Bax, A. (1997) Two-dimensional NMR methods for determining c1 angles of aromatic residues in proteins from three-bond JC¢Cg and JNCgcouplings. J. Am. Chem. Soc. 119, 1803–1804. 39. Tycko, R., Blanco, F. J., and Ishii, Y. (2000) Alignment of biopolymers in strained gels: A new way to create detectable dipole-dipole couplings in high-resolution biomolecular NMR. J. Am. Chem. Soc. 122, 9340–9341. 40. Sass, H. J., Musco, G., Stahl, S. J., Wingfield, P. T., and Grzesiek, S. (2000) Solution NMR of proteins within polyacrylamide gels: diffusional properties and residual alignment by mechanical stress or embedding of oriented purple membranes. J. Biomol. NMR 18, 303–309. 41. Chou, J. J., Gaemers, S., Howder, B., Louis, J. M., Bax, A. (2001) A simple apparatus for generating stretched polyacrylamide gels, yielding uniform alignment of proteins and detergent micelles. J. Biomol. NMR 21, 377–382. 42. Weigelt, J. (1998) Single Scan, Sensitivity- and Gradient-Enhanced TROSY for Multidimensional NMR Experiments. J. Am. Chem. Soc. 120, 10778–10779. 43. Cornilescu, G., Marquardt, J. L., Ottiger, M., and Bax, A. (1998) Validation of protein structure from anisotropic carbonyl chemical shifts in a dilute liquid crystalline phase. J. Am. Chem. Soc. 120, 6836–6837. 44. Lakomek, N. A., Lange, O. F., Walter, K. F., Fares, C., Egger, D., Lunkenheimer, P., Meiler, J., Grubmuller, H., Becker, S., de Groot, B. L., and Griesinger, C. (2008) Residual dipolar couplings as a tool to study molecular recognition of ubiquitin. Biochem. Soc. Trans. 36, 1433–1437. 45. Loria, J. P., Rance, M., and Palmer, A. G. (1999) A relaxation-compensated Carr-PurcellMeiboom-Gill sequence for characterizing chemical exchange by NMR spectroscopy. J. Am. Chem. Soc. 121, 2331–2332. 46. Allerhand, A., and Thiele, E. (1966) Analysis of Carr-Purcell Spin-Echo NMR Experiments on Multiple-Spin Systems. II. The Effect of Chemical Exchange. J. Chem. Phys. 45, 902–916. 47. Golovanov, A. P., Hautbergue, G. M., Wilson, S. A., and Lian, L. Y. (2004) A simple method for improving protein solubility and long-term stability. J. Am. Chem. Soc. 126, 8933–8939.

Chapter 11 Preparation of the Modular Multi-Domain Protein RPA for Study by NMR Spectroscopy Chris A. Brosey, Marie-Eve Chagot, and Walter J. Chazin Abstract The integrity and propagation of the genome depend upon the fidelity of DNA processing events, such as replication, damage recognition, and repair. Requisite to the numerous biochemical tasks required for DNA processing is the generation and manipulation of single-stranded DNA (ssDNA). As the primary eukaryotic ssDNA-binding protein, Replication Protein A (RPA) protects ssDNA templates from stray nuclease cleavage and untimely reannealment. More importantly, RPA also serves as a platform for organizing access to ssDNA for readout of the genetic code, recognition of aberrations in DNA, and processing by enzymes. We have proposed that RPA’s ability to adapt to such a broad spectrum of multiprotein machinery arises in part from its modular organization and interdomain flexibility. While requisite for function, RPA’s modular flexibility has presented many challenges to providing a detailed characterization of the dynamic architecture of the full-length protein. To enable the study of RPA’s interdomain dynamics and responses to ssDNA binding by biophysical methods including NMR spectroscopy, we have successfully produced recombinant full-length RPA in milligram quantities at natural abundance and enriched with NMR-active isotopes. Key words: Replication Protein A, DNA processing, Protein modularity, Isotopic labeling, Recombinant expression, Protein purification, NMR spectroscopy

1. Introduction As the primary eukaryotic single-stranded DNA (ssDNA)-binding protein, Replication Protein A (RPA) prevents reannealment of unwound DNA strands, controls access to DNA templates, and serves as a scaffold for the assembly and disassembly of DNA processing machinery (1, 2). A heterotrimer, RPA’s three subunits (RPA70, RPA32, and RPA14) contain seven structured domains interconnected by flexible linkers. Three of these domains form

Alexander Shekhtman and David S. Burz (eds.), Protein NMR Techniques, Methods in Molecular Biology, vol. 831, DOI 10.1007/978-1-61779-480-3_11, © Springer Science+Business Media, LLC 2012

181

182

C.A. Brosey et al.

the trimeric core of the protein (70C, 32D, 14), from which emanate the flexibly linked N-terminal domains of RPA70 (70N, 70A, 70B), as well as the disordered N-terminal and structured C-terminal domains of RPA32 (32N and 32C, respectively). Binding of ssDNA is facilitated by domains 70A, 70B, 70C, and 32D, which together occupy an occluded site size of 30 nucleotides (1). Interactions with other DNA-processing proteins are primarily mediated by domains 70N and 32C, and the principal DNA-binding domains 70A and 70B (1, 2). As a universal participant in DNA processing, RPA must interact with a wide array of structurally unique multiprotein complexes. The flexible, modular organization of the protein is thought to be critical for enabling such structural adaptability (3). Although high-resolution X-ray or NMR structures of all individual RPA domains are available (4–8), the dynamic interdomain organization of full-length RPA and the accompanying structural alterations imposed by DNA processing have not been extensively characterized. The full-length protein’s intrinsic flexibility poses several challenges to study by X-ray diffraction; and at 116 kDa, RPA falls outside the size limit of conventional NMR methods (30–40 kDa). Application of advanced NMR approaches, however, namely, deuterium labeling and TROSY- or CRINEPT-based techniques, has allowed this size limitation to be extended to proteins in excess of 100 kDa (9–12). This, combined with the discrete distribution of molecular mass among RPA domains (50 kDa for the trimer core and 10–14 kDa for the remaining domains), makes feasible characterization of the full-length protein by NMR (13). Here, we describe the production of full-length RPA by recombinant expression in Escherichia coli and subsequent purification of the protein by a series of FPLC steps. The protocols provided include those required for preparation of 2H-, 15N-enriched RPA for study by NMR spectroscopy.

2. Materials 2.1. Cell Transformation

1. RPA pET15b plasmid (see Note 1). 2. BL21(DE3) pLyS competent cells: 100-μL aliquots stored at −80°C. 3. LB medium plates: 10 g/L tryptone, 10 g/L NaCl, 5 g/L yeast extract, 15 g/L agar dissolved in Milli-Q water (filtered to a resistance of 18.3 MΩ-cm) and autoclaved at 121°C for 15 min. Add antibiotic stocks (ampicillin and chloramphenicol) at 1:1,000 dilution when the medium has cooled to 50–60°C (14) (see Note 2).

11

Preparation of the Modular Multi-Domain Protein RPA…

183

4. 1,000× Ampicillin stock: 100 mg/mL in Milli-Q water, sterilize by filtration at 0.2 μm (see Note 3). 5. 1,000× Chloramphenicol stock: 34 mg/mL in ethanol, sterilize by filtration at 0.2 μm (see Note 3). 6. SOC recovery medium: 20 g/L tryptone, 0.5 g/L NaCl, 5 g/L yeast extract, 2.5 mM KCl, 5 mM MgCl2, 5 mM MgSO4, 20 mM glucose dissolved in Milli-Q water and autoclaved at 121°C for 15 min (14). 2.2. Cell Expression Testing

1. RPA pET15b BL21(DE3) pLysS LB plate (Subheading 2.1). 2. Sterile 10-mL test culture tubes. 3. LB medium: 10 g/L tryptone, 10 g/L NaCl, 5 g/L yeast extract dissolved in Milli-Q water and autoclaved at 121°C for 15 min. 4. 1,000× Antibiotic stocks (Subheading 2.1). 5. 1 M IPTG: Sterilize by filtration at 0.2 μm and store at −20°C. 6. 2× SDS loading buffer: 100 mM Tris–HCl, pH 6.8, 4% (w/v) SDS (electrophoresis grade), 0.2% (w/v) bromophenol blue, 20% (w/v) glycerol, 200 mM β-mercaptoethanol (βME, added fresh) (14). 7. 8 M urea. 8. Precast 4–12% Bis-Tris SDS-PAGE gel (Invitrogen). 9. 1× MES SDS running buffer (Invitrogen). 10. 1× Prestained molecular weight standards. 11. SimplyBlue SafeStain.

2.3. Preparation of Culture Media

1. LB medium (Subheading 2.2).

2.3.1. LB Medium

3. 500-mL Erlenmeyer flask with baffles.

2. 1,000× Antibiotic stocks (Subheading 2.1). 4. Six 2.8-L Fernbach flasks with baffles.

2.3.2. Minimal Medium

1. Milli-Q water (900 mL/L medium). 2. 10× M9 salts: 5 g/L NaCl, 30 g/L KH2PO4, 60 g/L Na2HPO4 dissolved in Milli-Q water, adjusted to pH 7.4 with 10 M NaOH, and autoclaved at 121°C for 15 min. 3. 1 M MgSO4: Sterilize filter at 0.2 μm and store at room temperature. 4. 1 M CaCl2: Sterilize filter at 0.2 μm and store at room temperature.

184

C.A. Brosey et al.

5. 20% (w/v) Glucose: Sterilize filter at 0.2 μm and store at room temperature. 6. 1 M Thiamine hydrochloride: Sterilize filter at 0.2 μm and store at room temperature. 7. 1,000× Antibiotic stocks (Subheading 2.1). 8.

15

NH4Cl.

9. 500-mL Erlenmeyer flask with baffles. 10. Six 2.8-L Fernbach flasks with baffles. 2.3.3. Deuterated Minimal Medium

1. 99% D2O. 2. Dry components: 0.5 g/L NaCl, 3 g/L KH2PO4, 6 g/L Na2HPO4, 0.24 g/L-MgSO4, 11.1 mg/L-CaCl2, 2 g/L-glucose, 0.337 g/L thiamine hydrochloride, 0.5 g/L 15NH4Cl, and 0.1 g/L ampicillin. 3. Six sterile vacuum filtration systems with 1-L storage containers. 4. Six sterile 2.8-L Fernbach flasks with baffles.

2.4. Starter Cultures

1. RPA pET15B BL21(DE3) pLysS LB plate (Subheading 2.1).

2.4.1. LB Medium

2. 250 mL LB starter culture (Subheading 2.3.1).

2.4.2. Minimal Medium and Deuterated Minimal Medium

1. RPA pET15B BL21(DE3) pLysS LB plate (Subheading 2.1). 2. Sterile 10-mL test culture tubes. 3. LB medium (Subheading 2.2). 4. 1,000× Antibiotic stocks (Subheading 2.1). 5. 250 mL Minimal medium starter culture (Subheading 2.3.2).

2.5. Large-Scale Cell Culture and Overexpression

1. Starter culture (Subheading 2.4). 2. 6 L Sterile media in Fernbach flasks (Subheading 2.3). 3. 1 M IPTG (Subheading 2.2). 4. Bleach or 1% Terg-a-Zyme solution for decontamination of spent media.

2.6. RPA Purification 2.6.1. Cell Lysis

1. Lysis buffer: Dissolve two complete EDTA-free protease inhibitor cocktail tablets (Roche) in 80 mL of Ni-NTA buffer A (Subheading 2.6.2) in a 150-mL glass beaker on ice immediately prior to use. 2. 100-mL Glass homogenizer. 3. Sonic dismembrator. 4. 25-mm diameter, 0.45-μm syringe filter.

2.6.2. Ni-NTA Chromatography

1. Refrigerated Äkta FPLC purification system and accessories. 2. Ni-NTA buffer A: 20 mM HEPES, pH 7.5, 500 mM NaCl, 5 mM βME, 10 μM ZnCl2, 10 mM imidazole; adjusted to

11

Preparation of the Modular Multi-Domain Protein RPA…

185

target pH with concentrated HCl, filtered at 0.45 μm under vacuum, and stored at 4°C (see Notes 4–7). 3. Ni-NTA buffer B: 20 mM HEPES, pH 7.5, 500 mM NaCl, 5 mM βME, 10 μM ZnCl2, 300 mM imidazole; adjusted to target pH with concentrated HCl, filtered at 0.45 μm under vacuum, and stored at 4°C (see Notes 4–7). 4. 25 mL Ni-NTA pre-packed FPLC column (Sigma–Aldrich). 2.6.3. Desalting Exchange and Heparin Chromatography

1. Centrifugal concentrators (15 mL, 30 kDa MWCO). 2. Refrigerated Äkta FPLC purification system and accessories. 3. Heparin buffer A: 20 mM HEPES, pH 7.5, 50 mM NaCl, 5 mM βME, 10 μM ZnCl2, 10% glycerol; adjusted to target pH with concentrated HCl, filtered at 0.45 μm under vacuum, and stored at 4°C (see Notes 6 and 7). 4. Heparin buffer B: 20 mM HEPES, pH 7.5, 1 M NaCl, 5 mM βME, 10 μM ZnCl2, 10% glycerol; adjusted to target pH with concentrated HCl, filtered at 0.45 μm under vacuum, and stored at 4°C (see Notes 6 and 7). 5. HiPrep 26/10 Desalting column (GE Healthcare). 6. HiTrap 5-mL Heparin HP column (GE Healthcare).

2.6.4. Superdex 200 Gel Filtration Chromatography

1. Centrifugal concentrators (15 mL, 30 kDa MWCO). 2. 0.22-μm centrifugal spin filters. 3. Refrigerated Äkta FPLC purification system and accessories. 4. Gel filtration buffer: 20 mM HEPES, pH 7.5, 100 mM NaCl, 5 mM βME, 10 μM ZnCl2, 200 mM arginine; adjusted to target pH with concentrated HCl, filtered at 0.45 μm under vacuum, and stored at 4°C (see Notes 6 and 7). 5. Superdex 200 HR 10/30 column (GE Healthcare).

2.7. Preparation of Samples for NMR

1. Centrifugal concentrators (15 mL, 30 kDa MWCO). 2. 5- and 4-mm NMR tubes. 3. 99% D2O.

3. Methods This section describes a protocol for the production of full-length RPA in E. coli and its subsequent purification, including preparation of 2H-,15N-enriched protein for study by NMR. As a rule, robust expression of full-length RPA in E. coli is challenging as RPA is a relatively large protein (>100 kDa) and its ssDNA binding properties are toxic to bacterial cells. Average yields of RPA overexpressed from the pET15b vector range from 17 mg for 6 L of

186

C.A. Brosey et al.

rich LB culture to 4–8 mg of 15N-enriched RPA for 6 L of minimal medium culture (1–2 NMR samples at approximately 130 μM concentration and 260-μL volume). Working with RPA cultures grown in deuterated minimal medium requires patience and careful monitoring, as a deuterated environment is particularly stressful to the bacterial metabolism. Consequently, deuterated cultures take much longer to reach their target induction densities and usually result in a diminished yield of the recombinant protein. Due to the high cost of D2O and time investment required for growth and expression, we found it beneficial to pilot a small-scale test culture (100 mL) to develop an expected timeline for the growth and to ensure that all reagents were functioning as expected. Subsequent purification of this culture allowed us to determine that the overall yield of RPA had not suffered significantly from production in a deuterated environment. In the expression protocol below, we describe the full 6 L production run; however, when embarking upon deuterium labeling for the first time, we highly recommend starting with the smaller pilot culture. As RPA is a trimeric, DNA-binding protein, the purification protocol below is designed to ensure samples free of contaminating ssDNA, as well as uniform stoichiometry among all three RPA subunits. Expression of RPA from the pET15b vector results in an excess of the RPA70 subunit, which can be successfully separated from the intact heterotrimer with heparin and gel filtration chromatography. The heparin purification step also selects for RPA free from ssDNA contamination. 3.1. Cell Transformation

Ensuring robust antibiotic selection of the RPA vector on solid medium is vital to enabling the success of subsequent liquid cultures (see Note 2). 1. Thaw 100 μL of BL21(DE3) pLyS competent cells on ice, gently combine with 100 ng of RPA pET15b vector, and incubate for 30 min on ice. 2. Heat shock the cells at 42°C for 45 s and incubate on ice for 2 min. Add 900 μL of sterile SOC recovery medium. 3. Incubate the cells for 1 h at 37°C and 200–230 rpm, then centrifuge the cells at 16,100 × g for 1 min at room temperature (see Note 8). Remove 900 μL of the clarified SOC medium and gently resuspend the cells in the remaining medium prior to plating on an LB medium plate. 4. Incubate the plates overnight at 37°C.

3.2. Cell Expression Testing

Expression testing allows confirmation of the expression capability of the transformed bacterial colonies prior to scaling up protein production. The testing also allows for selection of colonies with the most robust expression.

11

Preparation of the Modular Multi-Domain Protein RPA…

187

1. Prepare five LB test cultures as follows: Transfer 5 mL of LB medium to a 10-mL sterile culture tube and add 1:1,000 dilutions of ampicillin and chlormaphenicol antibiotic stocks. Inoculate each culture with a colony selected from the center of a freshly transformed RPA pET15b LB plate (see Note 9) and incubate the cultures at 37°C, 200–230 rpm, until they reach an A600 of 0.5–0.6 (approximately 3–4 h). 2. Transfer 250 μL of each LB test culture into an Eppendorf tube as a preinduction sample. Centrifuge the sample at 16,100 × g for 1 min, decant the supernatant, add 7 μL each of 2× SDSPAGE loading buffer and 8 M urea, and vortex to mix. 3. Add IPTG to a final concentration of 1 mM to the remainder of the test culture to induce expression and continue to incubate with shaking at room temperature for 3 h. Collect a final 250 μL postinduction sample and process as in step 2. 4. Boil pre- and postinduction SDS-PAGE samples for 5–10 min to denature the lysates and load 2–4 μL from each sample into a precast 4–12% Bis–Tris SDS-PAGE gel preloaded into an electrophoresis cell filled with 1× MES SDS running buffer. Reserve one lane for 5 μL of 1× prestained molecular weight standards. Run the gel at 200 V. 5. Remove the gel from the electrophoresis cell and place in a loosely capped container filled with Milli-Q water (see Note 10). Fix the gel by heating on a high setting in a microwave oven for 1 min, followed by 1 min of cooling. Exchange the water and repeat. Remove the final rinse and stain with SimplyBlue SafeStain for 20 min. Remove the stain and refill the container with deionized water to destain the gel. 6. The relative proportion of RPA32 and RPA14 subunits is too low to observe on the gel. RPA70 should be just distinguishable at the appropriate molecular weight in lanes containing postinduction samples. Select colonies exhibiting the most abundant RPA production for subsequent largescale expression. 3.3. Preparation of Culture Media 3.3.1. LB Medium

This medium serves for the production of unlabeled RPA. Preparation should include a 250-mL starter culture to accommodate a 6-L large-scale culture. 1. Dissolve LB components (Subheading 2.2) in Milli-Q water in a 500-mL baffled Erlenmeyer flask (starter culture) or 2.8-L baffled Fernbach flasks (large-scale culture), autoclave at 121°C for 15 min, and cool to 50–60°C. 2. Add ampicillin and chloramphenicol at 1:1,000 dilution immediately prior to inoculation.

188

C.A. Brosey et al.

3.3.2. Minimal Medium

This medium serves for the production of 15N-enriched RPA. Preparation should include a 250 mL starter culture to accommodate a 6 L large-scale culture. A 250 mL minimal medium culture also serves as an adaptation culture for production of deuterated protein. 1. Dilute 10× M9 salts in Milli-Q water to 1× in a 500-mL baffled Erlenmeyer flask (starter culture) or 2.8-L baffled Fernbach flasks (large-scale culture), autoclave at 121°C for 15 min, and cool to 50–60°C. 2. Add the following components immediately prior to inoculation: 0.5 g/L of 15NH4Cl (see Note 11), 2 mL/L of 1 M MgSO4, 100 μL/L of 1 M CaCl2, 10 mL/L of 20% glucose, 1 mL/L of 1 M thiamine hydrochloride, and antibiotic stocks at 1:1,000 dilution.

3.3.3. Deuterated Minimal Medium

This medium serves for six 1 L large-scale production cultures and is prepared immediately prior to inoculation after the success of the 250 mL minimal medium adaptation culture has been ascertained. 1. Dry autoclave six 2.8-L Fernbach flasks with baffles and allow to dry thoroughly overnight. 2. Dissolve all dry components in 6 L of 99% D2O (see Notes 12 and 13) and immediately sterilize the medium in 1-L batches by using sterile vacuum filtration systems (i.e., 1 L/unit). This apparatus filters the medium directly into a sterile 1-L bottle. Chloramphenicol is not included in the medium at this stage to ease the metabolic burden on the cells. 3. Carefully transfer each 1 L of sterile deuterated minimal medium to a dry, sterile Fernbach flask (see Note 14).

3.4. Starter Cultures 3.4.1. LB Medium

1. Inoculate a 250 mL LB starter culture (Subheading 3.3.1) directly with an RPA pET15b colony selected from the test expression. 2. Grow the culture overnight at 37°C, 200–230 rpm. The culture should be cloudy in the morning.

3.4.2. Minimal Medium and Deuterated Minimal Medium

1. Inoculate a 4 mL LB starter culture (4 mL LB + 1:1,000 ampicillin/chloramphenicol in a 10-mL culture tube) with an RPA pET15b colony selected from the test expression. Grow for 3–4 h at 37°C, 200–230 rpm, or until cloudy. 2. Inoculate the 250 mL minimal medium starter culture (Subheading 3.3.2) with the 4 mL LB starter culture and shake overnight at 37°C. The culture should be cloudy in the morning.

11

3.5. Large-Scale Cell Culture and Overexpression

Preparation of the Modular Multi-Domain Protein RPA…

189

1. Prepare six 1 L cultures of rich LB medium, minimal medium, or deuterated minimal medium as described above (Subheading 3.3) and inoculate each with 30 mL (40 mL for deuterated minimal medium) of the corresponding overnight starter culture. 2. Grow the cultures at 37°C, 200–230 rpm, until an A600 of 0.6–0.7 is reached (see Note 15). 3. Allow the cultures to equilibrate for half an hour with agitation at 18°C (or room temperature for deuterated minimal medium) prior to induction. Collect a preinduction SDS-PAGE sample as described above (Subheading 3.2) and induce the cells with 1 mM IPTG. Allow cells to express overnight (approximately 16–18 h). 4. The A600 at the end of the expression period should be 1.8–2.0 for LB medium cultures and 0.9–1.0 for standard and deuterated minimal media cultures. Collect postinduction SDS-PAGE samples from the cultures as described above (Subheading 3.2). Harvest the cultures by centrifuging at 10,000 × g for 20 min at 4°C. 5. Decant the supernatant and reserve the spent deuterated media for recycling (15). Spent LB or minimal media may be decontaminated by the addition of bleach or a 1% Terg-a-zyme solution for 30 min, and then discarded. If purification does not follow immediately, transfer the pellets to sterile 50-mL conical tubes and freeze at −80°C. Run pre- and postinduction SDSPAGE samples as in Subheading 3.2 to confirm the presence of RPA expression.

3.6. RPA Purification

3.6.1. Cell Lysis

Purification of RPA involves three primary steps: Ni-NTA affinity, heparin, and size-exclusion chromatography. For best results, the protocol should be completed over the course of 2 days, where Ni-NTA and heparin chromatography steps are accomplished the first day and the final gel filtration step is carried out on the second day. If necessary, the Ni-NTA and heparin steps may be divided into two separate days and protein fractions from each purification kept at 4°C overnight. Ideally, though, the time from cell lysis to the final gel filtration exchange should be kept to a minimum. 1. If cells have been frozen at −80°C, thaw the pellets by submerging the 50-mL conical tubes in cool water. Meanwhile, prepare and chill the lysis buffer and pre-chill the 100-mL glass homogenizer on ice (see Note 16). 2. Transfer all 6 L of RPA cell pellets into the 100-mL homogenizer, rinse the 50-mL conical tubes with ice-cold lysis buffer, and add the rinse and any remaining lysis buffer to the homogenizer.

190

C.A. Brosey et al.

3. Homogenize the lysate until smooth (10–15 strokes). 4. Return the lysate to the 150-mL glass beaker and pack this into a 2-L plastic beaker filled with an ice-water bath (see Note 17). Ensure that there is sufficient ice to securely brace the beaker and prevent floating. 5. Sonicate the lysate with a macrotip set at 60% power for 5.0 min of total process time (pulsing 5.0 s on and 5.0 s off). Pause the sonicator half-way through this cycle to replenish the ice-water bath and to check the temperature of the lysate (see Note 16). The lysate should become translucent and less viscous as the sonicator disrupts the cellular material. If the lysate viscosity remains unchanged after the cycle is complete, repeat the cycle once more, monitoring the ice-water bath and lysate temperature. 6. Clarify the lysate by centrifuging at 48,000 × g for 20 min at 4°C. Ensure that both centrifuge and rotor are pre-chilled to at least 4°C. 7. Decant and filter the clarified supernatant through a 0.45-μm membrane. Store on ice for immediate loading onto the FPLC Ni-NTA column. 3.6.2. Ni-NTA Chromatography

Steps 2–4 are implemented as a pre-programmed Äkta FPLC method. 1. Equilibrate the prepacked 25 mL Ni-NTA column with three column volumes (3 CVs) each of filtered Milli-Q water and Ni-NTA buffer A (see Note 18). 2. Load the filtered lysate onto the equilibrated Ni-NTA column at 1.0–1.5 mL/min. 3. Wash unbound lysate from the column with 4 CVs of Ni-NTA buffer A at 2.5 mL/min. 4. Elute RPA with a 4 CV gradient (0–100% Ni-NTA buffer B, 10–300 mM imidazole), collecting 6-mL fractions at 2.5 mL/min. 5. Assess the presence of RPA from the A280 chromatogram trace and SDS-PAGE of relevant fractions (sampling 5 μL of each fraction). Pool fractions containing all three RPA subunits for further processing (typically, a 60-mL pool).

3.6.3. Desalting Exchange and Heparin Chromatography

The charged DNA-binding clefts of RPA render it sensitive to the absence of ambient salt. Effective binding of RPA to the heparin matrix, however, requires a low salt content in the loading buffer. Direct dialysis into the loading buffer (heparin buffer A) usually provokes extensive precipitation of RPA. Buffer exchange by desalting, however, allows for the rapid and successful transfer of RPA into the loading buffer with minimal aggregation. Once the series of FPLC desalting runs are complete, it is imperative to load the exchanged protein directly onto the heparin column to restore a stabilizing ionic environment. As before, loading, washing, and

11

Preparation of the Modular Multi-Domain Protein RPA…

191

elution are implemented automatically using pre-programmed Äkta FPLC methods. 1. Pre-rinse two centrifugal concentrators (15 mL, 30 kDa MWCO) with Milli-Q water by centrifuging at 3,700 × g for 10 min at 4°C. Concentrate the Ni-NTA RPA pool to ~30 mL and store on ice for desalting into heparin buffer A (see Notes 19 and 20). 2. Equilibrate the HiPrep 26/10 Desalting column with 2 CVs of filtered Milli-Q water and 1.5 CVs of heparin buffer A. Equilibrate the HiTrap 5 mL Heparin HP column with 5 CVs of filtered Milli-Q water and 3 CVs of heparin buffer A (see Note 21). 3. Filter and load 10 mL of the Ni-NTA RPA concentrate onto the desalting column at 2.0 mL/min, collecting 4-mL fractions. The protein should elute within the first four fractions (16 mL) of the run. Re-equilibrate the column and repeat the run twice for the remaining 20 mL of RPA Ni-NTA concentrate. Store fractions on ice until all runs are complete. 4. Combine all three desalting pools (48 mL), filter at 0.45 μm, and load directly onto the equilibrated heparin column at 1.0 mL/min. 5. Wash out unbound sample with 3 CVs of heparin buffer A at 2.5 mL/min. 6. Elute RPA with a 20 CV gradient (0–100% heparin buffer B, 50 mM to 1 M NaCl), collecting 4-mL fractions at 2.5 mL/min. 7. Assess the presence of RPA from the A280 chromatogram trace and SDS-PAGE of relevant fractions (sampling 5 μL of each fraction). The elution should include two major peaks: the first corresponding to RPA70 exclusively and the second to trimeric RPA. Fractions containing all three RPA subunits are pooled for further final purification (typically, a 40-mL pool). 3.6.4. Superdex 200 Gel Filtration Chromatography

Gel filtration provides a final polishing step for the purification and ensures the removal of any trace RPA70 or low-molecular-weight contaminants. As before, loading and elution are implemented automatically using a pre-programmed Äkta FPLC method. 1. Equilibrate the Superdex 200 HR 10/30 column with 1.5 CVs of filtered Milli-Q water and 1.5 CVs of gel filtration buffer. 2. Pre-rinse two centrifugal concentrators (15 mL, 30 kDa MWCO) with Milli-Q by centrifuging at 3,700 × g for 10 min at 4°C. Concentrate the heparin RPA pool to ~300–500 μL (see Note 19). 3. Filter the concentrate by using a 0.22-μm centrifugal spin filter in a refrigerated (4°C) centrifuge and load onto the Superdex 200 HR 10/30 column at 0.3 mL/min, collecting 0.5-mL fractions for 1.5 CVs.

192

C.A. Brosey et al.

4. As before, assess the presence of RPA from the A280 trace and SDSPAGE of relevant fractions (sampling 5 μL of each fraction). 5. Before the final RPA fractions are pooled, acquire final A280 and A260 measurements by UV-Vis spectrophotometry to ensure that the selected fractions are DNA-free, as determined by A260/A280 ratios of 0.64 or less. 3.7. Preparation of Samples for NMR

1. Pre-rinse two centrifugal concentrators (15 mL, 30 kDa MWCO) with Milli-Q water by centrifuging at 3,700 × g for 10 min at 4°C. Concentrate the S200 RPA pool to ~500– 600 μL (see Note 19), monitoring the protein concentration by UV-Vis spectrophotometry (the gel filtration buffer may serve as a blank). 2. The target NMR concentration for RPA is ~100–130 μM (10–15 mg/mL) with a minimum sample volume of 260 μΛ (using a 4-mm-diameter NMR tube). Once the target concentration is reached, centrifuge 260–300 μL of the concentrate at 16,100 × g for 5 min at 4°C to remove any stray precipitation. 3. Load the protein into a standard 4-mm diameter NMR tube, which may be fitted with an adaptor to fit a 5-mm spinner or slipped into a 5-mm diameter tube containing 120 μL D2O without the need for the adaptor.

4. Notes 1. The tricistronic RPA pET15b vector was a gift from the lab of Alexey Bochkarev. 6×-His tags with thrombin cleavage sites precede the RPA70 and RPA14 subunits. The order of subunit open reading frames (ORFs) is as follows: RPA70, RPA14, RPA32. 2. Using freshly prepared ampicillin stock in the LB medium is important for ensuring robust RPA transformation. Even though the choice of the BL21(DE3) pLysS cell line is designed to circumvent leaky expression, even small amounts of noninduced RPA can potentially result in resistant cells with less than robust expression. 3. Antibiotic stocks may be aliquoted and stored at −20°C for future use. For long-term storage of ampicillin stocks, storage at −80°C is recommended. 4. The 70C domain of RPA contains a zinc-binding motif, for which ZnCl2 is included in the purification buffers. 5. The most effective imidazole concentrations in the Ni-NTA buffers will depend on how recently the Ni-NTA resin has been charged. For freshly charged resin, the imidazole concentration

11

Preparation of the Modular Multi-Domain Protein RPA…

193

for Ni-NTA buffer A is often raised to 30 mM for the first few purifications to compensate for the higher nonspecific affinity of the resin. 6. βME is added fresh to each buffer immediately prior to use. Buffers can be prepared without βME and stored at 4°C if their use is anticipated to last beyond 1–2 days. 7. Preparation of 1 L of each buffer should provide more than enough for the entire purification. 8. As the transformation efficiency of the RPA pET15b vector is low and the double-antibiotic selection with the BL21(DE3) pLysS strain is quite stringent, plating the entire transformation culture is recommended. 9. When transferring a selected colony to the test expression culture, be sure to leave a portion behind for future inoculation of the large-scale cultures. If the colony is too small to divide in this manner, the plate may be left at room temperature for half a day to allow the colony to regrow. 10. An empty gel tip box will suffice. The level of water should be enough to immerse the gel. 11. The 15NH4Cl is measured out and added directly to the sterile medium as a powder. This ensures that the labeled material is not wasted should a step prior to the inoculation fails. 12. As mentioned at the beginning of Subheading 3, the volume of deuterated minimal medium may be adjusted for small-scale testing. 13. Mixing of the deuterated minimal medium dry components should take place in a clean, dry container and be carried out as efficiently as possible to prevent exchange with ambient water vapor. If a container large enough to accommodate 6-L volume is not available, substitution of two 3-L containers with subsequent exchange and mixing between the two batches can be used to ensure the homogeneity of the medium across all cultures. 14. Performing this sterile transfer in the presence of a Bunsen burner flame is advised. 15. In our experience, growth in rich LB medium requires 3–4 h to reach the target A600 while growth in minimal medium requires 8–10 h. For deuterated minimal medium, this timeline extends to 1.5–2 days. To ensure that the induction occurred during daylight hours, cultures were switched to agitation at 20°C overnight after a full day of growth at 37°C and then returned to 37°C the next morning. The target A600 was reached during the afternoon of the second day. 16. RPA is susceptible to proteolytic cleavage, particularly at the unstructured 60–70 amino-acid linker that connects the 70N and 70A domains. Throughout the purification, it is essential

194

C.A. Brosey et al.

that all buffers are kept ice cold and that heating from other steps of the lysis (sonication, centrifugation) is kept to a minimum. 17. The ice-water bath serves as a heat sink during sonication of the lysate. 18. Initializing the FPLC system (pump washing, cleaning superloops, setting up fraction collectors), as well as equilibration of the Ni-NTA column, should occur prior to or concurrently with cell lysis to ensure that the clarified, filtered lysate can be loaded directly into the system as soon as it is available. 19. Centrifugal concentrators are usually spun in 10–15-min increments and carefully mixed with each addition of the Ni-NTA pool to prevent buildup and aggregation of RPA at the base of the concentrator. 20. The desalting resolution of the HiPrep 26/10 Desalting column (GE Healthcare) is 10 mL; that is, the column can effectively exchange 10 mL of injected sample into the target buffer without contamination from the original buffer. To avoid desalting too concentrated a volume of RPA and triggering aggregation, the Ni-NTA pool is processed in three sequential 10-mL batches. 21. As with the Ni-NTA step, initializing the FPLC system and equilibrating the desalting and heparin columns should occur prior to or concurrently with concentrating the RPA Ni-NTA pool to allow for immediate loading once the target volume is reached.

Acknowledgments The authors would like to thank Dr. Dalyir Pretto and Susan Meyn. This work was supported by the National Institutes of Health operating grant R01 GM65484 and graduate training grant T32 GM08320. References 1. Wold, M. S. (1997) Replication protein A: A heterotrimeric, single-stranded DNA-binding protein required for eukaryotic DNA metabolism. Annu. Rev. Biochem. 66, 61–92. 2. Fanning, E., Klimovich, V., and Nager, A. R. (2006) A dynamic model for replication protein A (RPA) function in DNA processing pathways. Nuc. Acids Res. 34, 4216–4137. 3. Stauffer, M. E., and Chazin, W. J. (2004) Structural mechanisms of DNA replication,

repair, and recombination. J. Biol. Chem. 279, 30915–30918. 4. Bochkarev, A., Pfuetzner, R. A., Edwards, A. M., and Frappier, L. (1997) Structure of the singlestranded-DNA-binding domain of replication protein A bound to DNA. Nature 385, 176–181. 5. Jacobs, D. M., Lipton, A. S., Isern, N. G., Daughdrill, G. W., Lowry, D. F., Gomes, X., and Wold, M. S. (1999) Human replication

11

6.

7.

8.

9.

Preparation of the Modular Multi-Domain Protein RPA…

protein A: Global fold of the N-terminal RPA70 domain reveals a basic cleft and flexible C-terminal linker. J. Biomol. NMR 14, 321–331. Bochkarev, A., Bochkareva, E., Frappier, L., and Edwards, A. M. (1999) The crystal structure of the complex of replication protein A subunits RPA32 and RPA14 reveals a mechanism for single-stranded DNA binding. EMBO J. 18, 4498–4504. Mer, G., Bochkarev, A., Gupta, R., Bochkareva, E., Frappier, L., Ingles, C. J., Edwards, A. M., and Chazin, W. J. (2000) Structural basis for the recognition of DNA repair proteins UNG2, XPA, and RAD52 by replication factor A. Cell 103, 449–456. Bochkareva, E., Korolev, S., Lees-Miller, S. P., and Bochkarev, A. (2002) Structure of the RPA trimerization core and its role in the multistep DNA-binding mechanism of RPA. EMBO J. 21, 1855–1863. Pervushin, K., Riek, R., Wider, G., and Wuthrich, K. (1997) Attenuated T2 relaxation by mutual cancellation of dipole-dipole coupling and chemical shift anisotropy indicates an avenue to NMR structures of very large biological macromolecules in solution. Proc. Natl. Acad. Sci. U.S.A. 94, 12366–12371.

195

10. Riek, R., Wider, G., Pervushin, K., and Wuthrich, K. (1999) Polarization transfer by cross-correlated relaxation in solution NMR with very large molecules. Proc. Natl. Acad. Sci. U.S.A. 96, 4918–4923. 11. Riek, R., Pervushin, K., and Wuthrich, K. (2000) TROSY and CRINEPT: NMR with large molecular and supramolecular structures in solution. Trends Biochem. Sci. 25, 462–468. 12. Tugarinov, V., Hwang, P. M., and Kay, L. E. (2004) Nuclear magnetic resonance spectroscopy of high-molecular-weight proteins. Annu. Rev. Biochem. 73, 107–146. 13. Brosey, C. A., Chagot, M. E., Ehrhardt, M., Pretto, D. I., Weiner, B. E., and Chazin, W. J. (2009) NMR analysis of the architecture and functional remodeling of a modular multidomain protein, RPA. J. Am. Chem. Soc. 131, 6346–6347. 14. Sambrook J and Russell D. (2001) Molecular cloning: A laboratory manual, vol. 3, 3rd ed. Cold Spring Harbor Laboratory Press, New York. 15. Li, M. X., Corson, D. C., and Sykes, B. D. (2002) Structure determination by NMR isotope labeling. Meth. Mol. Biol. 173, 255–265.

Chapter 12 NMR Studies of Protein–RNA Interactions Carla A. Theimer, Nakesha L. Smith, and May Khanna Abstract This chapter describes the preparation of NMR quantities of RNA purified to single-nucleotide resolution for protein–RNA interaction studies. The protocol is easily modified to make nucleotide-specific isotopically labeled RNAs or uniformly labeled RNA fragments for ligation to generate segmentally labeled RNAs. Key words: In vitro transcription, Single-nucleotide resolution, RNA synthesis, Protein–RNA interactions, RNA purification, Isotopic labeling

1. Introduction Understanding RNA–protein interactions and the structures of RNA–protein complexes is an important avenue of research. Ribonucleoprotein (RNP) complexes and RNA–protein interactions have been found to be central to many biological processes, ranging from genomic stability through telomeric maintenance (1–3), and alternative splicing of RNA by the spliceosome (4), to protein synthesis by the ribosome (5–8). Additionally, exciting new discoveries regarding small, noncoding RNAs, including snoRNAs, snRNAs, and RNA interference, continue to unfold. Many of these RNAs are associated with proteins at various stages of maturation, from trafficking to processing to their final destination, and any defect in these pathways can lead to serious diseases (9). Thus, dissecting the structure of RNA–protein interactions is crucial for deciphering the roles of these RNP complexes as well as identifying potential targets for pharmaceutical intervention. Protocols are well-established and documented for the generation of sufficient quantities of 13C-,15N-isotopically labeled proteins

Alexander Shekhtman and David S. Burz (eds.), Protein NMR Techniques, Methods in Molecular Biology, vol. 831, DOI 10.1007/978-1-61779-480-3_12, © Springer Science+Business Media, LLC 2012

197

198

C.A. Theimer et al.

from bacterial sources for NMR studies; the purification of such proteins uses removable protein tags (e.g., His6 and GST) and various permutations of affinity, ion exchange, and size-exclusion chromatography. In addition, methodologies for generating 13C-,15Nisotopically labeled amino acid-specific (10) or segmentally labeled proteins are also available (11, 12). Similar strategies for segmental and nucleotide-specific labeling are particularly important for the analyses of RNAs by using NMR spectroscopy (13, 14), given the extremely poor dispersion of RNA chemical shifts. Selectively labeled RNAs are particularly useful in combination with NMR experiments that utilize filtering and editing schemes to identify intramolecular NOEs (between labeled and unlabeled nucleotides in a single-RNA sequence for unambiguous assignment of overlapped chemical shifts) and intermolecular NOEs (between RNAs or in RNA– protein complexes) (15). However, since there is no commercially available source for chemically synthesized 13C-,15N-isotopically labeled RNA, these studies rely on the ability to synthesize large quantities of RNA in vitro in the laboratory. The following sections outline start-to-finish detailed protocols for the synthesis and purification of milligram quantities of RNA using in vitro transcription by T7 RNA polymerase, suitable for use in NMR studies (Fig. 1a).

Fig. 1. RNA synthesis and purification. (a) An RNA synthesis and purification flowchart, including the general steps from initial template design to NMR data collection. (b) DNA template design for RNA transcription. The RNA product is presented on the annealed DNA template/T7 Top promoter duplex in lower case letters. The DNA sequences of the T7 RNA polymerase promoter sequence and DNA template for transcription are presented in upper case letters. The two nucleotides at the 5¢ end of the template, which should have 2¢-OMe substitutions, are indicated by asterisks. (c) A representative 20% (19:1) acrylamide:bisacrylamide gel showing the observed bands from a typical analytical transcription experiment compared to the DNA template.

12

NMR Studies of Protein–RNA Interactions

199

2. Materials 2.1. DNA Template Design for RNA Synthesis

1. Template DNA: Complementary DNA template for RNA transcription, 0.1–1.0 mM in water, store at −20°C, Fig. 1b (see Note 1). 2. T7 Top DNA: Purified T7 RNA polymerase promoter DNA (coding strand, 5¢-TAATACGACTCACTATA-3¢), 1 mM in water, store at −20°C.

2.2. RNA Synthesis

1. 50 mM Annealed DNA template: T7 Top DNA stock solution in water, store at −20°C. 2. 10× Transcription buffer: 400 mM Tris–HCl, pH 8.0, 10 mM spermidine, 0.1% (w/v) Triton X-100; store at room temperature. 3. 100 mM dithiothreitol (DTT): Store in 1–2-mL aliquots at −20°C. 4. 100 mM ATP, CTP, GTP, and UTP: Dissolve nucleotides in water and adjust to pH 8.0 using 1M NaOH; store in 1–2-mL aliquots at −20°C (see Note 2). 5. 1M MgCl2: Store at room temperature. 6. T7 RNA polymerase/ribonuclease (RNAse) inhibitor mixture: T7 RNA polymerase in solution, dialyzed into 30 mM HEPES (adjusted to pH 7.5 with 1M NaOH), 0.1M potassium glutamate, 0.25 mM EDTA, 0.05% Tween-20, 1 mM DTT, 200 mM NaCl. Following dialysis, RNAse inhibitor (see Note 3) and glycerol (50% (w/v) final concentration) are added. Store in 1-mL aliquots at −80°C. 7. Gel running buffer (1× TBE): 90 mM Tris base, 90 mM boric acid, and 2 mM EDTA (using 0.5M EDTA, pH 8.0, stock solution). Typically made as a 5× or 10× concentrated stock and diluted as needed; store at room temperature. 8. Denaturing acrylamide gel solution: 20% acrylamide/bisacrylamide (19:1), 1× TBE, and 7.8 M urea; store at 4°C (see Note 4). 9. 10% (w/v) Ammonium persulfate solution (APS): Store at 4°C. 10. Denaturing gel loading buffer: 80% formamide, 10 mM EDTA (using 0.5M EDTA, pH 8.0, stock solution), 0.025% (w/v) xylene cyanol, and 0.025% (w/v) bromophenol blue; store at room temperature. 11. Toluidine blue stain: 0.25% (w/v) in water; prepare as needed, and store at room temperature. 12. 100% Ethanol; store at −20°C. 13. 0.5M EDTA stock solution: 0.5M EDTA, adjust to pH 8.0 using solid NaOH pellets. 14. N,N,N´,N´-tetramethyl-ethane-1,2-diamine (TEMED).

200

C.A. Theimer et al.

2.3. RNA Purification

1. Denaturing gel loading buffer, denaturing acrylamide gel solution, APS, and gel running solutions are identical to those described in Subheading 2.2. 2. Polyester-backed, silica-based, fluorescent thin-layer chromatography (TLC) plates. 3. Low-salt buffer: 10 mM monosodium phosphate/disodium phosphate buffer, pH 7.6 (adjusted with 1M NaOH or 1M HCl if the pH is off by more than 0.2 pH units), 1 mM EDTA, and 200 mM KCl; store at room temperature (see Note 5). 4. High-salt buffer: 10 mM monosodium phosphate/disodium phosphate buffer, pH 7.6 (adjusted with 1M NaOH or 1M HCl if the pH is off by more than 0.2 pH units), 1 mM EDTA, and 1.5M KCl; store at room temperature (see Note 5). 5. HiTrap-Q anion exchange column (5 mL prepacked). 6. 20% (w/v) Ethanol. 7. Amicon-pressurized stirred cell and 1,000 and/or 3,000 molecular weight cutoff (MWCO) membranes. 8. NMR buffer for preliminary RNA experiments or for RNA– protein interaction studies (typically, 10–20 mM sodium phosphate buffer, pH 6.0–7.0, 0–200 mM KCl). 9. 70% Ethanol. 10. 100% Ethanol. 11. Electroelution chamber.

2.4. Preliminary RNA Analysis by NMR

1. 99.99% D2O: Store in sealed ampoules at room temperature. 2. 4M potassium chloride: Store at room temperature. 3. Acid and base solutions for pH optimization: Usually, 0.1 and 1M hydrochloric acid and 0.1 and 1M potassium (or sodium) hydroxide; store at room temperature. 4. Sodium azide. 5. Standard and Shigemi NMR tubes.

2.5. RNA–Protein Interactions by NMR

1. Polyethyleneimine (PEI): 5% (w/v) solution at pH 7.9 (MW 50–100 K). 2. Solid NaCl. 3. Amicon-pressurized stirred cell and 1,000 and/or 3,000 MWCO membranes. 4. NMR buffer similar to that used for RNA experiments (see Subheading 2.3). 5. Shigemi NMR tubes, D2O matched. 6. Solid ammonium sulfate.

12

NMR Studies of Protein–RNA Interactions

201

3. Methods 3.1. DNA Template Design for RNA Synthesis

Commonly, a partially double-stranded DNA template is used for in vitro transcription of large quantities of RNA, since only the promoter region of the template DNA must be double stranded for T7 RNA polymerase to bind and initiate transcription (Fig. 1b) (16). Standard PCR can be used to make the DNA template completely double stranded if it improves transcription efficiency and the overall yield for a particular template, although we do not often find this to be necessary. The biggest issue for synthesizing RNA by using in vitro transcription is RNA product heterogeneity due to in vitro transcription artifacts (Fig. 1c). It has been demonstrated that, in vitro, T7 RNA polymerase can produce off-target products as a result of 5¢-heterogeneity, 3¢-heterogeneity, and RNAtemplated RNA addition (16–19). T7 RNA polymerase only synthesizes RNA products that contain at least one 5¢-G nucleotide. Increased transcription efficiency is observed with two or three 5¢-G nucleotides, but multiple G nucleotides at the 5¢-end of the RNA has been shown to cause 5¢-heterogeneity (additional G nucleotides) in the RNA product (18). To strike a balance between transcription efficiency and artifact generation, we prefer, when possible, to start sequences with no more than two G nucleotides in a row. 3¢-heterogeneity appears to be a result of the runoff transcription mechanism and often results in the nontemplated addition of one or two nucleotides (N + 1 and, less often, N + 2 products) (16). This problem can be diminished or overcome completely by having the DNA template synthesized with 2¢-methoxyl groups on the two terminal 5¢ nucleotides (20, 21), and we strongly recommend that template DNAs be synthesized with this modification; Fig. 1b. Typically, we see RNA-templated RNA addition products that vary in size from ~10–20 nucleotides longer than the expected transcribed product. Since this phenomenon is sequence dependent, some sequences display no extraneous long products and some sequences make large amounts of long products (17, 19). A study has been performed on DNA sequence mutations that reduce or abolish this behavior (22); when sequence alteration is not feasible, there are transcription conditions that can help to reduce this problem (19). In addition, strategies to eliminate 5¢- and 3¢-heterogeneity in RNA samples using ribozyme-based cleavages are also performed, as described in detail elsewhere (13). Finally, mass spectrometry is an excellent technique to check that the final RNA product is the correct length and has the correct sequence composition. The DNA template is complementary to the sequence of the RNA and the top strand of the T7 promoter sequence (Fig. 1b). Both DNA sequences for RNA transcription can either be ordered

202

C.A. Theimer et al.

from an oligonucleotide synthesis company typically at a 250-nmol to 1-mmol scale, unpurified, or synthesized on site if you have access to a DNA synthesizer. The chemically synthesized DNA should be desalted by the company and arrive as a lyophilized pellet. 1. Dissolve the pellet in 100–300 mL of water. Depending on the quality of the DNA synthesis, it may be necessary to purify the DNA template using denaturing polyacrylamide gel electrophoresis (PAGE), although we do not always do so. Use a UV/ VIS spectrophotometer and the molar extinction coefficient of the specific DNA sequence to calculate the concentration of the DNA template (see Note 6) and adjust to ~0.2–1 mM. A dilution between 1:100 and 1:1,000 should be sufficient for spectrometry. 2. Prepare 1 mL of 50 mM annealed stock DNA for transcription, which is enough for 50 mL of transcription at a final template concentration of 1 mM. Pipette 50 mL of 1 mM T7 Top DNA (total 50 nmol) into a sterile labeled Eppendorf tube. Add 50 nmol of the template DNA for RNA transcription (based on the calculated concentration) and water to make a total volume of 1 mL. 3. Vortex the tube containing the DNAs and heat it at 95°C for 5 min. 4. Allow the annealed DNA template to cool to room temperature. Use the DNA immediately or store for later use at 4 or −20°C. 3.2. RNA Synthesis

The transcription reaction is essentially the same for making selectively or uniformly 13C-,15N-isotopically labeled RNA as it is for unlabeled RNA. All nucleotides are prepared and stored separately so that one or more unlabeled nucleotides can be replaced with 13 C-,15N-isotopically labeled nucleotides for making selectively labeled RNA samples or all four unlabeled nucleotides can be replaced with 13C-,15N-isotopically labeled nucleotides for uniform labeling. The primary differences are as follows: (1) since labeled nucleotides are prohibitively expensive, we typically run labeled transcriptions at a concentration of 2 mM for each nucleotide and (2) we often explore additional transcription optimization conditions (additional transcription components), such as adding inorganic pyrophosphatase (IPP) or polyethylene glycol (avg. MW 8,000), for improved transcription efficiency (23). Before performing large-scale transcription reactions (10– 50 mL), it is necessary to determine the optimum transcription conditions for every new DNA template (Fig. 1c). The conditions that need to be considered include NTP concentration (2–4 mM),

12

NMR Studies of Protein–RNA Interactions

203

magnesium chloride concentration (15–50 mM), annealed DNA template concentration (0.5–1 mM), and T7 RNA polymerase/ RNAse inhibitor mixture concentration. We typically use a concentration of 4 mM for each NTP when working with unlabeled nucleotides, and ~2 mL of T7 RNA polymerase/RNAse inhibitor mixture per 100 mL test reaction. Test reactions are performed in 1.5-mL Eppendorf tubes (see Note 7). 1. Thaw the T7 RNA polymerase/RNAse inhibitor mixture at −20°C, and then keep on ice; handle very gently (see Note 8). 2. Thaw all other frozen components on ice (except for the T7 RNA polymerase/RNAse inhibitor mixture), and vortex them briefly to ensure homogeneity (see Note 9). 3. For a single-DNA template, run between 8 and 16 different test reaction conditions, each reaction consisting of a total of 100 mL. Add 10 mL of 10× transcription buffer (1× final concentration) and 2.5 mL of 100 mM DTT (2.5 mM final concentration) to each of the labeled test reaction tubes. 4. Based on the exact conditions to be tested, add 1–2 mL of 50 mM annealed DNA template (0.5–1.0 mM final concentration), 2–4 mL of each 100 mM NTP stock (2.0–4.0 mM final concentration), and 1.5–5.0 mL of 1M MgCl2 (10–50 mM final concentration) to each tube. 5. Add water to bring the volume to 100 mL, leaving space for the T7 RNA polymerase/RNAse inhibitor mixture to be added later. Vortex the samples and centrifuge in a standard tabletop at 14,000 × g for 3 min at room temperature to collect the solution droplets. 6. Add 1–5 mL of T7 RNA polymerase/RNAse inhibitor mixture to the samples, pipetting the solution up and down gently to mix in the polymerase and to prevent denaturation of the protein. Always add the T7 RNA polymerase/RNAse inhibitor mixture last. 7. Incubate the samples at 37°C for 2–4 h. 8. When the test transcription reactions are complete, centrifuge in a standard tabletop at 14,000 × g for 3 minutes to pellet precipitated magnesium pyrophosphate, and transfer 10 mL of each reaction into clean labeled 1.5-mL Eppendorf tubes and add 10 mL of denaturing gel loading buffer to each tube. A standard sample is also made containing 5 mL of the annealed DNA template, 5 mL of water, and 10 mL of denaturing gel loading buffer. The samples are heated at ~95°C for 5 min immediately prior to loading on the denaturing gel. 9. These instructions assume the use of a vertical electrophoresis system (for example: ASU-250, C.B.S. Scientific) and 17-cm-length gels, although any vertical gel apparatus with a reasonable gel

204

C.A. Theimer et al.

size can be used. Rinse the glass plates and spacers with water, followed by ethanol, and wipe dry immediately prior to use. Although very small gels can be made using standard protein SDS-gel apparatus, we do not advise this as the resolution on such gels is generally too low for single-nucleotide resolution and thus not satisfactory for this purpose. 10. Prepare a 0.75-mm-thick 20% gel by mixing 50 mL of denaturing acrylamide gel solution with 500 mL of 10% APS. To this solution, mix in 50 mL of TEMED and immediately pour the gel. These gels are a single layer and the comb (10, 14, or 20 well) is inserted immediately after the entire gel is poured. The gel takes approximately 30 min to polymerize. The percentage of acrylamide:bisacrylamide solution used depends on the size of the RNA product that is run on the gel. We typically use 20% for RNA transcripts up to 40–50 nucleotides long, 15% for RNA transcripts between 45 and 70 nucleotides, and 10–12% for longer RNA transcripts. 11. When the gel is polymerized, remove the comb and carefully rinse the wells with water by using a 30-mL syringe equipped with a 22-gauge needle. Place the gel in the apparatus and fill the buffer chambers with gel running buffer. The gel should be pre-run at 150 V (or 15 W) for 15–20 min and the wells rinsed again with gel running buffer prior to loading the samples. 12. Once the samples are loaded, run the gel for 2–3 h at 150 V or until the bromophenol blue dye front (dark blue) is within ~3 cm of the bottom of the gel. 13. Stain the gel with toluidine blue stain on an orbital shaker for 15 min. Destain the gel with water (multiple exchanges) until you observe good contrast between the dark blue nucleic acid bands and the background of the gel. The optimal transcription conditions are chosen based on the test conditions that produce the highest intensity of the RNA product band. 14. Once the optimal solution conditions have been identified, the large-scale transcription reaction can be performed. Generally, we perform a 30 mL transcription reaction in a sterile, disposable, blue-capped 50-mL centrifuge tube. We find that this is a large enough transcription volume to generate a reasonable quantity of RNA for NMR samples (150–400 nmol of RNA) for most RNA sequences, although (rarely) some RNA sequences transcribe much better or much worse than this. The large-scale reaction is a direct scale-up of the small-scale reaction. The only differences are the volumes of reagents used and that it is usually not necessary to centrifuge the solution after vortex mixing. The T7 RNA polymerase/RNAse inhibitor mixture is still the last ingredient added before incubation and the tube is swirled gently after adding the polymerase.

12

NMR Studies of Protein–RNA Interactions

205

15. Incubate the reactions for 4–8 h at 37°C. The reactions can be incubated overnight, although this can be risky if there is any possibility of RNAse contamination or if the RNA sequence is prone to undesirable side reactions, like RNA-primed RNA addition by T7 RNA polymerase, as described above (see Note 10). 16. Centrifuge the reaction solution at 4,000 × g for 5 min at 4°C and decant the supernatant, which contains the RNA product, into a sterile 250-mL centrifuge bottle. There is a large pellet from precipitated inorganic pyrophosphate; this does not contain any RNA and can be discarded. 17. Add 1/10th the volume of 0.5M EDTA, pH 8.0 (1 mL of EDTA for every 10 mL of transcription), swirl to mix, and then add 2.5–3 times the volume of cold 100% ethanol (~75–90 mL of ethanol for a 30 mL transcription) to the transcription solution. 18. Place the centrifuge bottle containing the RNA product at −20°C overnight to precipitate (see Note 11). 3.3. RNA Purification

Denaturing PAGE is our primary means of purifying large quantities of RNA to single-nucleotide resolution for NMR. Although we have not experimented recently with currently available preparative-scale HPLC columns, in the past we found that HPLC purification would not purify RNAs larger than ~30 nucleotides to single-nucleotide resolution on a preparative scale. In addition, while we frequently purify small quantities of RNA using native PAGE, the native gels must be run in the cold room (4°C), typically take a long time to run, and rarely yield single-nucleotide resolution when the single-band products are checked on analytical denaturing gels for purity. 1. Centrifuge the bottle at 14,000 × g for 45 min at 4°C. 2. Very gently, decant the supernatant, as soon as the centrifuge stops running (see Note 12). There should be a visible white pellet on the wall of the bottle. The size of the pellet is not a direct reflection of product yield, since the majority of the size of the pellet is due to salt precipitation. 3. If the pellet is very large, put 20–50 mL of cold 70% (w/v) ethanol in the bottle very gently to wash away excess salt. Centrifuge with the pellet located against the outer wall at 14,000 × g for 30 min at 4°C, and again decant the supernatant immediately (see Note 13). 4. Dry the pellet by placing the capless centrifuge bottle underneath the hood, angled so that the residual ethanol is not lying directly on top of the pellet. Evaporating off all residual ethanol typically takes about 2 h if the supernatant was properly poured off without disturbing the pellet.

206

C.A. Theimer et al.

5. When the pellet has completely dried, put 2–3 mL of water in the bottle directly over the pellet and allow it to sit for 10 min. Pipette up and down gently to resuspend and transfer to a sterile, disposable 15-mL centrifuge tube. Rinse the bottle with an additional 1–2 mL of water to gather up any remaining sample and transfer to the centrifuge tube. 6. Add denaturing gel loading buffer to the sample. Use 1:1 RNA solution:gel loading buffer; although if the RNA solution volume is too large, you can use 2:1 RNA solution:gel loading buffer and it loads and runs fine on the purification gels. 7. These instructions assume the use of gel electrophoresis equipment for large, sequencing gels, 20 cm (w) × 42 cm (l) (for example: DDH-400-20, C.B.S. Scientific), spacers, and 3-well preparative scale (3-mm thick) combs. Rinse the glass plates, combs, and spacers with water, followed by ethanol, and wipe dry immediately prior to use. 8. Pour the number of gels needed to obtain the appropriate purification level (see Note 14). Preparative-scale gels typically take ~400 mL of acrylamide solution to fill each gel. Place 400 mL of denaturing acrylamide gel solution into a beaker, add 4 mL of 10% APS solution, and stir briefly to mix the APS in evenly. Add 400 mL of TEMED and stir the solution again briefly. In general, a 1/100 ratio of APS and 1/1,000 ratio of TEMED to gel solution are used for polymerization. The gel must be poured immediately at this ratio of APS and TEMED. The gels take 2 h to completely polymerize and cool, although they can sit for longer or even overnight. 9. Remove the combs and clean the wells by using a wash bottle of water. It is important to also carefully clean the back plate of the gel, which is in contact with the ceramic heat exchange plate, to prevent poor contact and smiling of the gel. Set up the gels in the sequencing apparatus and fill the upper and lower buffer chambers with gel running buffer. The gels should be pre-electrophoresed for 20–30 min at 20 W per gel, and the wells thoroughly rinsed with 1× TBE prior to loading the RNA samples. 10. Heat the RNA samples at ~95°C for 5 min immediately prior to loading on the purification gels. Load the gels with a standard 1,000 mL pipettor, since the tip easily fits down into the 3-mm wells. 11. The size of the RNA dictates how long the gels should be run and the wattage. Typically, we run these gels overnight (~18 h) at 20 W per gel for smaller RNAs (15–25 nucleotides), 25 W per gel for medium-sized RNAs (25–40 nucleotides), and 30 W per gel for larger RNAs (over 40 nucleotides). The gels are monitored based on the location of the dyes and the previous observation from test transcription gels, of where the RNA runs compared to the xylene cyanol and bromophenol blue dyes (see Note 15).

12

NMR Studies of Protein–RNA Interactions

207

12. Remove the gel from the apparatus, carefully remove one glass plate, and cover the gel with plastic wrap. Flip the gel over (plastic wrap on the bottom) onto a fluorescent TLC plate and gently remove the top glass plate. When a handheld shortwavelength UV light (254 nm) is shone onto the gel, the plate glows green and the RNA band is visible as a grey to black shadow on the fluorescent green background (UV shadowing). Carefully cut the product bands out of the gel with a clean razor blade and transfer them to a sterile, disposable 50-mL centrifuge tube and discard the rest of the gel. Repeat for all of the gels (exposure to UV light should be minimized to avoid UV damage of RNA). 13. Elute the RNA product by using a 4-trap Elutrap electroelution chamber (Whatman), with 1× TBE as the running buffer. Gel slices should be cut into small pieces but not crushed, and placed into the gel holding chamber between the BT1 and BT2 membranes (see Note 16). 14. Run the elution at 150 V and collect the RNA sample from the trap at 2–3-h intervals, typically for 9 h. For longer RNAs, we frequently reduce the voltage to 50 V and run overnight to ensure complete elution of the product from the gel slices. The progress of the elution can be tracked by calculating the amount of RNA in the eluant at each time point, based on UV absorbance at 260 nm. Time points should be stored at −20°C until all of the RNA has been collected, and the elution is finished. 15. Thaw and pool all of the eluted RNA fractions. Load this material onto an HiTrap-Q anion-exchange column hooked up to a peristaltic pump (flow rate 3–5 mL/min) and pre-equilibrated with low-salt buffer. Wash the column with ten-column volumes (50 mL) or more of low-salt buffer. Elute the RNA with high-salt buffer and collect 4-mL fractions. The purified RNA is typically found entirely in the second and third fractions. The column must always be thoroughly rinsed with low- and high-salt buffers between samples and immediately stored in 20% ethanol when not in use to prevent contamination. 16. Pool the two RNA containing fractions (8 mL) in a 50-mL centrifuge tube, add 24 mL of cold 100% ethanol, and store the RNA overnight at −20°C to precipitate. 17. Remove the centrifuge tube directly from the freezer, make a weight-matched balance tube, and centrifuge at 15,000 × g for 45 min at 4°C. Very gently, decant the supernatant as soon as the centrifuge stops running. At this point, it is possible to again perform a 70% ethanol wash as described above (step 3). 18. Dry the pellet by placing the capless centrifuge tube underneath the hood, angled so that the residual ethanol is not lying directly on top of the pellet. Evaporating off all residual ethanol typically takes about 2 h.

208

C.A. Theimer et al.

19. Dissolve the RNA sample in 50 mL of water or NMR buffer, if you already know what the required buffer should be, based on the protein to be investigated. For new RNAs under investigation, desalt the RNA (step 21) into water for pH and salt titrations in the NMR to determine optimal conditions for obtaining a single-RNA conformation in solution. 20. Anneal the RNA by placing the 50-mL tube at ~95°C for 5 min and then slow cool to room temperature on the benchtop (see Note 17). 21. When cool, load the solution into an Amicon-stirred cell (MWCO 3,000 or 1,000 membrane, depending on the size of your RNA), pressurize the chamber with 55 psi of nitrogen or argon, and stir. Concentrate the RNA down to ~1–2 mL and then add fresh buffer or water up to the 50-mL line. Perform at least three washes to remove excess salt. Concentrate the RNA down to ~0.5–1 mL (~0.2–2 mM) and transfer to a sterile 1.5-mL Eppendorf tube. The RNA is ready for NMR studies and should be stored at −20°C when not in use. 3.4. Preliminary RNA Analysis by NMR

Once an initial unlabeled RNA sample is made, a basic set of NMR experiments is run, including 1D 1H NMR experiments collected in 95% H2O/5% D2O (5–10°C) for pH and salt titrations, NOESYs collected in 95% H2O/5% D2O (5–10°C) and D2O (20–30°C), and a TOCSY collected in D2O (20–30°C), to assess the properties of the RNA in solution at NMR concentrations. Optimal salt and pH conditions are assessed by the number, intensity, and line widths of the detectable imino proton resonances compared to the number of imino protons which are expected to be protected from rapid exchange due to hydrogen bonding in Watson-Crick and noncanonical base pairs (Fig. 2). Generally, we find that the optimal pH for RNA samples falls between 6.0 and 7.0 and the optimal salt conditions vary from 0 to 200 mM monovalent salt, depending on the ability of the sequence to form alternative conformations (see Note 18). The optimal solution conditions are typically dictated by the protein of interest, but it is important to be aware of the expected behavior of the RNA under the appropriate solution conditions. Degradation products and alternative conformations (including unwanted dimerization) can be identified both from additional imino proton resonances in the 1D and 2D spectra collected in 95% H2O/5% D2O (Fig. 2). In addition, the 1D 1H spectrum helps identify any small-molecule contaminants in the RNA sample. The most common contaminants in RNA samples (and their causes) are as follows: acrylamide and/or urea (insufficient volume of low-salt wash of the anion-exchange column before eluting the RNA product), EDTA and/or ethanol (too few buffer exchanges during desalting and concentration), and ethanol and/or glycerol (insufficient soaking and rinsing of the Amicon membrane prior to installing in the

12

NMR Studies of Protein–RNA Interactions

209

Fig. 2. 1D and 2D imino proton NMR spectra of RNA. The secondary structure of a hairpin RNA and the imino proton region of 1D and 2D NOESY 1H spectra (500 MHz) of the RNA in 95% H2O/5% D2O at 10°C demonstrate the presence of sharp imino proton resonances. A clear sequential walk through the stem base pairs is indicated for the expected hairpin stem. The smaller imino proton resonances and NOE peaks are due to an alternative conformation in solution.

stirred cell). EDTA can be eliminated from the high- and low-salt anion-exchange buffers if it is a recurring contaminant in the final samples. Ethanol contamination can easily be removed by freezing the sample in liquid nitrogen, lyophilizing the sample to dryness, and resuspending in 95% H2O/5% D2O (repeated as necessary). Glycerol typically requires exhaustive dialysis or buffer exchanges in the Amicon-stirred cell to be completely removed. 1. For titrations, the RNA sample was previously concentrated into water during the last steps of RNA purification (Subheading 3.3, steps 19–21). Add 5% (w/v) 99.99% D2O to the RNA sample (usually, ~0.1–0.3 mM RNA for solution condition optimization) and adjust to the starting pH (~pH 5.5) using acid and base solutions (see Note 19). 2. Transfer the sample to a standard NMR tube, insert the NMR tube into the spectrophotometer, and allow the sample to equilibrate to 10°C for 5 min before the instrument is set up for data collection. 3. Collect a baseline 1D imino proton spectrum (using 1,1 echo water suppression) for a low-salt, low-pH (~5.5–6.0), RNA sample at 10°C; usually, 128 scans are sufficient (see Note 20).

210

C.A. Theimer et al.

4. After each spectrum is collected, transfer the RNA sample into a 1.5-mL Eppendorf tube, adjust the pH by 0.5 pH units using the appropriate acid and base solutions, return the sample to the NMR tube, and allow it to equilibrate to 10°C for 5 min in the spectrophotometer before the setup for data collection. Due to increased RNA degradation under basic conditions, we typically do not take RNA samples above pH 8.0 unless absolutely necessary. 5. After the NMR spectra are collected and compared, identify an optimal pH based on the number of expected and observed peaks, intensities of the peaks, line widths, and pH at which there is the least peak overlap in terms of chemical shift. The optimal pH for most RNA samples typically lies between pH 6.0 and 7.0. 6. For salt titrations, use a new RNA sample that is already adjusted to the correct pH. The same procedure is followed for salt titrations. Typically, use either 25- or 50-mM increments for the monovalent salt concentration, aliquoted from a 4 M KCl stock, and choose the best conditions again based on the number of expected and observed peaks, intensities of the peaks, line widths, and monovalent salt concentration at which there is the least peak overlap in terms of chemical shift. If the optimal conditions for the RNA are consistent with the conditions needed for protein stability and protein–RNA interactions, all of the RNA from the NMR samples can then be pooled with the stock RNA and exchanged into the appropriate NMR buffer at this point as described in RNA purification (steps 19–21). 3.5. RNA–Protein Interactions by NMR

This section focuses on preparing the protein for investigating RNA– protein interactions. Protein purifications are well-established and isotopic labeling of proteins has been reviewed extensively (12, 24, 25). Other than the standard issues and concerns for preparing NMR quality protein samples, there are two issues that also need to be considered. First, RNAses can bind to or copurify with the protein of interest and may potentially degrade the RNA during complex formation. Second, high-affinity cellular RNA can copurify with the protein. Usually, several different purification steps, such as affinity tag purification, followed by ion-exchange and size-exclusion chromatography, are sufficient to ensure complete removal of RNAses (26). If it is also necessary to remove RNA bound to protein, this can be done through treatment with 0.1–0.5% PEI (27) to ensure removal of all RNA bound to the protein of interest. Uniformly 15N-labeled protein is typically obtained by overexpression from E. coli in M9 minimal medium and purified using affinity columns for either His6- or GST-tagged proteins. This step is often followed by size-exclusion chromatography. Depending on the purity of the protein, as judged by Coomassie staining of SDS

12

NMR Studies of Protein–RNA Interactions

211

polyacrylamide gels, it may be necessary to include a third-column purification step, such as anion-exchange chromatography. Following these purification steps, there should be no RNAse contamination left in the purified protein. However, there may still be nucleic acid contamination, which can be detected by running the purified protein on an agarose or acrylamide gel and staining with ethidium bromide. If nucleic acid contamination is detected, a PEI precipitation following the first affinity column can ensure complete removal. 1. Pool the appropriate fractions that contain the protein of interest from the affinity column and add solid NaCl to a final concentration of 1M. 2. Place all samples at 4°C, add PEI to a final concentration of 0.5% (w/v), and incubate the solution for 1 h. 3. Centrifuge the solution at 15,000 × g for 35 min at 4°C to remove the precipitated PEI–nucleic acid complexes. 4. Recover the protein from the supernatant through precipitation by adding ammonium sulfate to a final concentration of 75% (see Note 21). 5. Centrifuge the protein solution at 10,000 × g for 20 min at 4°C, decant the supernatant, and recover the pelleted protein by dissolving in low-salt buffer (Subheading 2.3), if the next column is an anion-exchange column, or in the final NMR buffer (Subheading 2.3), if the next column is a size-exclusion column. The final buffer should be identical to the buffer chosen for optimal RNA stability (see Subheading 3.4). 6. A final column purification step, such as an ion-exchange column, can be inserted at this step, if the previous purification steps do not yield protein of sufficient purity. 7. Pool all the fractions that contain the protein of interest and concentrate the protein to 0.2–1 mM concentration using either Amicon-pressurized stir cells or disposable Amicon ultra centrifugal filters (Ultracel). 8. Monitor complex formation between the RNA and protein by acquiring a 1H–15N heteronuclear single-quantum correlation (15N-HSQC) spectrum of the 15N-labeled protein (isotopic labeling of proteins has been extensively reviewed elsewhere (11)). 9. First, collect the 15N-HSQC spectrum of the free protein. The 2D 15N-HSQC should be acquired from 6 to 13 ppm in the hydrogen dimension and 95–140 ppm in the nitrogen dimension, which is typically referred to as the fingerprint region of proteins (see Note 22). 10. Prepare the protein sample in a Shigemi NMR tube using 300 mL of a 0.2 mM protein sample in 5% D2O to obtain a good quality spectrum and define the effect of buffers on the overall fold of the protein.

212

C.A. Theimer et al.

11. Following the initial test of protein folding, measure the chemical shift perturbation in the 15N-HSQC of the amide protons from the free protein upon titration with unlabeled RNA. The protein should be between 0.1 and 0.5 mM and the unlabeled RNA has to be concentrated to obtain different ratios for the titration ranging from 0.25 to 2:1 of RNA:protein without requiring the need for large volumes of RNA solution to be added to the NMR sample (see Note 23). 12. Since each 15N peak on the protein represents a distinct amide peak of the protein, this titration can yield information on the position of the RNA binding site at the protein interface as well as the binding affinity, based on the NMR timescale. 13. The reverse titration (protein into the RNA sample) can also be performed. First, using the same purification protocol, obtain a concentrated, unlabeled protein (1–2 mM) that can be titrated into a 13C-,15N-labeled RNA sample, which is purified as described above for unlabeled RNA (see Subheading 3.3). 14. Obtain a 13C-HSQC of RNA, which typically exhibits distinct peaks for the carbons attached to the aromatic and H1¢ protons. Binding to the protein should cause perturbations of these peaks, and thus yields information of the binding site on the RNA (28) (see Note 24). Once a few sites on the protein or RNA have been identified using the chemical shift perturbation method, other more targeted experiments narrow down the binding region (for a review of different experiments and how they can be applied to solve the RNA–protein complex, see ref. 26). Ultimately, measuring NOEs between the RNA and the protein provides the most important details for the structural calculations of the RNA– protein complex. Filtered and edited NOESY experiments (outlined in the introduction based on ref. 15) have been instrumental in identifying NOEs for RNA structure determination and greatly simplify the identification of NOEs between the RNA and protein components of the RNA–protein complex.

4. Notes 1. All water for RNA studies must be distilled deionized 18.0 MW-cm quality from a purification system that is properly maintained (cartridges replaced as suggested by manufacturer). Large bottles of autoclaved water are kept on hand in the laboratory for solution making, and all solutions that come into contact with RNA are sterile filtered using Nalgene sterile filter units with a 0.2-mm pore size prior to storage. Unless otherwise noted, all solutions should be handled in this manner and “water” in the

12

NMR Studies of Protein–RNA Interactions

213

context of this protocol indicates autoclaved water of this quality which has been sterile filtered prior to use. As long as the water quality is maintained, we have not found it necessary to treat water with diethylpyrocarbonate (DEPC) in order to prevent RNAse contamination and degradation of RNA samples. 2. It is important that the pH of the nucleotides is adjusted using sodium hydroxide and not Tris base. High Tris base concentrations appear to inhibit transcription by T7 RNA polymerase. 3. For the final solution of T7 RNA polymerase/RNAse inhibitor mixture, a tube (10,000 U) of SUPERase·In RNase Inhibitor (Ambion/Applied Biosystems) is mixed with the enzyme prior to the addition of glycerol to a final concentration of 50% (w/v), prior to making aliquots and storing at −80°C. RNAse inhibitor is not essential, but is added as a precaution against any residual RNAse from the T7 RNA polymerase enzyme purification. The final concentration of T7 RNA polymerase is rarely measured; we use test transcriptions to identify the amount of each preparation to use for optimal RNA transcription efficiency. 4. It is important when making this solution that it is not diluted to volume until all of the urea has dissolved (very-high-quality ultrapure urea should be used) and that it is not heated. When using stored acrylamide–urea solutions, the solution must be examined to ensure that urea has not precipitated out of solution. It is for this reason that we store these solutions in transparent brown glass bottles. In addition, since acrylamide is a cumulative neurotoxin, it is important that gloves are worn at all times even when it is polymerized. We prefer to buy premixed acrylamide to reduce handling. 5. The low- and high-salt buffers for anion exchange can use different buffer salts and sodium chloride, rather than potassium chloride. We use phosphate buffer and potassium chloride for this column because this is basically the last step of RNA purification and they are the usual buffer and salt compounds that we use for RNA NMR studies. 6. If the extinction coefficient for a DNA is not provided in the documentation from the company, it can be calculated (for DNA or RNA) using the extinction coefficients for individual NMPs (A = 15.4 M-1-cm-1, C = 7.4 M-1-cm-1, G = 11.8 M-1-cm-1, T = 9.6 M-1-cm-1, U = 9.9 M-1-cm-1) or a good approximation is to set the extinction coefficient at 10 M-1-cm-1 multiplied by the number of nucleotides. It is not advisable to use the general assumption that an A260 of 1 corresponds to 50 mg/mL as this is not accurate for short DNAs or short RNAs. 7. All disposable plasticware (pipette tips and Eppendorf tubes) that do not come presterilized and certified RNAse and DNAse free should be autoclaved prior to use and kept in sealed

214

C.A. Theimer et al.

containers to prevent contamination. We also cover the openings of glassware with aluminum foil and autoclave all glassware in the laboratory routinely after washing and rinsing with distilled deionized water. 8. Although it is possible to use purchased T7 RNA polymerase for in vitro transcription, the amounts necessary for large-scale synthesis of RNA for NMR tend to be prohibitively expensive. Therefore, most laboratories, ours included, purify their own supply of T7 RNA polymerase for transcription. The plasmid for His6-tagged T7 RNA polymerase is readily available from any number of academic sources. In general, we do not quantitate the T7 RNA polymerase; we simply use analytical transcription reactions to determine the appropriate amount of T7 RNA polymerase to be used for optimal transcription efficiency. 9. These solutions do not freeze in a homogeneous fashion, nor do they thaw that way. Before pipetting out of a thawed tube of solution, you must make sure that the solution is completely thawed, then vortex mix the tube briefly to make sure that it is well-mixed, and centrifuge the solution briefly to push all the droplets to the bottom of the tube. 10. If the reaction is working, it will start to get cloudy 1–2 h after you put the reaction into the water bath. The reaction gets cloudy because, as nucleotide triphosphates are linked together to form RNA (which has only one phosphate between each nucleotide), the other two phosphate groups are released as pyrophosphate (PPi). The pyrophosphate interacts with the magnesium in solution to form an insoluble magnesium pyrophosphate complex. The more precipitation you observe, the more nucleotide triphosphates have been converted into RNA. 11. If you use 2.5–3× the volume of 100% ethanol, your final solution will be 70–75% ethanol. It is not recommended to leave a sample in ethanol for more than a few days. The longer the sample sits, the more salt precipitates out of solution. The greater the amount of salt in the precipitate, the bigger effect it has on the separation efficiency and running of the purification gels. Ideal timing for RNA ethanol precipitation is either overnight at −20°C or for a few hours at −80°C. 12. It is important to pour off the supernatant as soon as the rotor stops spinning, before the pellet detaches from the wall of the bottle. 13. This step removes excess salt and helps the purification gels to run with better resolution. It is critical that you do not resuspend the pellet or rinse it off the wall with rough handling or it is very difficult to recover without significant loss of sample. 14. The number of gels needed to purify your RNA needs to be determined before you run your entire RNA sample on the gels. Each purification gel has three wells that can be loaded with

12

NMR Studies of Protein–RNA Interactions

215

different amounts of your dissolved RNA: loading buffer mixture. Load three different amounts of RNA in the three wells and run one gel to see which amount gives you a good-sized RNA band. Generally, loading amounts of the dissolved RNA solution that correspond to ~2, 4, and 6 mL of the original transcription give you a good idea what the proper amount for optimal purification might be. The RNA band should be somewhere between the width of a pencil and the width of your finger. Much less than this is hard to find on the gels and any more than that eliminates single-nucleotide resolution. The optimal loading amount is used for subsequent gels for this specific RNA. 15. To obtain the full separation capacity of the 42-cm-long gel, it is important to run the RNA to the bottom 1/3 of the gel or as close to the bottom as possible. The location of the RNA of interest on the gel with respect to the dyes should be noted at the test-preparative-gel stage. When purifying RNAs that run very close to the dye front (an ~27-nucleotide RNA runs on top of the xylene cyanol dye on a 20% 19:1 denaturing acrylamide gel), it is necessary to omit the overlapping dye from the samples completely and run a lane on one of the gels that contains only dye as a marker. 16. The MWCO for the BT1 membrane is ~5,000. To prevent loss of smaller product RNAs (~30 nucleotides or less), the BT1 membrane should be replaced with thoroughly rinsed 1,000 MWCO dialysis tubing for these samples. 18-mm wide dialysis tubing fits exactly in the trap chambers and only needs to be cut at the open ends to fit the length of the BT1 membrane. It is also possible to make a second trap by placing an additional BT1 membrane after the next insert in the trap. It is absolutely critical that elution traps are not overloaded with gel slices, which significantly reduces elution efficiency. 17. The conditions for annealing should be based on the behavior of the RNA in question. RNAs that are intended to form unimolecular structures should be annealed under dilute conditions (50 mL) while RNAs that are intended to form duplexes (dimers) should be annealed in more concentrated solutions (2–5 mL). In addition, snap cooling on ice can be used to affect the conformational exchange in solution. The timing of adding monovalent salts (before annealing or immediately after snap cooling) can be used to bias the sample toward forming a single conformation as needed. The appropriate annealing conditions require trial and error for each RNA sample. 18. We typically include 0.2% (w/v) sodium azide in RNA samples to prevent bacterial contamination. Usually, the sodium azide is included in the NMR buffer solution used for the final stages of RNA concentration and desalting (RNA purification: steps 19–21), after the optimal NMR solution conditions have been identified. While divalent salts, such as MgCl2, do stabilize

216

C.A. Theimer et al.

RNA structures and are often needed for larger RNA structures to form, completely small RNAs typically fold without magnesium, and the NMR samples last longer (degrade less quickly) if magnesium is excluded from the NMR buffer, whenever possible. 19. It is very important that, when the pH of RNA samples in water is being adjusted, you are careful not to overshoot the desired pH. Alternately adding acid and base to reach the appropriate pH results unnecessarily increases the salt concentration, which also affects the intensity of the imino proton resonance peaks. 20. It is important to ensure that the sweep width is wide enough not to miss any unusual imino proton chemical shifts (protonated cytosine residues, for example). Therefore, we routinely collect out to 18 ppm initially, although almost all imino proton peaks are found between ~9 and 15 ppm. 21. PEI can be very difficult to remove from a protein preparation, as it cannot be dialyzed out effectively. Ammonium sulfate precipitation is the standard method of separating the protein of interest from the PEI, with the protein precipitating and the PEI remaining in the supernatant solution. The precise percentage of ammonium sulfate necessary is protein dependent and the optimal percentage to be used should be tested on small samples of the protein to ensure maximal protein recovery. 22. Examining the fingerprint region of the free protein spectrum: If the peaks are well-dispersed with minimal overlap in the center region of the spectrum, this is an indication that the protein is well-folded. Each peak in this spectrum represents a unique amide peak from the protein. If the protein is not well-folded, it is still worth testing the titration with RNA, as it is possible that the RNA may induce conformational change of the protein upon binding. 23. It is best to start with a lower end of the range of protein concentrations since formation of RNA:RNA multimers can be an issue at high concentrations. This titration also gives some idea of the stoichiometry of RNA:protein complex. If a large amount of RNA (high RNA:protein ratio) must be added to observe chemical shift perturbations, this may be an indication that measuring NOEs between the RNA and protein is difficult. 24. One must keep in mind that it is possible to get shifted resonances (on the protein or the RNA) not only in the binding site, but also in other regions due to allosteric modulation or, more likely, changes in the pH or monovalent salt concentration of the solution. This is why it is particularly important to try to have both the RNA and protein prepared such that their final solution conditions are identical.

12

NMR Studies of Protein–RNA Interactions

217

References 1. Autexier, C., and Triki, I. (1999) Tetrahymena telomerase ribonucleoprotein RNA–protein interactions. Nucl. Acids Res. 27, 2227–2234. 2. Bachand, F., Triki, I., and Autexier, C. (2001) Human telomerase RNA–protein interactions. Nucl. Acids Res. 29, 3385–3393. 3. Greider, C. W., and Blackburn, E. H. (1987) The telomere terminal transferase of Tetrahymena is a ribonucleoprotein enzyme with two kinds of primer specificity. Cell 51, 887–898. 4. Staley, J. P., and Woolford, J. L., Jr. (2009) Assembly of ribosomes and spliceosomes: complex ribonucleoprotein machines. Curr. Opin. Cell Biol. 21, 109–118. 5. Ban, N., Nissen, P., Hansen, J., Moore, P. B., and Steitz, T. A. (2000) The complete atomic structure of the large ribosomal subunit at 2.4 A resolution. Science 289, 905–920. 6. Cech, T. R. (2000) Structural biology. The ribosome is a ribozyme, Science 289, 878–879. 7. Schluenzen, F., Tocilj, A., Zarivach, R., Harms, J., Gluehmann, M., Janell, D., Bashan, A., Bartels, H., Agmon, I., Franceschi, F., and Yonath, A. (2000) Structure of functionally activated small ribosomal subunit at 3.3 angstroms resolution. Cell 102, 615–623. 8. Wimberly, B. T., Brodersen, D. E., Clemons, W. M., Jr., Morgan-Warren, R. J., Carter, A. P., Vonrhein, C., Hartsch, T., and Ramakrishnan, V. (2000) Structure of the 30S ribosomal subunit. Nature 407, 327–339. 9. Ule, J. (2008) Ribonucleoprotein complexes in neurologic diseases. Current Opinion in Neurobiology 18, 516–523. 10. Whittaker, J. W. (2007) Selective isotopic labeling of recombinant proteins using amino acid auxotroph strains. Methods. Mol. Biol. 389, 175–188. 11. Cowburn, D., Shekhtman, A., Xu, R., Ottesen, J. J., and Muir, T. W. (2004) Segmental isotopic labeling for structural biological applications of NMR. Methods Mol. Biol. 278, 47–56. 12. Liu, D., Xu, R., and Cowburn, D. (2009) Segmental isotopic labeling of proteins for nuclear magnetic resonance. Methods Enzymol. 462, 151–175. 13. Lu, K., Miyazaki, Y., and Summers, M. F. (2010) Isotope labeling strategies for NMR studies of RNA. J. Biomol. NMR 46, 113–125. 14. Nelissen, F. H., van Gammeren, A. J., Tessari, M., Girard, F. C., Heus, H. A., and Wijmenga, S. S.

(2008) Multiple segmental and selective isotope labeling of large RNA for NMR structural studies. Nucl. Acids Res. 36, e89. 15. Peterson, R. D., Theimer, C. A., Wu, H., and Feigon, J. (2004) New applications of 2D filtered/edited NOESY for assignment and structure elucidation of RNA and RNA–protein complexes. J. Biomol. NMR 28, 59–67. 16. Milligan, J. F., Groebe, D. R., Witherell, G. W., and Uhlenbeck, O. C. (1987) Oligoribonucleotide synthesis using T7 RNA polymerase and synthetic DNA templates. Nucl. Acids Res. 15, 8783–8798. 17. Cazenave, C., and Uhlenbeck, O. C. (1994) RNA template-directed RNA synthesis by T7 RNA polymerase. Proc. Natl. Acad. Sci. U.S.A. 91, 6972–6976. 18. Pleiss, J. A., Derrick, M. L., and Uhlenbeck, O. C. (1998) T7 RNA polymerase produces 5¢ end heterogeneity during in vitro transcription from certain templates. RNA 4, 1313–1317. 19. Triana-Alonso, F. J., Dabrowski, M., Wadzack, J., and Nierhaus, K. H. (1995) Self-coded 3¢-extension of run-off transcripts produces aberrant products during in vitro transcription with T7 RNA polymerase. J. Biol. Chem. 270, 6298–6307. 20. Kao, C., Rudisser, S., and Zheng, M. (2001) A simple and efficient method to transcribe RNAs with reduced 3¢ heterogeneity. Methods 23, 201–205. 21. Kao, C., Zheng, M., and Rudisser, S. (1999) A simple and efficient method to reduce nontemplated nucleotide addition at the 3 terminus of RNAs transcribed by T7 RNA polymerase. RNA 5, 1268–1272. 22. Nacheva, G. A., and Berzal-Herranz, A. (2003) Preventing nondesired RNA-primed RNA extension catalyzed by T7 RNA polymerase. Eur. J. Biochem./FEBS 270, 1458–1465. 23. Cunningham, P. R., and Ofengand, J. (1990) Use of inorganic pyrophosphatase to improve the yield of in vitro transcription reactions catalyzed by T7 RNA polymerase. BioTechniques 9, 713–714. 24. Gardner, K. H., and Kay, L. E. (1998) The use of 2H, 13C, 15N multidimensional NMR to study the structure and dynamics of proteins. Ann. Rev. Biophys. Biomol. Struct. 27, 357–406. 25. Marley, J., Lu, M., and Bracken, C. (2001) A method for efficient isotopic labeling of recombinant proteins. J. Biomol. NMR 20, 71–75.

218

C.A. Theimer et al.

26. Wu, H., Finger, L. D., and Feigon, J. (2005) Structure determination of protein/RNA complexes by NMR. Methods Enzymol. 394, 525–545. 27. Marenchino, M., Armbruster, D. W., and Hennig, M. (2009) Rapid and efficient purification of RNA-binding proteins: application to

HIV-1 Rev. Protein Expression and Purification 63, 112–119. 28. Khanna, M., Wu, H., Johansson, C., CaizerguesFerrer, M., and Feigon, J. (2006) Structural study of the H/ACA snoRNP components Nop10p and the 3¢ hairpin of U65 snoRNA. RNA 12, 40–52.

Chapter 13 Preparation and Optimization of Protein–DNA Complexes Suitable for Detailed NMR Studies My D. Sam and Robert T. Clubb Abstract This chapter describes the methods to form and optimize samples of protein–DNA complexes that are suitable for detailed structure and dynamics studies by NMR spectroscopy. Key words: Protein–DNA complex, NMR, Structure, Intermolecular NOEs

1. Introduction Interactions between proteins and DNA molecules play an essential role in a wide range of important biological processes, including gene expression and genomic replication, recombination, and repair. Of particular interest are site-specific DNA-binding transcription factors, which regulate gene expression by recognizing specific nucleotide sequences. These proteins locate, and bind to, the correct site within the genome despite the presence of a vast number of competitor sites with similar geometries and electrostatic surfaces. Over the past several decades, X-ray crystallography has been used extensively to determine a large number of high-resolution structures of protein–DNA complexes (1, 2). This work has provided a wealth of detailed stereochemical information about binding site recognition, which typically is achieved through complementary hydrogen-bonding and van der Waals interactions that can be maximized by protein folding and/or DNA distortions (3, 4). However, crystallography provides only a static view of a protein– DNA complex, and thus little insight into the conformational dynamics that underpin macromolecular recognition (5).

Alexander Shekhtman and David S. Burz (eds.), Protein NMR Techniques, Methods in Molecular Biology, vol. 831, DOI 10.1007/978-1-61779-480-3_13, © Springer Science+Business Media, LLC 2012

219

220

M.D. Sam and R.T. Clubb

NMR spectroscopy is a powerful tool that can be used to investigate protein–DNA recognition in the solution state. When the spectra obtained are of good quality, NMR can be used to elucidate high-resolution structures and atomic-level conformational dynamics (6, 7). NMR can also be applied to investigate other key aspects of recognition, including the basis of nonspecific binding (8), hydration lifetimes (9), on/off rates of binding, and the process by which a protein locates its binding site (10). Even when the quality of the NMR spectra are poor due to resonance line broadening, structural models of a protein–DNA complex can be generated using chemical shift mapping techniques. In recent years, the size and complexity of protein–DNA complexes amenable for NMR studies have increased due to several methodological advances, such as selective isotopic labeling, residual dipolar coupling measurements (11), paramagnetic relaxation enhancement methods (12), and transverse relaxation-optimized experiments, that exploit ultrahigh magnetic field strengths (13, 14). However, one of the largest obstacles to successfully studying a protein–DNA complex by NMR is the preparation of sufficiently stable, concentrated, and homogeneous samples of the complex. For a protein–DNA complex to be suitable for detailed NMR studies, it typically must satisfy several criteria. The components and nature of the interaction should be well-defined. In particular, biochemical experiments should have been performed to clearly delineate the specific nucleotide sequence recognized by the protein, as well as the stoichiometry and affinity of the resulting complex. The same molecular weight limitations that hinder NMR studies of other macromolecules apply, so the final size of the complex is also an important consideration. Ideally, the complex should have a dissociation constant (Kd) in the submicromolar range, thus making it more likely that it will be in the slow-exchange regime on the chemical shift timescale. However, weaker affinity complexes that are in fast exchange have also been successfully studied. To optimize the production and spectral qualities of a complex, large quantities of DNA and isotopically labeled protein are also needed to enable different preparative procedures and conditions to be tested. Pilot studies typically make use of purified 15N-enriched protein and a range of DNA species that differ in their length and sequence. In our experience, the greatest chance of success occurs when the protein is soluble in its DNA-free state and its 1H–15N HSQC spectrum is well-resolved. However, DNA is highly soluble and as a result the aggregation behavior of the protein may be greatly reduced upon complex formation. To create a stable complex suitable for NMR studies, the DNA molecule must have the appropriate nucleotide sequence and length to form productive contacts with the protein. For sequencespecific DNA binding proteins, this information can be obtained from previously reported biochemical studies, which should define

13

Preparation and Optimization of Protein–DNA…

221

the specificity, affinity, and stoichiometry of the complex. If the protein is known to bind to several DNA sites, then an alignment of their nucleotide sequences may reveal conserved positions essential for binding. This knowledge is helpful later in optimizing the spectra of the protein–DNA complex, since it identifies nucleotides within the DNA molecule that can presumably be altered without affecting stability. To reduce spectral overlap, the minimal DNA sequence with good binding affinity for the protein should be used to make the complex. If the structure of the protein in the DNA-free state is known, a model of it docked to B-form DNA should be constructed. This may help to determine the minimum DNA length that can be used, and whether additional nucleotides outside of the known binding site are required to form nonspecific stabilizing contacts. In practice, DNA molecules studied in our group almost always contain a G:C base pair (bp) at each end, which limits fraying by increasing the melting temperature. Minor changes in the DNA and protein sequences can dramatically affect the NMR spectrum of a protein–DNA complex and are therefore parameters that can be optimized. A common mistake is to choose a DNA fragment that is too long with unnecessary base pairs at either its 3¢ or 5¢ end. This can be problematic as longer DNA fragments can contain weaker, “cryptic” binding sites for the protein that become occupied at the high protein concentrations present in the NMR sample (typically > 0.5 mM). For example, a protein that forms numerous interactions with an A-T sequence located at the center of the primary site might also bind to a secondary A-T dinucleotide sequence present elsewhere in a longer DNA fragment. If this occurs, the multiple binding modes of the protein cause resonance line broadening. Modeling studies and a comparison of the DNA-binding sites can be used to identify potential “cryptic” sites, which can then be eliminated by altering the nucleotide sequence of the DNA molecule. A nucleotide sequence comparison can also identify dsDNA molecules that have NMR spectra that can more readily be assigned. For example, it may be preferable to maximize the number of thymine bases in the sequence, as its methyl groups are good anchor points in the assignment process. The length and sequence of the protein can also be adjusted to improve spectral quality. Typically, this involves deleting unstructured amino acids at the polypeptide termini to reduce spectral overlap. However, even subtle single amino acid changes can have a dramatic impact on spectral quality. For example, in our studies of an ARID-DNA complex, a single phenylalanine-to-leucine mutation was found to dramatically reduce line broadening, salvaging a protein–DNA complex that was originally ill suited for structural analysis by NMR (15). This biochemical approach is not a general method, but may prove useful in the spectral optimization of other protein complexes that suffer from interfacial line broadening caused by dynamic changes in proximal aromatic rings.

222

M.D. Sam and R.T. Clubb

Fig. 1. Flowchart showing the procedures used to form and optimize protein–DNA complexes for NMR studies.

In this chapter, we outline the approaches we typically use to form protein–DNA complexes suitable for NMR studies. The overall procedure for this protocol is outlined in Fig. 1.

2. Materials The exact reagents used to form protein–DNA complexes suitable for high-resolution solution-state NMR studies vary depending upon the specific system that is being studied. In this section, the materials used to produce the Integrase(Int)-DNA complex are described (16). 2.1. Binding Affinity Measurements

1. 2× binding buffer: 40 mM Tris–HCl, pH 7.5, 40 mM NaCl, 40 mM KCl, 10% (w/v) glycerol, 2 mM EDTA, and 2 mM DTT. 2.

32

P-labeled DNA: Labeled at its 5¢-termini with T4 polynucleotide kinase and g-32P-ATP.

3. PhosphorImager (Molecular Dynamics Inc.): To quantify radioactivity in the gels. 4. 1× TBE: For 1 L of 10× TBE, dissolve 108 g of Tris base, 55 g of boric acid, and 7.4 g of disodium EDTA salt. Dilute tenfold to get 1× TBE. 5. Bovine serum albumin (BSA): Stock concentration 1 mg/mL in H2O. 6. Poly dI/dC: Stock concentration 0.5 mg/mL.

13

Preparation and Optimization of Protein–DNA…

223

7. Protein stock solution: 20 mM (4× of the highest protein concentration used in the binding assay). For protein–DNA complexes with nanomolar affinity, a typical titration range is as follows (nM): 5,000, 1,000, 200, 100, 50, 25, 5, 1, 0.5, and 0. Protein should be dissolved in a solution that is most stable for that particular protein. This protein buffer is diluted in the binding assay and replaced with binding buffer solution. 8. 6× DNA loading dyes: 0.03% (w/v) xylene cyanol FF and 0.03% (w/v) bromphenol blue in 10 mM Tris, pH 8, and 30% (w/v) glycerol. 9. 1 M Tris, pH 8: Dissolve 121.1 g of Trizma (MW = 121.1 g/mol) in 800 mL of H2O, titrate with concentrated HCl until pH 8 is achieved, and then bring final volume to 1 L. 2.2. Preparation of Purified SingleStranded DNA

1. DNA oligonucleotides: 1 mmol scale synthesis (Integrated DNA Technologies, IDT) (see Note 1). 2. 17% acrylamide–urea gel: For 400 mL total volume, mix 168.2 g of urea, 170 mL of 40% acrylamide (37.5:1), 40 mL of 10× TBE (Subheading 2.1), 60 mL of H2O, 1 mL of 10% (w/v) APS, and 100 mL of TEMED. 3. DNA loading buffer: 7 M urea, 50 mM Tris, pH 8.0 (using 1 M Tris, pH 8; Subheading 2.1), 5 mM EDTA (using 0.5 M EDTA, pH 8), and 10% (w/v) glycerol. 4. 0.5 M EDTA, pH 8: Add 186.1 g of EDTA disodium salt and ~20 g of NaOH pellets to 700 mL of H2O (EDTA dissolves as it approaches pH 8), and then bring the solution to 1 L once pH 8 is achieved. 5. 6x DNA loading dyes (see Subheading 2.1). 6. 1× TBE (see Subheading 2.1). 7. FLEX TLC plates. 8. Electroelution chamber. 9. Dialysis buffer: 50 mM Tris–HCl, pH 7.5, 200 mM NaCl, and 2 mM EDTA (using 0.5 M EDTA, pH 8).

2.3. Preparation of Duplex DNA for NMR Studies

1. Annealing buffer: Same as dialysis buffer (Subheading 2.2).

2.4. Preparation of the Protein–DNA Complex

1. High-salt protein buffer: 50 mM Hepes, pH 7.0 (using 1 M Hepes, pH 7), 500 mM NaCl, and 1 mM DTT.

2. D2O.

2. 1 M Hepes, pH 7: Stock solution is adjusted to pH with NaOH. 3. High-salt DNA buffer: 50 mM Tris, pH 7.5 (using 1 M Tris, pH 7.5, see Subheading 2.1, except adjust pH to 7.5), 500 mM NaCl, and 0.1 mM EDTA.

224

M.D. Sam and R.T. Clubb

4. Low-salt buffer: 25 mM Hepes, pH 7.0 (using 1 M Hepes, pH 7), 15 mM NaCl, 2 mM DTT, 7% D2O, and 0.01% NaN3. 5. Centricon YM-3 Bioseparations).

centrifugal

filter

device

(Amicon

6. ~50 mM protein solution. 7. ~50 mM dsDNA solution.

3. Methods Our lab has solved the structures of six protein–DNA complexes. Four have been determined by NMR spectroscopy and two have been determined by X-ray crystallography (17–22). Below, we describe the procedures we generally use to form complexes between a sequence-specific binding protein and a duplex-DNA molecule. Unless otherwise stated, the procedures described below are used to produce samples of the complex between the Int protein and its cognate DNA site (17–22). Four procedures are presented: (1) an electrophoretic mobility shift assay (EMSA) for affinity and specificity measurements, (2) methods to purify singlestranded DNA (ssDNA), (3) methods to prepare duplex DNA (dsDNA), and (4) the procedures used to assess the spectral quality of a protein–DNA complex to determine if additional NMR studies are warranted. 3.1. Binding Affinity Measurements

Biochemical assays should be available to rapidly estimate the affinity of wild-type and mutant proteins for different DNA molecules. A variety of methods can be employed to measure binding, such as the EMSA, isothermal titration calorimetry (ITC), fluorescence anisotropy, surface plasmon resonance (SPR; e.g., Biacore), and fluorescence quenching (if the protein contains an appropriately positioned tryptophan). However, we favor the EMSA because it is robust and simple to perform (23). This procedure has been described in detail previously (24) and is outlined below. 1. Mix the following components: 12 mL of 2× binding buffer, 3 mL of 1 mg/mL BSA, 2 mL of 0.5 mg/mL Poly dI/dC (competitor DNA) for a total of 17 mL. 2. Add to this mixture the protein stock solution and the appropriate amount of H2O to achieve a final volume of 23 mL (see Subheading 2.1 and Note 2). 3. Incubate on ice for 20 min. 4. Add 1 mL of 32P-labeled DNA probe (~4,000 cpm/mL). 5. Incubate on ice for 20 min.

13

Preparation and Optimization of Protein–DNA…

225

6. Prepare an 8% polyacrylamide gel and pre-electrophorese the gel by running it for 30–60 min at 10 V/cm in 1× TBE at ambient temperature or 4°C (see Note 3). 7. Load the reaction mixtures onto an 8% polyacrylamide/TBE gel at 4°C (see Note 4). Load 6× DNA loading dyes in a separate lane as a reference to track the migration of the free DNA. The gel run time should be optimized for each specific system so as to resolve the species of interest and to be as short as possible. A typical gel run time is ~1.5 h at constant voltage (10 V/cm), but the voltage should be reduced if the gel becomes warm during electrophoresis. 8. Quantify the amount of free and bound DNA in each lane by using a PhosphorImager system or the equivalent. 9. Determine the dissociation constant by fitting to the following equation: θ = [L ]/ ([L ]+ K d ), where q, [L], and Kd are the fraction of DNA bound, the total protein concentration in the reaction, and the dissociation constant, respectively. q is equal to the counts present in the shifted band divided by the total counts for the DNA (free plus shifted bands). 3.2. Preparation of Purified SingleStranded DNA

In this section, we discuss how to purify large quantities of commercially available ssDNA oligonucleotides for NMR studies. We initially purchase the ssDNA in a crude form at a cost of ~$2 per nucleotide for 1 mmole of material. ssDNA with lengths less than 20 base pairs is purified on 20% acrylamide–urea gels and with lengths longer than 20 bp is purified on 17% gels. The ssDNA is then eluted from the gel and dialyzed into native buffer for further use. The procedure we use generally yields ssDNA that is >98% pure for molecules up to 40 bp in length. For ssDNA shorter than 15 bp, more conventional approaches are sufficient to produce ssDNA suitable for NMR studies (see Note 5). 1. Prepare one 17% acrylamide/urea gel with a single lane (see Note 6). 2. Dissolve the DNA oligonucleotide (1 mmol or 3–6 mg) in 2 mL of DNA loading buffer. 3. Load 20 mL of 6× DNA loading dyes on the right and left edge of the gel within the single lane. The migration of the xylene cyanol FF and bromophenol blue dyes along the gel gives an estimate of how far the DNA has migrated (see Note 7). 4. Run the gel in 1× TBE. It typically takes ~10–13 h for ssDNA (depending on the length) to migrate ~3/4 of the entire gel length. For a single gel running at a constant 50 W, the voltage is ~600–700 V (65–80 mA) (see Note 8). 5. Transfer the gel onto saran wrap by removing the top gel plate and placing a layer of saran wrap directly on top of the gel.

226

M.D. Sam and R.T. Clubb

Invert the gel onto FLEX TLC plates with the saran wrap on top of the TLC plates. Remove the remaining gel plate and in a darkroom, use a handheld UV lamp (254 nm) to locate the ssDNA. Quickly excise the DNA using a razor blade. Cut the excised gel containing the desired ssDNA into ½-in. pieces to ensure efficient DNA electroelution. The purpose of the FLEX TLC plate is to enhance the DNA signals during exposure to UV lights. 6. Assemble an Elutrap™ electroelution device (Whatman). Membranes for trapping DNA are BT1 (14 nucleotide cutoff) and BT2 (cellulose acetate membrane). Alternatively, BT1 can be replaced with a low-molecular-weight cutoff dialysis membrane. 7. Elute the DNA in 1× TBE. Run the Elutrap at 150 V for 8–10 h (remove eluted DNA from the Elutrap two to three times during this period). 8. Dialyze extensively with dialysis buffer to remove all denaturing reagents from the ssDNA. 9. Determine the DNA concentration from a UV absorbance reading at 260 nm (A260). The extinction coefficient may be calculated online using a program provided by IDT (http:// www.idtdna.com/analyzer/Applications/OligoAnalyzer/). 3.3. Preparation of Duplex DNA for NMR Studies

Complementary purified ssDNA molecules are annealed to produce the appropriate DNA duplex. NMR and/or chromatographic approaches are then used to verify duplex formation prior to forming the protein–DNA complex. The steps used to generate dsDNA are outlined below. 1. Dissolve complementary ssDNA molecules in annealing buffer to a final concentration of ~100 mM. The samples should be free of urea and EDTA. 2. Heat the sample to ~95–100°C for 10 min and slowly cool to ambient temperature in the heat block. A water bath may be used in place of a heating block for larger annealing volumes. 3. Perform NMR experiments to ensure that the sample has properly annealed and that no excess ssDNA is present. Add an appropriate amount of D2O to maintain the field lock, and acquire an 1H 2D TOCSY spectrum (mixing time ~40 ms). Acquire spectra of the ssDNA components of the duplex as well. Compare the three spectra; signals from the ssDNA spectra appearing in the duplex spectrum indicate ssDNA excess. Typically, the H5-H6 cross peaks of the cytosine bases are used to discriminate between the single-stranded and doublestranded forms of DNA (see Note 9).

3.4. Preparation of the Protein–DNA Complex

Our group follows a conservative approach when forming protein– DNA complexes to minimize sample loss due to precipitation (17–22) (see Note 10). In this procedure, dilute concentrations of

13

Preparation and Optimization of Protein–DNA…

227

the components are mixed in the presence of high salt, followed by concentration and removal of the salt to form the final NMR sample. Typically, the starting concentration of the dsDNA is ~50–100 mM and it is dissolved in at least 150 mM NaCl, near physiological pH (see Note 11). Pilot studies should be performed prior to embarking on a large-scale production of the complex as we have found that the order of addition of the protein and DNA components can have a dramatic impact on the results obtained. To test for precipitation of the sample upon component mixing, slowly titrate the protein into a ~20 mL sample of the DNA (the final volume of the fully formed complex should be ~40 mL). The concentration of the components is as described above and enough protein should be added to generate a 1:1 complex. The reverse titration in which dsDNA is titrated into a sample of the protein should also be performed. In both cases, carefully observe if any precipitation occurs during the mixing procedure and, more importantly, if any precipitation remains after the titration is complete. A light microscope can provide an easy way to assay for the presence of precipitation. As already mentioned, the order of addition can be critical. For example, in our studies of the Excisionase(Xis)-DNA complex, the addition of dsDNA to a solution of Xis resulted in irreversible precipitation, whereas the reverse titration, titrating Xis into a solution of dsDNA, yielded a soluble complex after mixing (20). To obtain an NMR sample once the protocol for complex formation has been optimized, we typically mix the DNA and protein components such that the final volume of the complex is ~10 mL (~25 mM of the complex). The salt and unwanted buffer components are then removed by dialysis or using a centrifugal filter unit. This step is important for complex stability as the presence of salt tends to destabilize the protein–DNA complex by shielding electrostatic interactions. Generally, to obtain a sample with good NMR spectral properties, several variants of the complex are studied that differ in their pH, complex concentration, and ionic strength. Initial screening is typically performed using ~200–500 mM samples of the complex, which are then concentrated further to construct the final sample once the best conditions are discovered (see Note 12). 1. Prepare ~10 mL of 50 mM Int protein solution dissolved in high-salt protein buffer. Prepare ~10 mL of 50 mM dsDNA solution dissolved in high-salt DNA buffer (see Note 11). 2. Prepare the final NMR sample by slowly titrating 10 mL of 50 mM Int into 10 mL of 50 mM dsDNA. Add ~0.5 mL of protein, mix, and repeat until the titration is complete. The specific order of addition and the conditions used should have been optimized as described above.

228

M.D. Sam and R.T. Clubb

3. Exchange the final sample into low-salt buffer using a protein concentrator (see Note 12). Concentrate the sample to a volume suitable for NMR experiments. Monitor the total amount of complex at each step of this process by measuring the A260 (see Note 13). 3.5. Assessing the Quality of the Protein– DNA Complex

1. To assess the quality of the DNA spectrum, record 1D 1H spectra using a 1331 pulse adjusted to maximally excite the imino protons (25). 2. Compare the spectra of the complex and the isolated dsDNA molecule to assess protein binding. 3. If sufficient material is present, use a similar excitation scheme to record a 2D 1H NOESY spectrum, which reveals whether the appropriate imino–imino cross peaks are present. Ascertain the quality of the protein spectrum by comparing the 1H–15N HSQC spectra of the free and DNA-bound forms of the protein. The spectrum of the complex should be well-resolved, exhibit uniform signal intensities, and differ substantially from the NMR spectrum of the DNA-free protein (Fig. 2). In favorable cases, it may be possible to observe intermolecular NOEs using protein–DNA complexes containing only 15 N-labeled protein. Depending on the structure of the complex, intermolecular NOE cross peaks are sometimes observed in the 2D 1H NOESY spectrum between the imino protons of

Fig. 2. The 1H–15N HSQC spectrum of the Int-DNA complex, which exhibits good dispersion and uniform line widths. Data for this complex were suitable to determine a highresolution NMR structure (17–22).

13

Preparation and Optimization of Protein–DNA…

229

the DNA and protein protons that resonate upfield of 1.2 ppm. This is because the most upfield resonances in the 1H spectrum of DNA are the thymine H5 methyl groups (1.2 and 1.6 ppm). Intermolecular NOEs between the protein amide and DNA imino protons can sometimes also be seen in the 3D 15N-edited NOESY spectrum of the 15N-labeled complex. However, a full assessment of whether a complex is suitable for structure determination by NMR requires the acquisition of the appropriate 2D and 3D edited and filtered NOESY experiments using samples in which the protein is labeled with 13C and 15N.

4. Notes 1. A variety of companies sell pure or partially purified singlestranded oligonucleotides that are synthesized using phosphoramidite chemistry (e.g., IDT, Invitrogen, Sigma, and Applied Biosystems). To save money, we typically obtain standard, desalted, unpurified oligonucleotides, which are then further purified in-house. Olignonucleotides greater than 60 nt in length can be produced via enzymatic reactions, which may also be applied to produce DNA molecules enriched with 13C and 15N (26, 27). 2. In this procedure, binding isotherms are generated by varying the protein concentration (0, 0.5, 1, 5, 25, 50, 100, 200, 1,000, 5,000 nM protein). 3. Alternatively, Tris–acetate–EDTA (TAE) may be used for the gel casting mixture and running buffer. 4. A 5–15% gel could be used depending on the respective sizes of the individual components and formed complex. 5. A classical and more straightforward method to purify ssDNA from a crude synthesis is to use an HPLC reverse-phase column (C4 to C18; porous hydrocarbon silica gel) with 0.1 M triethylammonium acetate (TEAA; mobile phase A) and ace-tonitrile (mobile phase B). A complete purification protocol and a column selection guide are described in Current Protocols Nucleic Acid Chemistry by Andrus et al. (28). A Mono Q column (GE, Mono Q HR5/5) on an FPLC can be used to purify ssDNA under denaturing conditions. Buffers for this purification scheme are as follows: (a) Buffer A: 50 mM Tris, pH 7.5, 6 M urea, and (b) Buffer B: 50 mM Tris, pH 7.5, 6 M urea, 1.5 M NaCl. No more than 1.5 mmol of ssDNA should be loaded onto this column for good separation. A 25 mL gradient (flow rate = 1 mL/min) from 10 to 30% buffer B should be sufficient to ensure good separation of ssDNA from impurities. It should be noted that all buffers and resuspended DNA

230

M.D. Sam and R.T. Clubb

should be filtered through a 0.2-mm filter before applying to the column. 6. If the DNA is longer than ~20 nucleotides, a 20% acrylamide gel should be used instead. 7. On a 10% gel, xylene cyanol migrates equivalent to a 55-nucleotide ssDNA molecule and bromophenol blue migrates equivalent to a 12-nucleotide ssDNA molecule. On a 20% gel, xylene cyanol and bromophenol blue migrate equivalent to 28- and 8-nucleotide ssDNA molecules, respectively. 8. Power settings for a single 20% acrylamide/7 M urea gel should be set to 30 W and maximum voltage and current. For a single gel running at a constant 30 W, the voltage is ~600–650 V and the current is 45–50 mA. 9. An alternative approach to ensure that solutions of dsDNA do not contain excess ssDNA is to provide a 10% excess of one of the DNA strands in the annealing reaction. After annealing, a Mono Q column is then used to separate the dsDNA molecule from excess ssDNA under native conditions. The following buffers can be used: Buffer A: 50 mM Tris, pH 7.5, and 1 mM EDTA; Buffer B: 50 mM Tris, pH 7.5, 1 mM EDTA, and 1.5 M NaCl. For good separation of the two DNA species, no more than 2 mg of DNA should be loaded onto a 1 mL Mono Q column. ssDNA typically elutes around 25% buffer B (0.375 M NaCl) and dsDNA typically elutes at ~35% buffer B (0.525 M NaCl). A 25 mL gradient from 10 to 45% NaCl running at 1 mL/min works well for oligos between 10 and 30 bp. 10. A variety of approaches can be used to successfully make protein–DNA complexes for NMR studies. A strategy commonly used in the literature is to titrate a ~1 mM 15N-labeled sample of the protein with a concentrated stock solution of the DNA molecule. 1H–15N HSQC spectra are recorded at various points during the titration until the desired stoichiometry is reached. Although the titration method is simple, mixing concentrated protein and DNA samples can cause the resulting complex to precipitate. 11. These conditions prevent a shift in the equilibrium from dsDNA to ssDNA that occurs at low-salt concentrations. A similar protein concentration is used when forming the complex; however, the salt and pH in the protein solution can vary and are chosen to maximize the stability and solubility of the free protein. 12. Frequently, the components of the complex are precious and therefore methods that minimize sample loss during concentration are desired. One trick to concentrate small-volume samples (< 2 mL) is to partially evaporate the complex. In this procedure, a weak flow of nitrogen gas is blown over the

13

Preparation and Optimization of Protein–DNA…

231

sample while it rests in the NMR tube. This can be accomplished by passing the gas through drawn-out pipette that is in turn inserted into the NMR tube. The buffer components are also concentrated during this process, and therefore the initial buffer conditions must be chosen accordingly. 13. A rough estimate of the concentration of the complex can be obtained by measuring its A260 immediately after mixing the components. The amount of the material present at this point in the preparation procedure is known. Therefore, it is possible to estimate the extinction coefficient of the complex by determining the optical absorbance of the complex at 260 and 280 nm. Once estimated, the extinction coefficient enables the concentration of the complex to be readily determined as it is concentrated to a volume suitable for NMR. It also enables the yield of the concentration procedure to be determined as the total amount of complex before and after concentrating can be determined.

Acknowledgments We thank Dr. Evgeny Fadeev for making Fig. 2. This work was supported by a grant from the National Institutes of Health to R.T.C. (R01 AI52217). References 1. Pabo, C. O., and Sauer, R. T. (1992) Transcription Factors-Structural Families and Principles of DNA Recognition. Annu. Rev. Biochem. 61, 1053–95. 2. Garvie, C. W., and Wolberger, C. (2001) Recognition of specific DNA sequences. Mol. Cell. 8, 937–946. 3. Nadassy, K., Wodak, S. J., and Janin, J. (1999) Structural features of protein-nucleic acid recognition sites. Biochemistry 38, 1999–2017. 4. Jen-Jacobson, L. (1997) Protein-DNA recognition complexes: conservation of structure and binding energy in the transition state. Biopolymers 44, 153–180. 5. Boehr, D. D., Nussinov, R., and Wright, P. E. (2009) The role of dynamic conformational ensembles in biomolecular recognition. Nat. Chem. Biol. 5, 789–796. 6. Billeter, M., Qian, Y. Q., Otting, G., Muller, M., Gehring, W., and Wuthrich, K. (1993) Determination of the nuclear magnetic resonance

7.

8.

9.

10.

solution structure of an Antennapedia homeodomain-DNA complex. J. Mol. Biol. 234, 1084–1093. Omichinski, J. G., Clore, G. M., Schaad, O., Felsenfeld, G., Trainor, C., Appella, E., Stahl, S. J., and Gronenborn, A. M. (1993) NMR structure of a specific DNA complex of Zn-containing DNA binding domain of GATA-1. Science 261, 438–446. Kalodimos, C. G., Biris, N., Bonvin, A. M., Levandoski, M. M., Guennuegues, M., Boelens, R., and Kaptein, R. (2004) Structure and flexibility adaptation in nonspecific and specific protein-DNA complexes. Science 305, 386–389. Qian, Y. Q., Otting, G., and Wuthrich, K. (1993) NMR detection of hydration water in the intermolecular interface of a protein-DNA complex. J. Am. Chem. Soc. 115, 1189–1190. Iwahara, J., and Clore, G. M. (2006) Direct observation of enhanced translocation of a

232

11.

12.