Globins and Other Nitric Oxide-Reactive Proteins, Part B (Methods in Enzymology, Volume 437)

VOLUME FOUR HUNDRED AND THIRT Y-SEVEN METHODS IN ENZYMOLOGY Globins and Other Nitric Oxide-Reactive Proteins, Part B...

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VOLUME FOUR HUNDRED AND THIRT Y-SEVEN

METHODS

IN

ENZYMOLOGY Globins and Other Nitric Oxide-Reactive Proteins, Part B

METHODS IN ENZYMOLOGY Editors-in-Chief

JOHN N. ABELSON AND MELVIN I. SIMON Division of Biology California Institute of Technology Pasadena, California Founding Editors

SIDNEY P. COLOWICK AND NATHAN O. KAPLAN

VOLUME FOUR HUNDRED AND THIRT Y-SEVEN

METHODS

IN

ENZYMOLOGY Globins and Other Nitric Oxide-Reactive Proteins, Part B EDITED BY

ROBERT K. POOLE Department of Molecular Biology and Biotechnology The University of Sheffield Sheffield, UK

AMSTERDAM • BOSTON • HEIDELBERG • LONDON NEW YORK • OXFORD • PARIS • SAN DIEGO SAN FRANCISCO • SINGAPORE • SYDNEY • TOKYO Academic Press is an imprint of Elsevier

Academic Press is an imprint of Elsevier 525 B Street, Suite 1900, San Diego, California 92101-4495, USA 84 Theobald’s Road, London WC1X 8RR, UK

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⬁

Copyright # 2008, Elsevier Inc. All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the Publisher. The appearance of the code at the bottom of the first page of a chapter in this book indicates the Publisher’s consent that copies of the chapter may be made for personal or internal use of specific clients. This consent is given on the condition, however, that the copier pay the stated per copy fee through the Copyright Clearance Center, Inc. (www.copyright.com), for copying beyond that permitted by Sections 107 or 108 of the U.S. Copyright Law. This consent does not extend to other kinds of copying, such as copying for general distribution, for advertising or promotional purposes, for creating new collective works, or for resale. Copy fees for pre-2008 chapters are as shown on the title pages. If no fee code appears on the title page, the copy fee is the same as for current chapters. 0076-6879/2008 $35.00 Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone: (þ44) 1865 843830, fax: (þ44) 1865 853333, E-mail: permissions@elsevier. com. You may also complete your request on-line via the Elsevier homepage (http://elsevier.com), by selecting ‘‘Support & Contact’’ then ‘‘Copyright and Permission’’ and then ‘‘Obtaining Permissions.’’ For information on all Elsevier Academic Press publications visit our Web site at www.books.elsevier.com ISBN: 978-0-12-374278-0 PRINTED IN THE UNITED STATES OF AMERICA 08 09 10 11 9 8 7 6 5 4 3 2 1

CONTENTS

Contributors Preface Volumes in Series

xvii xxvii xxix

Section I. Nitric Oxide-Metabolising and Detoxifying Enzymes

1

1. Structural Studies on Flavodiiron Proteins

3

Joaõ B. Vicente, Maria Armeńia Carrondo, Miguel Teixeira, and Carlos Frazaõ 1. Introduction 2. Crystallization of Flavodiiron Proteins 3. Diffraction Data Collection, Structure Determination, and Refinement 4. Overall Description of Structures 5. Conclusion References

2. Biochemical, Spectroscopic, and Thermodynamic Properties of Flavodiiron Proteins

4 4 7 8 16 17

21

Joaõ B. Vicente, Marta C. Justino, Vera L. Gonçalves, Lı´gia M. Saraiva, and Miguel Teixeira 1. Introduction 2. Cloning of Genes Encoding Flavodiiron Proteins and Their Truncated Domains 3. Production and Purification of Recombinant Flavodiiron Proteins 4. Biochemical Characterization of Flavodiiron Proteins 5. Spectroscopic Properties 6. Redox Properties 7. Conclusions Acknowledgments References

22 24 25 26 29 32 37 42 42

v

vi

Contents

3. Kinetic Characterization of the Escherichia coli Nitric Oxide Reductase Flavorubredoxin

47

Joaõ B. Vicente, Francesca M. Scandurra, Elena Forte, Maurizio Brunori, Paolo Sarti, Miguel Teixeira, and Alessandro Giuffre` 1. Introduction 2. Amperometric Measurements 3. Spectroscopic Measurements 4. Conclusions Acknowledgments References

4. Escherichia coli Cytochrome c Nitrite Reductase NrfA

48 49 51 61 61 61

63

Thomas A. Clarke, Paul C. Mills, Susie R. Poock, Julea N. Butt, Myles R. Cheesman, Jeffrey A. Cole, Jay C. D. Hinton, Andrew M. Hemmings, ¨derberg, Stephen Spiro, Jessica Van Gemma Kemp, Christopher A. G. So Wonderen, and David J. Richardson 1. Introduction 2. Measurement of Cytochrome c Nitrite Reductase-Dependent Consumption of Nitric Oxide in Whole Cells 3. Growth of E. coli Optimized for Cytochrome c Nitrite Reductase Production for Use in Enzyme Purification 4. Purification of Cytochrome c Nitrite Reductase 5. Assaying the Cytochrome c Nitrite Reductase 6. Crystallization of E. coli Cytochrome c Nitrite Reductase 7. Concluding Remarks Acknowledgments References

5. The Respiratory Nitric Oxide Reductase (NorBC) from Paracoccus denitrificans

64 66 66 68 69 73 74 76 76

79

Sarah J. Field, Faye H. Thorndycroft, Andrey D. Matorin, David J. Richardson, and Nicholas J. Watmough 1. Introduction 2. Purification of Native NorBC from Paracoccus denitrificans 3. Purification of Recombinant NorBC 4. Amperometric Assays of NO Consumption 5. Pseudoazurin as an Electron Donor in Assays of NorBC 6. Preparation of NOR for Spectroscopic Investigation 7. Electron Paramagnetic Resonance Spectroscopy 8. Concluding Remarks Acknowledgments References

80 82 85 86 88 91 96 98 99 99

Contents

6. Redox-Controlled Dinitrosyl Formation at the Diiron-Oxo Center of NorA

vii

103

Rainer Cramm and Katja Strube 1. Introduction 2. Genetic Context and Expression of the NorA Gene in R. eutropha 3. Purification of NorA 4. Disulfide Bridges in NorA 5. Iron Analysis and Preparation of Apo-NorA 6. Interconversion of Redox Forms of NorA 7. Generation of NorA-DNIC In Vitro 8. Preparation of NorA-DNIC Formed In Vivo 9. Quantification of NO from NorA-DNIC 10. Outlook References

7. Purification and Functional Analysis of Fungal Nitric Oxide Reductase Cytochrome P450nor

104 105 106 107 108 109 111 113 113 114 114

117

Li Zhang and Hirofumi Shoun 1. Introduction 2. Screening of P450nor Activity 3. Gas Analysis 4. Purification of P450nor 5. Nitric Oxide Reductase Activity Assay 6. Protein Sequencing 7. Isolation of cDNA 8. Subcellular Fractionation of T. cutaneum 9. Site-Directed Mutagenesis 10. Expression of Recombinant Proteins 11. Purification of Recombinant Proteins 12. Titration of NAD Analogs 13. Stopped-Flow Rapid Scan Analysis 14. Other Analysis 15. Conclusion Acknowledgments References

8. A Quantitative Approach to Nitric Oxide Inhibition of Terminal Oxidases of the Respiratory Chain

118 119 119 120 121 122 123 124 125 126 127 128 130 131 131 131 131

135

Maria G. Mason, Rebecca S. Holladay, Peter Nicholls, Mark Shepherd, and Chris E. Cooper 1. Introduction 2. Evaluation of Current Techniques for Measuring pNO, pO2, and KM (O2)

136 137

viii

Contents

3. 4. 5. 6.

Nitric Oxide Donor Compounds Nitric Oxide Kinetics Oxygen Kinetics Optical Detection of Enzyme Intermediates in the Presence of Oxygen and NO Appendices Acknowledgments References

Section II. Sensor Proteins 9. Cloning, Expression, and Purification of the N-terminal Heme-Binding Domain of Globin-Coupled Sensors

138 139 149 151 153 156 156

161 163

Jennifer A. Saito, Tracey Allen K. Freitas, and Maqsudul Alam 1. Introduction 2. Bioinformatic Search of Globin-Coupled Sensors 3. Functional Analysis of Globin-Coupled Sensors Acknowledgments References

10. Oxygen-Sensing Histidine-Protein Kinases: Assays of Ligand Binding and Turnover of Response-Regulator Substrates

164 164 166 171 171

173

Marie-Alda Gilles-Gonzalez, Gonzalo Gonzalez, Eduardo Henrique Silva Sousa, and Jason Tuckerman 1. Introduction 2. Assays Acknowledgments References

11. Reactions of Nitric Oxide and Oxygen with the Regulator of Fumarate and Nitrate Reduction, a Global Transcriptional Regulator, during Anaerobic Growth of Escherichia coli

174 175 187 187

191

Jason C. Crack, Nick E. Le Brun, Andrew J. Thomson, Jeffrey Green, and Adrian J. Jervis 1. Introduction 2. Production of 4Fe-FNR Protein 3. Determination of Iron and Acid-Labile Sulfide Content of FNR 4. UV-Visible Absorbance Spectra of FNR 5. Cluster Reaction with Nitric Oxide and Oxygen 6. Purification of 2Fe-FNR 7. Detection of Other Reaction Products 8. Conclusions References

192 194 197 198 198 204 204 206 207

Contents

12. Genome-Wide Identification of Binding Sites for the Nitric Oxide-Sensitive Transcriptional Regulator NsrR

ix

211

Sam Efromovich, David Grainger, Diane Bodenmiller, and Stephen Spiro 1. Introduction 2. Strain Construction 3. Reference and Control Samples 4. Culture Conditions 5. Immunoprecipitation of DNA Targets Associated with NsrR 6. DNA Labeling, Microarray Hybridization, and Processing 7. Visualization and Analysis of DNA Microarray Data 8. A New Statistical Methodology for Treatment of Chip-on-Chip Data 9. Conclusions Acknowledgments References

13. Characterization of the Nitric Oxide-Reactive Transcriptional Activator NorR

212 214 216 217 218 219 220 222 231 231 231

235

Benoıˆt D’Autreáux, Nick Tucker, Stephen Spiro, and Ray Dixon 1. 2. 3. 4.

Introduction Measurement of NorR Activity In Vivo Measurement of Transcriptional Activation by NorR In Vitro Detection of the Ferrous-Nitrosyl Form of NorR by In Vivo Electron Paramagnetic Resonance (EPR) 5. In Vitro Reconstitution of the Iron Center in NorR 6. Measurement of NO Affinity 7. Standardization of the NO Electrode 8. Determination of NorRFe(NO) Kd 9. Conclusions Acknowledgment References

Section III. Advanced Spectroscopic Methods 14. Hemoglobins from Mycobacterium tuberculosis and Campylobacter jejuni: A Comparative Study with Resonance Raman Spectroscopy

236 237 238 240 242 243 246 247 248 248 249

253

255

Changyuan Lu, Tsuyoshi Egawa, Masahiro Mukai, Robert K. Poole, and Syun-Ru Yeh 1. Hemoglobin Superfamily: An Overview 2. Microbial Hemoglobins 3. Resonance Raman Spectroscopy: Applications in Hemeproteins

256 257 258

x

Contents

4. Structures and Functions of Microbial Hemoglobins 5. Closing Remarks Acknowledgments References

266 281 282 282

15. The Power of Using Continuous-Wave and Pulsed Electron Paramagnetic Resonance Methods for the Structure Analysis of Ferric Forms and Nitric Oxide-Ligated Ferrous Forms of Globins 287 Sabine Van Doorslaer and Filip Desmet 1. Introduction 2. Electron Paramagnetic Resonance in a Nutshell 3. EPR Studies of NO-Ligated Globins 4. EPR Studies of Ferric globins 5. Spin-Labeling Heme Proteins 6. Future Challenges and Possibilities Acknowledgments References

16. Oxygen Binding to Heme Proteins in Solution, Encapsulated in Silica Gels, and in the Crystalline State

288 289 295 301 304 305 305 306

311

Luca Ronda, Stefano Bruno, Serena Faggiano, Stefano Bettati, and Andrea Mozzarelli 1. Oxygen-Binding Curves to Heme Proteins 2. Determination of OBCs for Hemoglobin in Solution 3. Determination of K1 for Hemoglobin in Solution in the Absence of Allosteric Effectors 4. Determination of OBCs for T State Hemoglobin Gels in the Absence and Presence of Allosteric Effectors 5. Determination of OBCs for T State Hemoglobin Crystals 6. Determination of OBCs for Hemocyanin in Solution and in Silica Gels Acknowledgments References

17. Characterization of Ligand Migration Mechanisms inside Hemoglobins from the Analysis of Geminate Rebinding Kinetics

313 316 318 318 320 323 325 325

329

Stefania Abbruzzetti, Stefano Bruno, Serena Faggiano, Luca Ronda, Elena Grandi, Andrea Mozzarelli, and Cristiano Viappiani 1. 2. 3. 4.

Introduction Principles of Nanosecond Laser Flash Photolysis Basic Experimental Layouts Encapsulation of Hbs in Silica Gels

330 330 331 335

Contents

5. Enhancement of Geminate Rebinding and Advantages of Gel Encapsulation 6. Extraction of Kinetic Information Acknowledgments References

18. Ligand Dynamics in Heme Proteins Observed by Fourier Transform Infrared Spectroscopy at Cryogenic Temperatures

xi

336 337 342 342

347

Karin Nienhaus and G. Ulrich Nienhaus 1. Introduction 2. Materials 3. Fourier Transform Infrared Cryospectroscopy 4. Low-Temperature FTIR Spectroscopy on NO-Ligated Heme Proteins 5. Concluding Remarks Acknowledgments References

19. Time-Resolved X-Ray Crystallography of Heme Proteins

348 349 353 365 373 374 374

379

Vukica Sˇrajer and William E. Royer, Jr. 1. Introduction 2. Experiment 3. Data Processing and Analysis 4. A Case Study: Scapharca Dimeric Hemoglobin 5. Conclusions Acknowledgments References

20. Structural Dynamics of Myoglobin

379 381 385 388 391 393 393

397

M. Brunori, D. Bourgeois, and B. Vallone 1. Background 2. Crystallographic Studies of Myoglobin States 3. Experimental Approaches Acknowledgments References

21. Use of the Conjugate Peak Refinement Algorithm for Identification of Ligand-Binding Pathways in Globins

398 399 400 413 413

417

Stephen D. Golden and Kenneth W. Olsen 1. Introduction 2. Exploration of Oxygen-Binding Pathways in Myoglobin

418 418

xii

Contents

3. Theoretical Models 4. Potential Energy Function 5. Transition Pathways 6. Methods 7. Results 8. Conclusions References

22. Finding Gas Migration Pathways in Proteins Using Implicit Ligand Sampling

419 420 421 425 429 432 433

439

Jordi Cohen, Kenneth W. Olsen, and Klaus Schulten 1. Introduction 2. Methods 3. Example Calculation: Truncated Hemoglobin (trHb) from Paramecium caudatum 4. Discussion Acknowledgments References

440 442 446 449 455 456

23. Identification of Ligand-Binding Pathways in Truncated Hemoglobins Using Locally Enhanced Sampling Molecular Dynamics

459

Stephen D. Golden and Kenneth W. Olsen 1. Introduction 2. Molecular Dynamics 3. Locally Enhanced Sampling Molecular Dynamics 4. Methods 5. Results 6. Conclusions References

24. Nitric Oxide Reactivity with Globins as Investigated Through Computer Simulation

460 462 465 466 468 471 472

477

Marcelo A. Marti, Luciana Capece, Axel Bidon-Chanal, Alejandro Crespo, Victor Guallar, F. Javier Luque, and Dario A. Estrin 1. 2. 3. 4.

Introduction Molecular Dynamics (MD) Methods Quantum Mechanical-Molecular Mechanical Methods Illustrative Examples

478 479 485 488

Contents

5. Ligand Migration Profiles from MSMD and PELE Simulations: Exploring Ligand Entry Pathways in M. tuberculosis trHbN 6. Conclusions Acknowledgments References

25. Microbial Responses to Nitric Oxide and Nitrosative Stress: Growth, ‘‘Omic,’’ and Physiological Methods

xiii

490 494 495 495

499

Steven T. Pullan, Claire E. Monk, Lucy Lee, and Robert K. Poole 1. Introduction 2. Methods 3. Nitric Oxide, NO-Releasing Agents, and Nitrosating Agents 4. Illustrative Results from Applications of These Methods References

26. Analysis of Nitric Oxide-Dependent Antimicrobial Actions in Macrophages and Mice

500 504 512 514 516

521

Andre´s Vazquez-Torres, Tania Stevanin, Jessica Jones-Carson, Margaret Castor, Robert C. Read, and Ferric C. Fang 1. NO-Dependent Antimicrobial Actions of Murine Macrophages 2. NO-Dependent Antimicrobial Actions of Human Macrophages 3. NO-dependent Antimicrobial Actions in Laboratory Mice References

27. Measuring Nitric Oxide Metabolism in the Pathogen Neisseria meningitidis

522 528 532 536

539

Melanie J. Thomson, Tania M. Stevanin, and James W. B. Moir 1. 2. 3. 4.

Introduction Safety Aspects of Handling N. meningitidis in the Laboratory Metabolism of Neisseria sp. Experimental Approaches to Analyzing Nitrogen Metabolism Relevant to NO 5. Simultaneous Measurement of Oxygen and NO during Pure Culture of N. meningitidis 6. Measurement of NO Production/Disappearance in Tissue Culture Using Human Monocyte-Derived Macrophages References

540 541 541 544 547 555 558

xiv

Contents

28. Localization of S-Nitrosothiols and Assay of Nitric Oxide Synthase and S-Nitrosoglutathione Reductase Activity in Plants

561

Francisco J. Corpas, Alfonso Carreras, Francisco J. Esteban, Mounira Chaki, Raquel Valderrama, Luis A. Del Rıó, and Juan B. Barroso 1. Introduction 2. Determination of L-Arginine-Dependent NOS Activity by Ozone Chemiluminescence in Plant Tissues 3. Assay of GSNOR Activity 4. Localization of S-Nitrosothiols and S-Nitrosoglutathione in Plant Tissues by Confocal Laser-Scanning Microscopy (CLSM) 5. Conclusion Acknowledgments References

29. Methods for Nitric Oxide Detection during Plant–Pathogen Interactions

562 563 566 567 571 572 572

575

E. Vandelle and M. Delledonne 1. 2. 3. 4. 5. 6.

Introduction Nitric Oxide Detection by Mass Spectrometry Nitric Oxide Detection by Laser Photoacoustic Spectroscopy Nitric Oxide Detection by Chemiluminescence Nitric Oxide Detection by Hemoglobin Conversion Nitric Oxide Detection by Electron Paramagnetic Resonance (EPR) Spin Trap 7. Nitric Oxide Detection Using Diaminofluoresceins 8. Conclusion References

30. Bioimaging Techniques for Subcellular Localization of Plant Hemoglobins and Measurement of Hemoglobin-Dependent Nitric Oxide Scavenging In Planta

576 577 579 582 583 585 587 590 591

595

Kim H. Hebelstrup, Erik stergaard-Jensen, and Robert D. Hill 1. Introduction 2. Measuring Hemoglobin-Dependent NO Scavenging 3. Techniques for Determination of Subcellular Localization of Plant Hemoglobins 4. Imaging of Hemoglobin-Dependent NO Scavenging in Arabidopsis Plants 5. Engineering of GLB1-GFP/GLB2-GFP Constructs and Microscopic Analysis of A. thaliana Plants Expressing GFP-Tagged Hemoglobin References

596 596 597 598 600 603

Contents

31. Use of Recombinant Iron-Superoxide Dismutase as A Marker of Nitrative Stress

xv

605

Estı´baliz Larrainzar, Estı´baliz Urarte, In˜igo Auzmendi, Idoia Ariz, Cesar Arrese-Igor, Esther M. Gonza´lez, and Jose F. Moran 1. Introduction 2. Immunodetection of Nitrated Proteins: Metal-Mediated Tyrosine Nitration of BSA 3. Tyrosine Nitration of Purified Recombinant Vu_FeSOD Affects its Enzymatic Activity 4. Tyrosine Nitration in Vu_FeSOD can be Estimated Using Antibodies Against 3-Nitrotyrosine 5. SIN-1-Dependent Vu_FeSOD Nitration can be Detected by the Loss of Enzymatic Activity Acknowledgments References Author Index Subject Index

606 608 610 612 612 616 616 619 647

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CONTRIBUTORS

Stefania Abbruzzetti Dipartimento di Fisica, Universita` degli Studi di Parma, Parma, Italy, and NEST CNR-INFM, Pisa, Italy Maqsudul Alam Department of Microbiology, and Advanced Studies in Genomics, Proteomics, and Bioinformatics, College of Natural Sciences, University of Hawaii, Honolulu, Hawaii Cesar Arrese-Igor Departamento de Ciencias del Medio Natural, Universidad Pu´blica de Navarra, Campus de Arrosadia, E-31006 Pamplona, Navarre, Spain Idoia Ariz Instituto de Agrobiotecnologıá, Universidad Pu´blica de Navarra-CSIC-Gobierno de Navarra, Campus de Arrosadıá, E-31006 Pamplona, Navarre, Spain In˜igo Auzmendi Departamento de Ciencias del Medio Natural, Universidad Pu´blica de Navarra, Campus de Arrosadia, E-31006 Pamplona, Navarre, Spain Juan B. Barroso Grupo de Senãlizacioń Molecular y Sistemas Antioxidantes en Plantas, Unidad ´ rea de Bioquı´mica y Biologıá Molecular, Universidad Asociada al CSIC (EEZ), A de Jaeń, E-23071 Jaeń, Spain Stefano Bettati Department of Biochemistry and Molecular Biology, University of Parma, Parma, Italy Axel Bidon-Chanal Departament de Fisicoquı´mica, Facultat de Farma`cia, Universitat de Barcelona, Barcelona, Spain Diane Bodenmiller School of Biology, Georgia Institute of Technology, Atlanta, Georgia D. Bourgeois Institut de Biologie Structurale Jean-Pierre Ebel, CEA, CNRS, Universite´ Joseph Fourier and European Synchrotron Radiation Facility, Grenoble Cedex, France

xvii

xviii

Contributors

Nick E. Le Brun Centre for Metalloprotein Spectroscopy and Biology, School of Chemical Sciences and Pharmacy, University of East Anglia, Norwich, United Kingdom Stefano Bruno Department of Biochemistry and Molecular Biology, University of Parma, Parma, Italy, and Dipartimento di Biochimica e Biologia Molecolare, Universita` degli Studi di Parma, Parma, Italy M. Brunori Dipartimento di Scienze Biochimiche ‘‘A. Rossi Fanelli,’’ Universita` di Roma ‘‘La Sapienza,’’ Roma, Italy Maurizio Brunori Department of Biochemical Sciences, CNR Institute of Molecular Biology and Pathology and Istituto Pasteur–Fondazione Cenci Bolognetti Sapienza, University of Rome, Rome, Italy Julea N. Butt Centre for Metalloprotein Spectroscopy and Biology, School of Biological Sciences, and School of Chemical Sciences and Pharmacy, University of East Anglia, Norwich, United Kingdom Luciana Capece Departamento de Quı´mica Inorgańica, Analı´tica y Quı´mica Fı´sica/INQUIMAECONICET, Facultad de Ciencias Exactas y Naturales, Universidad de Buenos Aires, Ciudad Universitaria, Buenos Aires, Argentina Alfonso Carreras Grupo de Senãlizacioń Molecular y Sistemas Antioxidantes en Plantas, Unidad ´ rea de Bioquı´mica y Biologıá Molecular, Universidad Asociada al CSIC (EEZ), A de Jaeń, E-23071 Jaeń, Spain Maria Armeńia Carrondo Instituto de Tecnologia Quı´mica e Biolo´gica, Universidade Nova de Lisboa, Oeiras, Portugal Margaret Castor University of Washington School of Medicine, Seattle, Washington Mounira Chaki Grupo de Senãlizacioń Molecular y Sistemas Antioxidantes en Plantas, Unidad ´ rea de Bioquı´mica y Biologıá Molecular, Universidad Asociada al CSIC (EEZ), A de Jaeń, E-23071 Jaeń, Spain Myles R. Cheesman Centre for Metalloprotein Spectroscopy and Biology, School of Chemical Sciences and Pharmacy, University of East Anglia, Norwich, United Kingdom

Contributors

xix

Thomas A. Clarke Centre for Metalloprotein Spectroscopy and Biology, School of Biological Sciences, University of East Anglia, Norwich, United Kingdom Jordi Cohen Beckman Institute, University of Illinois, Urbana, Illinois Jeffrey A. Cole School of Biosciences, University of Birmingham, Edgbaston, Birmingham Chris E. Cooper Department of Biological Sciences, University of Essex, Colchester, United Kingdom Francisco J. Corpas Departamento de Bioquimica, Biologıá Celular y Molecular de plantas, Estacioń Experimental del Zaidıń, CSIC, Spain Jason C. Crack Centre for Metalloprotein Spectroscopy and Biology, School of Chemical Sciences and Pharmacy, University of East Anglia, Norwich, United Kingdom Rainer Cramm Institut fu¨r Biologie/Mikrobiologie, Humboldt-Universita¨t zu Berlin, Berlin, Germany Alejandro Crespo Departamento de Quı´mica Inorgańica, Analı´tica y Quı´mica Fı´sica/INQUIMAECONICET, Facultad de Ciencias Exactas y Naturales, Universidad de Buenos Aires, Ciudad Universitaria, Buenos Aires, Argentina Benoıˆt D’Autreáux Laboratoire Stress Oxydant et Cancer, Service de Biologie Inte´grative et Geńe´tique Molećulaire, Institut de Biologie et de Technologies de Saclay, CEA-Saclay, Gif-sur-Yvette Cedex, France M. Delledonne Dipartimento Scientifico e Tecnologico, Universita` degli Studi di Verona, Verona, Italy Filip Desmet University of Antwerp, Department of Physics, SIBAC Laboratory, Antwerp, Belgium Ray Dixon John Innes Centre, Colney, Norwich, United Kingdom Sabine Van Doorslaer University of Antwerp, Department of Physics, SIBAC Laboratory, Antwerp, Belgium

xx

Contributors

Sam Efromovich Department of Mathematical Sciences, University of Texas at Dallas, Richardson, Texas Tsuyoshi Egawa Department of Physiology and Biophysics, Albert Einstein College of Medicine, Bronx, New York Francisco J. Esteban Grupo de Senãlizacioń Molecular y Sistemas Antioxidantes en Plantas, Unidad ´ rea de Bioquı´mica y Biologıá Molecular, Universidad Asociada al CSIC (EEZ), A de Jaeń, E-23071 Jaeń, Spain Dario A. Estrin Departamento de Quı´mica Inorgańica, Analı´tica y Quı´mica Fı´sica/INQUIMAECONICET, Facultad de Ciencias Exactas y Naturales, Universidad de Buenos Aires, Ciudad Universitaria, Buenos Aires, Argentina Serena Faggiano Department of Biochemistry and Molecular Biology, University of Parma, Parma, Italy, and Dipartimento di Biochimica e Biologia Molecolare, Universita` degli Studi di Parma, Parma, Italy Ferric C. Fang University of Washington School of Medicine, Seattle, Washington Sarah J. Field Center for Metalloprotein Spectroscopy and Biology, School of Biological Sciences, University of East Anglia, Norwich, United Kingdom Elena Forte Department of Biochemical Sciences, CNR Institute of Molecular Biology and Pathology and Istituto Pasteur–Fondazione Cenci Bolognetti Sapienza, University of Rome, Rome, Italy Carlos Frazaõ Instituto de Tecnologia Quı´mica e Biolo´gica, Universidade Nova de Lisboa, Oeiras, Portugal Tracey Allen K. Freitas Department of Microbiology, University of Hawaii, Honolulu, Hawaii Marie-Alda Gilles-Gonzalez Department of Biochemistry, University of Texas Southwestern Medical Center, Dallas, Texas Alessandro Giuffre` Department of Biochemical Sciences, CNR Institute of Molecular Biology and Pathology and Istituto Pasteur–Fondazione Cenci Bolognetti Sapienza, University of Rome, Rome, Italy

Contributors

xxi

Stephen D. Golden Department of Chemistry, Loyola University Chicago, Chicago, Illinois Vera L. Gonçalves Instituto de Tecnologia Quı´mica e Biolo´gica, Universidade Nova de Lisboa, Oeiras, Portugal Esther M. Gonza´lez Departamento de Ciencias del Medio Natural, Universidad Pu´blica de Navarra, Campus de Arrosadia, E-31006 Pamplona, Navarre, Spain Gonzalo Gonzalez Department of Biochemistry, University of Texas Southwestern Medical Center, Dallas, Texas David Grainger School of Biosciences, University of Birmingham, Edgbaston, Birmingham, United Kingdom Elena Grandi Dipartimento di Fisica, Universita` degli Studi di Parma, Parma, Italy Jeffrey Green Department of Molecular Biology and Biotechnology, University of Sheffield, Sheffield, United Kingdom Victor Guallar Catalan Institute for Research and Advanced Studies (ICREA), Computational Biology Program, Barcelona Supercomputing Center, Barcelona, Spain Kim H. Hebelstrup Department of Plant Science, University of Manitoba, Winnipeg, Manitoba, Canada Andrew M. Hemmings Centre for Metalloprotein Spectroscopy and Biology, School of Biological Sciences, and School of Chemical Sciences and Pharmacy, University of East Anglia, Norwich, United Kingdom Robert D. Hill Department of Plant Science, University of Manitoba, Winnipeg, Manitoba, Canada Jay C. D. Hinton Institute of Food Research, Norwich, United Kingdom Rebecca S. Holladay Department of Biological Sciences, University of Essex, Colchester, United Kingdom Adrian J. Jervis Department of Molecular Biology and Biotechnology, University of Sheffield, Sheffield, United Kingdom

xxii

Contributors

Jessica Jones-Carson University of Colorado Health Sciences Center, Aurora, Colorado Marta C. Justino Instituto de Tecnologia Quı´mica e Biolo´gica, Universidade Nova de Lisboa, Oeiras, Portugal Gemma Kemp Centre for Metalloprotein Spectroscopy and Biology, School of Chemical Sciences and Pharmacy, University of East Anglia, Norwich, United Kingdom Estı´baliz Larrainzar Departamento de Ciencias del Medio Natural, Universidad Pu´blica de Navarra, Campus de Arrosadia, E-31006 Pamplona, Navarre, Spain Lucy Lee Department of Molecular Biology and Biotechnology, University of Sheffield, Sheffield, United Kingdom Changyuan Lu Department of Physiology and Biophysics, Albert Einstein College of Medicine, Bronx, New York F. Javier Luque Departament de Fisicoquı´mica, Facultat de Farma`cia, Universitat de Barcelona, Barcelona, Spain Marcelo A. Marti Departamento de Quı´mica Inorgańica, Analı´tica y Quı´mica Fı´sica/INQUIMAECONICET, Facultad de Ciencias Exactas y Naturales, Universidad de Buenos Aires, Ciudad Universitaria, Buenos Aires, Argentina Maria G. Mason Department of Biological Sciences, University of Essex, Colchester, United Kingdom Andrey D. Matorin Center for Metalloprotein Spectroscopy and Biology, School of Biological Sciences, University of East Anglia, Norwich, United Kingdom Paul C. Mills Centre for Metalloprotein Spectroscopy and Biology, School of Biological Sciences, University of East Anglia, Norwich, United Kingdom James W. B. Moir Department of Biology, University of York, Heslington, York Claire E. Monk Department of Molecular Biology and Biotechnology, University of Sheffield, Sheffield, United Kingdom

Contributors

xxiii

Jose F. Moran Instituto de Agrobiotecnologıá, Universidad Pu´blica de Navarra-CSIC-Gobierno de Navarra, Campus de Arrosadıá, E-31006 Pamplona, Navarre, Spain Andrea Mozzarelli Department of Biochemistry and Molecular Biology, University of Parma, Parma, Italy, and Dipartimento di Biochimica e Biologia Molecolare, Universita` degli Studi di Parma, Parma, Italy Masahiro Mukai Mitsubishi Kagaku Institute of Life Sciences, Minamiooya, Machida, Tokyo, Japan Peter Nicholls Department of Biological Sciences, University of Essex, Colchester, United Kingdom Karin Nienhaus Institute of Biophysics, University of Ulm, Ulm, Germany G. Ulrich Nienhaus Institute of Biophysics, University of Ulm, Ulm, Germany, and Department of Physics, University of Illinois at Urbana–Champaign, Urbana, Illinois Erik stergaard-Jensen Department of Molecular Biology, University of Aarhus, Aarhus, Denmark Kenneth W. Olsen Department of Chemistry, Loyola University Chicago, Chicago, Illinois Susie R. Poock Centre for Metalloprotein Spectroscopy and Biology, School of Biological Sciences, University of East Anglia, Norwich, United Kingdom Robert K. Poole Department of Molecular Biology and Biotechnology, University of Sheffield, Sheffield, United Kingdom Steven T. Pullan Department of Molecular Biology and Biotechnology, University of Sheffield, Sheffield, United Kingdom Robert C. Read University of Sheffield, Sheffield, United Kingdom David J. Richardson Center for Metalloprotein Spectroscopy and Biology, School of Biological Sciences, University of East Anglia, Norwich, United Kingdom Luis A. Del Rıó Departamento de Bioquimica, Biologıá Celular y Molecular de plantas, Estacioń Experimental del Zaidıń (EEZ), CSIC, Apartado 419, E-18080 Grannada, Spain

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Contributors

Luca Ronda Department of Biochemistry and Molecular Biology, University of Parma, Parma, Italy, and Dipartimento di Biochimica e Biologia Molecolare, Universita` degli Studi di Parma, Parma, Italy William E. Royer, Jr. Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, Massachusetts Jennifer A. Saito Department of Microbiology, University of Hawaii, Honolulu, Hawaii Lı´gia M. Saraiva Instituto de Tecnologia Quı´mica e Biolo´gica, Universidade Nova de Lisboa, Oeiras, Portugal Paolo Sarti Department of Biochemical Sciences, CNR Institute of Molecular Biology and Pathology and Istituto Pasteur–Fondazione Cenci Bolognetti Sapienza, University of Rome, Rome, Italy Francesca M. Scandurra Department of Biochemical Sciences, CNR Institute of Molecular Biology and Pathology and Istituto Pasteur–Fondazione Cenci Bolognetti Sapienza, University of Rome, Rome, Italy Klaus Schulten Beckman Institute, University of Illinois, Urbana, Illinois Vukica Sˇrajer Center for Advanced Radiation Sources, University of Chicago, Illinois ¨derberg Christopher A. G. So Centre for Metalloprotein Spectroscopy and Biology, School of Biological Sciences, University of East Anglia, Norwich, United Kingdom Mark Shepherd University of Sheffield, Western Bank, Sheffield, United Kingdom Hirofumi Shoun Department of Biotechnology, University of Tokyo, Bunkyo-ku, Tokyo, Japan Eduardo Henrique Silva Sousa Department of Biochemistry, University of Texas Southwestern Medical Center, Dallas, Texas Stephen Spiro Department of Molecular and Cell Biology, University of Texas at Dallas, Richardson, Texas

Contributors

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Tania M. Stevanin School of Medicine and Biomedical Science, University of Sheffield, Sheffield, United Kingdom Tania Stevanin University of Sheffield, Sheffield, United Kingdom Katja Strube Institut fu¨r Biologie/Mikrobiologie, Humboldt-Universita¨t zu Berlin, Berlin, Germany Miguel Teixeira Instituto de Tecnologia Quı´mica e Biolo´gica, Universidade Nova de Lisboa, Oeiras, Portugal Andrew J. Thomson Centre for Metalloprotein Spectroscopy and Biology, School of Chemical Sciences and Pharmacy, University of East Anglia, Norwich, United Kingdom Faye H. Thorndycroft Center for Metalloprotein Spectroscopy and Biology, School of Biological Sciences, University of East Anglia, Norwich, United Kingdom Melanie J. Thomson Department of Biology, University of York, Heslington, York Nick Tucker John Innes Centre, Colney, Norwich, United Kingdom Jason Tuckerman Department of Biochemistry, University of Texas Southwestern Medical Center, Dallas, Texas Estı´baliz Urarte Instituto de Agrobiotecnologıá, Universidad Pu´blica de Navarra-CSIC-Gobierno de Navarra, Campus de Arrosadıá, E-31006 Pamplona, Navarre, Spain Raquel Valderrama Grupo de Senãlizacioń Molecular y Sistemas Antioxidantes en Plantas, Unidad ´ rea de Bioquı´mica y Biologıá Molecular, Universidad Asociada al CSIC (EEZ), A de Jaeń, E-23071 Jaeń, Spain B. Vallone Dipartimento di Scienze Biochimiche ‘‘A. Rossi Fanelli,’’ Universita` di Roma ‘‘La Sapienza,’’ Roma, Italy E. Vandelle Dipartimento Scientifico e Tecnologico, Universita` degli Studi di Verona, Verona, Italy

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Contributors

Andre´s Vazquez-Torres University of Colorado Health Sciences Center, Aurora, Colorado Cristiano Viappiani Dipartimento di Fisica, Universita` degli Studi di Parma, Parma, Italy, and NEST CNR-INFM, Pisa, Italy Joaõ B. Vicente Instituto de Tecnologia Quı´mica e Biolo´gica, Universidade Nova de Lisboa, Oeiras, Portugal Nicholas J. Watmough Center for Metalloprotein Spectroscopy and Biology, School of Biological Sciences, University of East Anglia, Norwich, United Kingdom Jessica Van Wonderen Centre for Metalloprotein Spectroscopy and Biology, School of Chemical Sciences and Pharmacy, University of East Anglia, Norwich, United Kingdom Syun-Ru Yeh Department of Physiology and Biophysics, Albert Einstein College of Medicine, Bronx, New York Li Zhang Department of Biology, University of Kentucky, Lexington, Kentucky

PREFACE

The genesis of ideas for these two volumes of Methods in Enzymology appears to be a talk (subtitled Bloody Bacteria) that I presented at the Agouron Institute meeting in Santa Fe, New Mexico, in April 2006. The topic of the meeting was Oxygen, but my message was not how microbial hemoglobins manage oxygen but rather how the primary function of many such hemoglobins is nitric oxide detoxification. Despite my straying from my brief, John Abelson and Mel Simon generously invited me to consider editing a volume of Methods in Enzymology to cover these emerging aspects of such a well-studied protein family. Further discussion of the proposal at the XIVth International Conference on Dioxygen Binding and Sensing Proteins at Stazione Zoologica Anton Dohrn in beautiful Napoli later that year—warmly hosted by Cinzia Verde and Guido di Prisco—generated much interest and support. The result was a two-volume heterodimer: I hope cooperativity can be found in Volumes 436 and 437. Just as the organizers of the Agouron Institute conference interpreted Oxygen with commendable flexibility, Methods in Enzymology has allowed some freedom in the definition of an enzyme. In 1994, when the topic Hemoglobins (Part C) was last covered explicitly in this series (Volume 232), some justification for labeling a hemoglobin as an enzyme might have been warranted. But as Maurizio Brunori pointed out in 1999 (Trends in Biochemical Sciences, 24, 158–161), the promotion of hemoglobin to the status of ‘‘honorary enzyme’’ had been conferred decades earlier by Monod, Wyman, and Changeux. In 2007, the idea that certain hemoglobins, even those not displaying allosteric heme–heme interactions, have enzymatic functions is well established, the most obvious examples being those hemoglobins that transform substrates into products, such as nitric oxide into nitrate. Other topics covered in these volumes are not new to the Methods in Enzymology series either. The most recent coverage of overtly related topics was Nitric Oxide (Part E) in Volume 396 (2005) and Oxygen Sensing in Volume 381 (2004). I hope, however, that the particular juxtaposition of topics in these two volumes will draw attention to the intimate links between globins, their gaseous ligands (nitric oxide, oxygen, and carbon monoxide), and the sensing and detoxification of these biologically critical small molecules. There is a strong microbial flavor in these volumes, reflecting some of the most exciting developments in recent years. Volume

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Preface

436 deals with some chemical and analytical aspects of nitric oxide and methods for bacterial and archaeal hemoglobins, as well as diverse (especially ‘‘newer’’) hemoglobins in plants and animals. Volume 437 covers various non-hemoglobin nitric oxide-detoxifying proteins, sensors for gaseous ligands, advanced spectroscopic tools, and aspects of the functions of these proteins in microbial and plant physiology. In each volume, some chapters serve not as methodological recipes but short reviews to place the methods in a proper framework. These volumes would not have been possible without the tremendous enthusiasm of so many colleagues, contributors, and friends around the world. I thank them all, and also Tari Broderick and Cindy Minor (Elsevier, San Diego, California), for their help and encouragement in leading these volumes to a successful and timely outcome. ROBERT K. POOLE

METHODS IN ENZYMOLOGY

VOLUME I. Preparation and Assay of Enzymes Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME II. Preparation and Assay of Enzymes Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME III. Preparation and Assay of Substrates Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME IV. Special Techniques for the Enzymologist Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME V. Preparation and Assay of Enzymes Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME VI. Preparation and Assay of Enzymes (Continued) Preparation and Assay of Substrates Special Techniques Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME VII. Cumulative Subject Index Edited by SIDNEY P. COLOWICK AND NATHAN O. KAPLAN VOLUME VIII. Complex Carbohydrates Edited by ELIZABETH F. NEUFELD AND VICTOR GINSBURG VOLUME IX. Carbohydrate Metabolism Edited by WILLIS A. WOOD VOLUME X. Oxidation and Phosphorylation Edited by RONALD W. ESTABROOK AND MAYNARD E. PULLMAN VOLUME XI. Enzyme Structure Edited by C. H. W. HIRS VOLUME XII. Nucleic Acids (Parts A and B) Edited by LAWRENCE GROSSMAN AND KIVIE MOLDAVE VOLUME XIII. Citric Acid Cycle Edited by J. M. LOWENSTEIN VOLUME XIV. Lipids Edited by J. M. LOWENSTEIN VOLUME XV. Steroids and Terpenoids Edited by RAYMOND B. CLAYTON xxix

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VOLUME XVI. Fast Reactions Edited by KENNETH KUSTIN VOLUME XVII. Metabolism of Amino Acids and Amines (Parts A and B) Edited by HERBERT TABOR AND CELIA WHITE TABOR VOLUME XVIII. Vitamins and Coenzymes (Parts A, B, and C) Edited by DONALD B. MCCORMICK AND LEMUEL D. WRIGHT VOLUME XIX. Proteolytic Enzymes Edited by GERTRUDE E. PERLMANN AND LASZLO LORAND VOLUME XX. Nucleic Acids and Protein Synthesis (Part C) Edited by KIVIE MOLDAVE AND LAWRENCE GROSSMAN VOLUME XXI. Nucleic Acids (Part D) Edited by LAWRENCE GROSSMAN AND KIVIE MOLDAVE VOLUME XXII. Enzyme Purification and Related Techniques Edited by WILLIAM B. JAKOBY VOLUME XXIII. Photosynthesis (Part A) Edited by ANTHONY SAN PIETRO VOLUME XXIV. Photosynthesis and Nitrogen Fixation (Part B) Edited by ANTHONY SAN PIETRO VOLUME XXV. Enzyme Structure (Part B) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME XXVI. Enzyme Structure (Part C) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME XXVII. Enzyme Structure (Part D) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME XXVIII. Complex Carbohydrates (Part B) Edited by VICTOR GINSBURG VOLUME XXIX. Nucleic Acids and Protein Synthesis (Part E) Edited by LAWRENCE GROSSMAN AND KIVIE MOLDAVE VOLUME XXX. Nucleic Acids and Protein Synthesis (Part F) Edited by KIVIE MOLDAVE AND LAWRENCE GROSSMAN VOLUME XXXI. Biomembranes (Part A) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME XXXII. Biomembranes (Part B) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME XXXIII. Cumulative Subject Index Volumes I-XXX Edited by MARTHA G. DENNIS AND EDWARD A. DENNIS VOLUME XXXIV. Affinity Techniques (Enzyme Purification: Part B) Edited by WILLIAM B. JAKOBY AND MEIR WILCHEK


VOLUME XXXV. Lipids (Part B) Edited by JOHN M. LOWENSTEIN VOLUME XXXVI. Hormone Action (Part A: Steroid Hormones) Edited by BERT W. O’MALLEY AND JOEL G. HARDMAN VOLUME XXXVII. Hormone Action (Part B: Peptide Hormones) Edited by BERT W. O’MALLEY AND JOEL G. HARDMAN VOLUME XXXVIII. Hormone Action (Part C: Cyclic Nucleotides) Edited by JOEL G. HARDMAN AND BERT W. O’MALLEY VOLUME XXXIX. Hormone Action (Part D: Isolated Cells, Tissues, and Organ Systems) Edited by JOEL G. HARDMAN AND BERT W. O’MALLEY VOLUME XL. Hormone Action (Part E: Nuclear Structure and Function) Edited by BERT W. O’MALLEY AND JOEL G. HARDMAN VOLUME XLI. Carbohydrate Metabolism (Part B) Edited by W. A. WOOD VOLUME XLII. Carbohydrate Metabolism (Part C) Edited by W. A. WOOD VOLUME XLIII. Antibiotics Edited by JOHN H. HASH VOLUME XLIV. Immobilized Enzymes Edited by KLAUS MOSBACH VOLUME XLV. Proteolytic Enzymes (Part B) Edited by LASZLO LORAND VOLUME XLVI. Affinity Labeling Edited by WILLIAM B. JAKOBY AND MEIR WILCHEK VOLUME XLVII. Enzyme Structure (Part E) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME XLVIII. Enzyme Structure (Part F) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME XLIX. Enzyme Structure (Part G) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME L. Complex Carbohydrates (Part C) Edited by VICTOR GINSBURG VOLUME LI. Purine and Pyrimidine Nucleotide Metabolism Edited by PATRICIA A. HOFFEE AND MARY ELLEN JONES VOLUME LII. Biomembranes (Part C: Biological Oxidations) Edited by SIDNEY FLEISCHER AND LESTER PACKER

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VOLUME LIII. Biomembranes (Part D: Biological Oxidations) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME LIV. Biomembranes (Part E: Biological Oxidations) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME LV. Biomembranes (Part F: Bioenergetics) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME LVI. Biomembranes (Part G: Bioenergetics) Edited by SIDNEY FLEISCHER AND LESTER PACKER VOLUME LVII. Bioluminescence and Chemiluminescence Edited by MARLENE A. DELUCA VOLUME LVIII. Cell Culture Edited by WILLIAM B. JAKOBY AND IRA PASTAN VOLUME LIX. Nucleic Acids and Protein Synthesis (Part G) Edited by KIVIE MOLDAVE AND LAWRENCE GROSSMAN VOLUME LX. Nucleic Acids and Protein Synthesis (Part H) Edited by KIVIE MOLDAVE AND LAWRENCE GROSSMAN VOLUME 61. Enzyme Structure (Part H) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME 62. Vitamins and Coenzymes (Part D) Edited by DONALD B. MCCORMICK AND LEMUEL D. WRIGHT VOLUME 63. Enzyme Kinetics and Mechanism (Part A: Initial Rate and Inhibitor Methods) Edited by DANIEL L. PURICH VOLUME 64. Enzyme Kinetics and Mechanism (Part B: Isotopic Probes and Complex Enzyme Systems) Edited by DANIEL L. PURICH VOLUME 65. Nucleic Acids (Part I) Edited by LAWRENCE GROSSMAN AND KIVIE MOLDAVE VOLUME 66. Vitamins and Coenzymes (Part E) Edited by DONALD B. MCCORMICK AND LEMUEL D. WRIGHT VOLUME 67. Vitamins and Coenzymes (Part F) Edited by DONALD B. MCCORMICK AND LEMUEL D. WRIGHT VOLUME 68. Recombinant DNA Edited by RAY WU VOLUME 69. Photosynthesis and Nitrogen Fixation (Part C) Edited by ANTHONY SAN PIETRO VOLUME 70. Immunochemical Techniques (Part A) Edited by HELEN VAN VUNAKIS AND JOHN J. LANGONE


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VOLUME 71. Lipids (Part C) Edited by JOHN M. LOWENSTEIN VOLUME 72. Lipids (Part D) Edited by JOHN M. LOWENSTEIN VOLUME 73. Immunochemical Techniques (Part B) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 74. Immunochemical Techniques (Part C) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 75. Cumulative Subject Index Volumes XXXI, XXXII, XXXIV–LX Edited by EDWARD A. DENNIS AND MARTHA G. DENNIS VOLUME 76. Hemoglobins Edited by ERALDO ANTONINI, LUIGI ROSSI-BERNARDI, AND EMILIA CHIANCONE VOLUME 77. Detoxication and Drug Metabolism Edited by WILLIAM B. JAKOBY VOLUME 78. Interferons (Part A) Edited by SIDNEY PESTKA VOLUME 79. Interferons (Part B) Edited by SIDNEY PESTKA VOLUME 80. Proteolytic Enzymes (Part C) Edited by LASZLO LORAND VOLUME 81. Biomembranes (Part H: Visual Pigments and Purple Membranes, I) Edited by LESTER PACKER VOLUME 82. Structural and Contractile Proteins (Part A: Extracellular Matrix) Edited by LEON W. CUNNINGHAM AND DIXIE W. FREDERIKSEN VOLUME 83. Complex Carbohydrates (Part D) Edited by VICTOR GINSBURG VOLUME 84. Immunochemical Techniques (Part D: Selected Immunoassays) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 85. Structural and Contractile Proteins (Part B: The Contractile Apparatus and the Cytoskeleton) Edited by DIXIE W. FREDERIKSEN AND LEON W. CUNNINGHAM VOLUME 86. Prostaglandins and Arachidonate Metabolites Edited by WILLIAM E. M. LANDS AND WILLIAM L. SMITH VOLUME 87. Enzyme Kinetics and Mechanism (Part C: Intermediates, Stereo-chemistry, and Rate Studies) Edited by DANIEL L. PURICH VOLUME 88. Biomembranes (Part I: Visual Pigments and Purple Membranes, II) Edited by LESTER PACKER

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VOLUME 89. Carbohydrate Metabolism (Part D) Edited by WILLIS A. WOOD VOLUME 90. Carbohydrate Metabolism (Part E) Edited by WILLIS A. WOOD VOLUME 91. Enzyme Structure (Part I) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME 92. Immunochemical Techniques (Part E: Monoclonal Antibodies and General Immunoassay Methods) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 93. Immunochemical Techniques (Part F: Conventional Antibodies, Fc Receptors, and Cytotoxicity) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 94. Polyamines Edited by HERBERT TABOR AND CELIA WHITE TABOR VOLUME 95. Cumulative Subject Index Volumes 61–74, 76–80 Edited by EDWARD A. DENNIS AND MARTHA G. DENNIS VOLUME 96. Biomembranes [Part J: Membrane Biogenesis: Assembly and Targeting (General Methods; Eukaryotes)] Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 97. Biomembranes [Part K: Membrane Biogenesis: Assembly and Targeting (Prokaryotes, Mitochondria, and Chloroplasts)] Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 98. Biomembranes (Part L: Membrane Biogenesis: Processing and Recycling) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 99. Hormone Action (Part F: Protein Kinases) Edited by JACKIE D. CORBIN AND JOEL G. HARDMAN VOLUME 100. Recombinant DNA (Part B) Edited by RAY WU, LAWRENCE GROSSMAN, AND KIVIE MOLDAVE VOLUME 101. Recombinant DNA (Part C) Edited by RAY WU, LAWRENCE GROSSMAN, AND KIVIE MOLDAVE VOLUME 102. Hormone Action (Part G: Calmodulin and Calcium-Binding Proteins) Edited by ANTHONY R. MEANS AND BERT W. O’MALLEY VOLUME 103. Hormone Action (Part H: Neuroendocrine Peptides) Edited by P. MICHAEL CONN VOLUME 104. Enzyme Purification and Related Techniques (Part C) Edited by WILLIAM B. JAKOBY


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VOLUME 105. Oxygen Radicals in Biological Systems Edited by LESTER PACKER VOLUME 106. Posttranslational Modifications (Part A) Edited by FINN WOLD AND KIVIE MOLDAVE VOLUME 107. Posttranslational Modifications (Part B) Edited by FINN WOLD AND KIVIE MOLDAVE VOLUME 108. Immunochemical Techniques (Part G: Separation and Characterization of Lymphoid Cells) Edited by GIOVANNI DI SABATO, JOHN J. LANGONE, AND HELEN VAN VUNAKIS VOLUME 109. Hormone Action (Part I: Peptide Hormones) Edited by LUTZ BIRNBAUMER AND BERT W. O’MALLEY VOLUME 110. Steroids and Isoprenoids (Part A) Edited by JOHN H. LAW AND HANS C. RILLING VOLUME 111. Steroids and Isoprenoids (Part B) Edited by JOHN H. LAW AND HANS C. RILLING VOLUME 112. Drug and Enzyme Targeting (Part A) Edited by KENNETH J. WIDDER AND RALPH GREEN VOLUME 113. Glutamate, Glutamine, Glutathione, and Related Compounds Edited by ALTON MEISTER VOLUME 114. Diffraction Methods for Biological Macromolecules (Part A) Edited by HAROLD W. WYCKOFF, C. H. W. HIRS, AND SERGE N. TIMASHEFF VOLUME 115. Diffraction Methods for Biological Macromolecules (Part B) Edited by HAROLD W. WYCKOFF, C. H. W. HIRS, AND SERGE N. TIMASHEFF VOLUME 116. Immunochemical Techniques (Part H: Effectors and Mediators of Lymphoid Cell Functions) Edited by GIOVANNI DI SABATO, JOHN J. LANGONE, AND HELEN VAN VUNAKIS VOLUME 117. Enzyme Structure (Part J) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME 118. Plant Molecular Biology Edited by ARTHUR WEISSBACH AND HERBERT WEISSBACH VOLUME 119. Interferons (Part C) Edited by SIDNEY PESTKA VOLUME 120. Cumulative Subject Index Volumes 81–94, 96–101 VOLUME 121. Immunochemical Techniques (Part I: Hybridoma Technology and Monoclonal Antibodies) Edited by JOHN J. LANGONE AND HELEN VAN VUNAKIS VOLUME 122. Vitamins and Coenzymes (Part G) Edited by FRANK CHYTIL AND DONALD B. MCCORMICK

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VOLUME 123. Vitamins and Coenzymes (Part H) Edited by FRANK CHYTIL AND DONALD B. MCCORMICK VOLUME 124. Hormone Action (Part J: Neuroendocrine Peptides) Edited by P. MICHAEL CONN VOLUME 125. Biomembranes (Part M: Transport in Bacteria, Mitochondria, and Chloroplasts: General Approaches and Transport Systems) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 126. Biomembranes (Part N: Transport in Bacteria, Mitochondria, and Chloroplasts: Protonmotive Force) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 127. Biomembranes (Part O: Protons and Water: Structure and Translocation) Edited by LESTER PACKER VOLUME 128. Plasma Lipoproteins (Part A: Preparation, Structure, and Molecular Biology) Edited by JERE P. SEGREST AND JOHN J. ALBERS VOLUME 129. Plasma Lipoproteins (Part B: Characterization, Cell Biology, and Metabolism) Edited by JOHN J. ALBERS AND JERE P. SEGREST VOLUME 130. Enzyme Structure (Part K) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME 131. Enzyme Structure (Part L) Edited by C. H. W. HIRS AND SERGE N. TIMASHEFF VOLUME 132. Immunochemical Techniques (Part J: Phagocytosis and Cell-Mediated Cytotoxicity) Edited by GIOVANNI DI SABATO AND JOHANNES EVERSE VOLUME 133. Bioluminescence and Chemiluminescence (Part B) Edited by MARLENE DELUCA AND WILLIAM D. MCELROY VOLUME 134. Structural and Contractile Proteins (Part C: The Contractile Apparatus and the Cytoskeleton) Edited by RICHARD B. VALLEE VOLUME 135. Immobilized Enzymes and Cells (Part B) Edited by KLAUS MOSBACH VOLUME 136. Immobilized Enzymes and Cells (Part C) Edited by KLAUS MOSBACH VOLUME 137. Immobilized Enzymes and Cells (Part D) Edited by KLAUS MOSBACH VOLUME 138. Complex Carbohydrates (Part E) Edited by VICTOR GINSBURG


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VOLUME 139. Cellular Regulators (Part A: Calcium- and Calmodulin-Binding Proteins) Edited by ANTHONY R. MEANS AND P. MICHAEL CONN VOLUME 140. Cumulative Subject Index Volumes 102–119, 121–134 VOLUME 141. Cellular Regulators (Part B: Calcium and Lipids) Edited by P. MICHAEL CONN AND ANTHONY R. MEANS VOLUME 142. Metabolism of Aromatic Amino Acids and Amines Edited by SEYMOUR KAUFMAN VOLUME 143. Sulfur and Sulfur Amino Acids Edited by WILLIAM B. JAKOBY AND OWEN GRIFFITH VOLUME 144. Structural and Contractile Proteins (Part D: Extracellular Matrix) Edited by LEON W. CUNNINGHAM VOLUME 145. Structural and Contractile Proteins (Part E: Extracellular Matrix) Edited by LEON W. CUNNINGHAM VOLUME 146. Peptide Growth Factors (Part A) Edited by DAVID BARNES AND DAVID A. SIRBASKU VOLUME 147. Peptide Growth Factors (Part B) Edited by DAVID BARNES AND DAVID A. SIRBASKU VOLUME 148. Plant Cell Membranes Edited by LESTER PACKER AND ROLAND DOUCE VOLUME 149. Drug and Enzyme Targeting (Part B) Edited by RALPH GREEN AND KENNETH J. WIDDER VOLUME 150. Immunochemical Techniques (Part K: In Vitro Models of B and T Cell Functions and Lymphoid Cell Receptors) Edited by GIOVANNI DI SABATO VOLUME 151. Molecular Genetics of Mammalian Cells Edited by MICHAEL M. GOTTESMAN VOLUME 152. Guide to Molecular Cloning Techniques Edited by SHELBY L. BERGER AND ALAN R. KIMMEL VOLUME 153. Recombinant DNA (Part D) Edited by RAY WU AND LAWRENCE GROSSMAN VOLUME 154. Recombinant DNA (Part E) Edited by RAY WU AND LAWRENCE GROSSMAN VOLUME 155. Recombinant DNA (Part F) Edited by RAY WU VOLUME 156. Biomembranes (Part P: ATP-Driven Pumps and Related Transport: The Na, K-Pump) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER

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VOLUME 157. Biomembranes (Part Q: ATP-Driven Pumps and Related Transport: Calcium, Proton, and Potassium Pumps) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 158. Metalloproteins (Part A) Edited by JAMES F. RIORDAN AND BERT L. VALLEE VOLUME 159. Initiation and Termination of Cyclic Nucleotide Action Edited by JACKIE D. CORBIN AND ROGER A. JOHNSON VOLUME 160. Biomass (Part A: Cellulose and Hemicellulose) Edited by WILLIS A. WOOD AND SCOTT T. KELLOGG VOLUME 161. Biomass (Part B: Lignin, Pectin, and Chitin) Edited by WILLIS A. WOOD AND SCOTT T. KELLOGG VOLUME 162. Immunochemical Techniques (Part L: Chemotaxis and Inflammation) Edited by GIOVANNI DI SABATO VOLUME 163. Immunochemical Techniques (Part M: Chemotaxis and Inflammation) Edited by GIOVANNI DI SABATO VOLUME 164. Ribosomes Edited by HARRY F. NOLLER, JR., AND KIVIE MOLDAVE VOLUME 165. Microbial Toxins: Tools for Enzymology Edited by SIDNEY HARSHMAN VOLUME 166. Branched-Chain Amino Acids Edited by ROBERT HARRIS AND JOHN R. SOKATCH VOLUME 167. Cyanobacteria Edited by LESTER PACKER AND ALEXANDER N. GLAZER VOLUME 168. Hormone Action (Part K: Neuroendocrine Peptides) Edited by P. MICHAEL CONN VOLUME 169. Platelets: Receptors, Adhesion, Secretion (Part A) Edited by JACEK HAWIGER VOLUME 170. Nucleosomes Edited by PAUL M. WASSARMAN AND ROGER D. KORNBERG VOLUME 171. Biomembranes (Part R: Transport Theory: Cells and Model Membranes) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 172. Biomembranes (Part S: Transport: Membrane Isolation and Characterization) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER


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VOLUME 173. Biomembranes [Part T: Cellular and Subcellular Transport: Eukaryotic (Nonepithelial) Cells] Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 174. Biomembranes [Part U: Cellular and Subcellular Transport: Eukaryotic (Nonepithelial) Cells] Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 175. Cumulative Subject Index Volumes 135–139, 141–167 VOLUME 176. Nuclear Magnetic Resonance (Part A: Spectral Techniques and Dynamics) Edited by NORMAN J. OPPENHEIMER AND THOMAS L. JAMES VOLUME 177. Nuclear Magnetic Resonance (Part B: Structure and Mechanism) Edited by NORMAN J. OPPENHEIMER AND THOMAS L. JAMES VOLUME 178. Antibodies, Antigens, and Molecular Mimicry Edited by JOHN J. LANGONE VOLUME 179. Complex Carbohydrates (Part F) Edited by VICTOR GINSBURG VOLUME 180. RNA Processing (Part A: General Methods) Edited by JAMES E. DAHLBERG AND JOHN N. ABELSON VOLUME 181. RNA Processing (Part B: Specific Methods) Edited by JAMES E. DAHLBERG AND JOHN N. ABELSON VOLUME 182. Guide to Protein Purification Edited by MURRAY P. DEUTSCHER VOLUME 183. Molecular Evolution: Computer Analysis of Protein and Nucleic Acid Sequences Edited by RUSSELL F. DOOLITTLE VOLUME 184. Avidin-Biotin Technology Edited by MEIR WILCHEK AND EDWARD A. BAYER VOLUME 185. Gene Expression Technology Edited by DAVID V. GOEDDEL VOLUME 186. Oxygen Radicals in Biological Systems (Part B: Oxygen Radicals and Antioxidants) Edited by LESTER PACKER AND ALEXANDER N. GLAZER VOLUME 187. Arachidonate Related Lipid Mediators Edited by ROBERT C. MURPHY AND FRANK A. FITZPATRICK VOLUME 188. Hydrocarbons and Methylotrophy Edited by MARY E. LIDSTROM VOLUME 189. Retinoids (Part A: Molecular and Metabolic Aspects) Edited by LESTER PACKER

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VOLUME 190. Retinoids (Part B: Cell Differentiation and Clinical Applications) Edited by LESTER PACKER VOLUME 191. Biomembranes (Part V: Cellular and Subcellular Transport: Epithelial Cells) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 192. Biomembranes (Part W: Cellular and Subcellular Transport: Epithelial Cells) Edited by SIDNEY FLEISCHER AND BECCA FLEISCHER VOLUME 193. Mass Spectrometry Edited by JAMES A. MCCLOSKEY VOLUME 194. Guide to Yeast Genetics and Molecular Biology Edited by CHRISTINE GUTHRIE AND GERALD R. FINK VOLUME 195. Adenylyl Cyclase, G Proteins, and Guanylyl Cyclase Edited by ROGER A. JOHNSON AND JACKIE D. CORBIN VOLUME 196. Molecular Motors and the Cytoskeleton Edited by RICHARD B. VALLEE VOLUME 197. Phospholipases Edited by EDWARD A. DENNIS VOLUME 198. Peptide Growth Factors (Part C) Edited by DAVID BARNES, J. P. MATHER, AND GORDON H. SATO VOLUME 199. Cumulative Subject Index Volumes 168–174, 176–194 VOLUME 200. Protein Phosphorylation (Part A: Protein Kinases: Assays, Purification, Antibodies, Functional Analysis, Cloning, and Expression) Edited by TONY HUNTER AND BARTHOLOMEW M. SEFTON VOLUME 201. Protein Phosphorylation (Part B: Analysis of Protein Phosphorylation, Protein Kinase Inhibitors, and Protein Phosphatases) Edited by TONY HUNTER AND BARTHOLOMEW M. SEFTON VOLUME 202. Molecular Design and Modeling: Concepts and Applications (Part A: Proteins, Peptides, and Enzymes) Edited by JOHN J. LANGONE VOLUME 203. Molecular Design and Modeling: Concepts and Applications (Part B: Antibodies and Antigens, Nucleic Acids, Polysaccharides, and Drugs) Edited by JOHN J. LANGONE VOLUME 204. Bacterial Genetic Systems Edited by JEFFREY H. MILLER VOLUME 205. Metallobiochemistry (Part B: Metallothionein and Related Molecules) Edited by JAMES F. RIORDAN AND BERT L. VALLEE


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VOLUME 206. Cytochrome P450 Edited by MICHAEL R. WATERMAN AND ERIC F. JOHNSON VOLUME 207. Ion Channels Edited by BERNARDO RUDY AND LINDA E. IVERSON VOLUME 208. Protein–DNA Interactions Edited by ROBERT T. SAUER VOLUME 209. Phospholipid Biosynthesis Edited by EDWARD A. DENNIS AND DENNIS E. VANCE VOLUME 210. Numerical Computer Methods Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 211. DNA Structures (Part A: Synthesis and Physical Analysis of DNA) Edited by DAVID M. J. LILLEY AND JAMES E. DAHLBERG VOLUME 212. DNA Structures (Part B: Chemical and Electrophoretic Analysis of DNA) Edited by DAVID M. J. LILLEY AND JAMES E. DAHLBERG VOLUME 213. Carotenoids (Part A: Chemistry, Separation, Quantitation, and Antioxidation) Edited by LESTER PACKER VOLUME 214. Carotenoids (Part B: Metabolism, Genetics, and Biosynthesis) Edited by LESTER PACKER VOLUME 215. Platelets: Receptors, Adhesion, Secretion (Part B) Edited by JACEK J. HAWIGER VOLUME 216. Recombinant DNA (Part G) Edited by RAY WU VOLUME 217. Recombinant DNA (Part H) Edited by RAY WU VOLUME 218. Recombinant DNA (Part I) Edited by RAY WU VOLUME 219. Reconstitution of Intracellular Transport Edited by JAMES E. ROTHMAN VOLUME 220. Membrane Fusion Techniques (Part A) Edited by NEJAT DU¨ZGU¨NES VOLUME 221. Membrane Fusion Techniques (Part B) Edited by NEJAT DU¨ZGU¨NES VOLUME 222. Proteolytic Enzymes in Coagulation, Fibrinolysis, and Complement Activation (Part A: Mammalian Blood Coagulation Factors and Inhibitors) Edited by LASZLO LORAND AND KENNETH G. MANN

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VOLUME 223. Proteolytic Enzymes in Coagulation, Fibrinolysis, and Complement Activation (Part B: Complement Activation, Fibrinolysis, and Nonmammalian Blood Coagulation Factors) Edited by LASZLO LORAND AND KENNETH G. MANN VOLUME 224. Molecular Evolution: Producing the Biochemical Data Edited by ELIZABETH ANNE ZIMMER, THOMAS J. WHITE, REBECCA L. CANN, AND ALLAN C. WILSON VOLUME 225. Guide to Techniques in Mouse Development Edited by PAUL M. WASSARMAN AND MELVIN L. DEPAMPHILIS VOLUME 226. Metallobiochemistry (Part C: Spectroscopic and Physical Methods for Probing Metal Ion Environments in Metalloenzymes and Metalloproteins) Edited by JAMES F. RIORDAN AND BERT L. VALLEE VOLUME 227. Metallobiochemistry (Part D: Physical and Spectroscopic Methods for Probing Metal Ion Environments in Metalloproteins) Edited by JAMES F. RIORDAN AND BERT L. VALLEE VOLUME 228. Aqueous Two-Phase Systems Edited by HARRY WALTER AND GO¨TE JOHANSSON VOLUME 229. Cumulative Subject Index Volumes 195–198, 200–227 VOLUME 230. Guide to Techniques in Glycobiology Edited by WILLIAM J. LENNARZ AND GERALD W. HART VOLUME 231. Hemoglobins (Part B: Biochemical and Analytical Methods) Edited by JOHANNES EVERSE, KIM D. VANDEGRIFF, AND ROBERT M. WINSLOW VOLUME 232. Hemoglobins (Part C: Biophysical Methods) Edited by JOHANNES EVERSE, KIM D. VANDEGRIFF, AND ROBERT M. WINSLOW VOLUME 233. Oxygen Radicals in Biological Systems (Part C) Edited by LESTER PACKER VOLUME 234. Oxygen Radicals in Biological Systems (Part D) Edited by LESTER PACKER VOLUME 235. Bacterial Pathogenesis (Part A: Identification and Regulation of Virulence Factors) Edited by VIRGINIA L. CLARK AND PATRIK M. BAVOIL VOLUME 236. Bacterial Pathogenesis (Part B: Integration of Pathogenic Bacteria with Host Cells) Edited by VIRGINIA L. CLARK AND PATRIK M. BAVOIL VOLUME 237. Heterotrimeric G Proteins Edited by RAVI IYENGAR VOLUME 238. Heterotrimeric G-Protein Effectors Edited by RAVI IYENGAR


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VOLUME 239. Nuclear Magnetic Resonance (Part C) Edited by THOMAS L. JAMES AND NORMAN J. OPPENHEIMER VOLUME 240. Numerical Computer Methods (Part B) Edited by MICHAEL L. JOHNSON AND LUDWIG BRAND VOLUME 241. Retroviral Proteases Edited by LAWRENCE C. KUO AND JULES A. SHAFER VOLUME 242. Neoglycoconjugates (Part A) Edited by Y. C. LEE AND REIKO T. LEE VOLUME 243. Inorganic Microbial Sulfur Metabolism Edited by HARRY D. PECK, JR., AND JEAN LEGALL VOLUME 244. Proteolytic Enzymes: Serine and Cysteine Peptidases Edited by ALAN J. BARRETT VOLUME 245. Extracellular Matrix Components Edited by E. RUOSLAHTI AND E. ENGVALL VOLUME 246. Biochemical Spectroscopy Edited by KENNETH SAUER VOLUME 247. Neoglycoconjugates (Part B: Biomedical Applications) Edited by Y. C. LEE AND REIKO T. LEE VOLUME 248. Proteolytic Enzymes: Aspartic and Metallo Peptidases Edited by ALAN J. BARRETT VOLUME 249. Enzyme Kinetics and Mechanism (Part D: Developments in Enzyme Dynamics) Edited by DANIEL L. PURICH VOLUME 250. Lipid Modifications of Proteins Edited by PATRICK J. CASEY AND JANICE E. BUSS VOLUME 251. Biothiols (Part A: Monothiols and Dithiols, Protein Thiols, and Thiyl Radicals) Edited by LESTER PACKER VOLUME 252. Biothiols (Part B: Glutathione and Thioredoxin; Thiols in Signal Transduction and Gene Regulation) Edited by LESTER PACKER VOLUME 253. Adhesion of Microbial Pathogens Edited by RON J. DOYLE AND ITZHAK OFEK VOLUME 254. Oncogene Techniques Edited by PETER K. VOGT AND INDER M. VERMA VOLUME 255. Small GTPases and Their Regulators (Part A: Ras Family) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 256. Small GTPases and Their Regulators (Part B: Rho Family) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL

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VOLUME 257. Small GTPases and Their Regulators (Part C: Proteins Involved in Transport) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 258. Redox-Active Amino Acids in Biology Edited by JUDITH P. KLINMAN VOLUME 259. Energetics of Biological Macromolecules Edited by MICHAEL L. JOHNSON AND GARY K. ACKERS VOLUME 260. Mitochondrial Biogenesis and Genetics (Part A) Edited by GIUSEPPE M. ATTARDI AND ANNE CHOMYN VOLUME 261. Nuclear Magnetic Resonance and Nucleic Acids Edited by THOMAS L. JAMES VOLUME 262. DNA Replication Edited by JUDITH L. CAMPBELL VOLUME 263. Plasma Lipoproteins (Part C: Quantitation) Edited by WILLIAM A. BRADLEY, SANDRA H. GIANTURCO, AND JERE P. SEGREST VOLUME 264. Mitochondrial Biogenesis and Genetics (Part B) Edited by GIUSEPPE M. ATTARDI AND ANNE CHOMYN VOLUME 265. Cumulative Subject Index Volumes 228, 230–262 VOLUME 266. Computer Methods for Macromolecular Sequence Analysis Edited by RUSSELL F. DOOLITTLE VOLUME 267. Combinatorial Chemistry Edited by JOHN N. ABELSON VOLUME 268. Nitric Oxide (Part A: Sources and Detection of NO; NO Synthase) Edited by LESTER PACKER VOLUME 269. Nitric Oxide (Part B: Physiological and Pathological Processes) Edited by LESTER PACKER VOLUME 270. High Resolution Separation and Analysis of Biological Macromolecules (Part A: Fundamentals) Edited by BARRY L. KARGER AND WILLIAM S. HANCOCK VOLUME 271. High Resolution Separation and Analysis of Biological Macromolecules (Part B: Applications) Edited by BARRY L. KARGER AND WILLIAM S. HANCOCK VOLUME 272. Cytochrome P450 (Part B) Edited by ERIC F. JOHNSON AND MICHAEL R. WATERMAN VOLUME 273. RNA Polymerase and Associated Factors (Part A) Edited by SANKAR ADHYA VOLUME 274. RNA Polymerase and Associated Factors (Part B) Edited by SANKAR ADHYA


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VOLUME 275. Viral Polymerases and Related Proteins Edited by LAWRENCE C. KUO, DAVID B. OLSEN, AND STEVEN S. CARROLL VOLUME 276. Macromolecular Crystallography (Part A) Edited by CHARLES W. CARTER, JR., AND ROBERT M. SWEET VOLUME 277. Macromolecular Crystallography (Part B) Edited by CHARLES W. CARTER, JR., AND ROBERT M. SWEET VOLUME 278. Fluorescence Spectroscopy Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 279. Vitamins and Coenzymes (Part I) Edited by DONALD B. MCCORMICK, JOHN W. SUTTIE, AND CONRAD WAGNER VOLUME 280. Vitamins and Coenzymes (Part J) Edited by DONALD B. MCCORMICK, JOHN W. SUTTIE, AND CONRAD WAGNER VOLUME 281. Vitamins and Coenzymes (Part K) Edited by DONALD B. MCCORMICK, JOHN W. SUTTIE, AND CONRAD WAGNER VOLUME 282. Vitamins and Coenzymes (Part L) Edited by DONALD B. MCCORMICK, JOHN W. SUTTIE, AND CONRAD WAGNER VOLUME 283. Cell Cycle Control Edited by WILLIAM G. DUNPHY VOLUME 284. Lipases (Part A: Biotechnology) Edited by BYRON RUBIN AND EDWARD A. DENNIS VOLUME 285. Cumulative Subject Index Volumes 263, 264, 266–284, 286–289 VOLUME 286. Lipases (Part B: Enzyme Characterization and Utilization) Edited by BYRON RUBIN AND EDWARD A. DENNIS VOLUME 287. Chemokines Edited by RICHARD HORUK VOLUME 288. Chemokine Receptors Edited by RICHARD HORUK VOLUME 289. Solid Phase Peptide Synthesis Edited by GREGG B. FIELDS VOLUME 290. Molecular Chaperones Edited by GEORGE H. LORIMER AND THOMAS BALDWIN VOLUME 291. Caged Compounds Edited by GERARD MARRIOTT VOLUME 292. ABC Transporters: Biochemical, Cellular, and Molecular Aspects Edited by SURESH V. AMBUDKAR AND MICHAEL M. GOTTESMAN VOLUME 293. Ion Channels (Part B) Edited by P. MICHAEL CONN

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VOLUME 294. Ion Channels (Part C) Edited by P. MICHAEL CONN VOLUME 295. Energetics of Biological Macromolecules (Part B) Edited by GARY K. ACKERS AND MICHAEL L. JOHNSON VOLUME 296. Neurotransmitter Transporters Edited by SUSAN G. AMARA VOLUME 297. Photosynthesis: Molecular Biology of Energy Capture Edited by LEE MCINTOSH VOLUME 298. Molecular Motors and the Cytoskeleton (Part B) Edited by RICHARD B. VALLEE VOLUME 299. Oxidants and Antioxidants (Part A) Edited by LESTER PACKER VOLUME 300. Oxidants and Antioxidants (Part B) Edited by LESTER PACKER VOLUME 301. Nitric Oxide: Biological and Antioxidant Activities (Part C) Edited by LESTER PACKER VOLUME 302. Green Fluorescent Protein Edited by P. MICHAEL CONN VOLUME 303. cDNA Preparation and Display Edited by SHERMAN M. WEISSMAN VOLUME 304. Chromatin Edited by PAUL M. WASSARMAN AND ALAN P. WOLFFE VOLUME 305. Bioluminescence and Chemiluminescence (Part C) Edited by THOMAS O. BALDWIN AND MIRIAM M. ZIEGLER VOLUME 306. Expression of Recombinant Genes in Eukaryotic Systems Edited by JOSEPH C. GLORIOSO AND MARTIN C. SCHMIDT VOLUME 307. Confocal Microscopy Edited by P. MICHAEL CONN VOLUME 308. Enzyme Kinetics and Mechanism (Part E: Energetics of Enzyme Catalysis) Edited by DANIEL L. PURICH AND VERN L. SCHRAMM VOLUME 309. Amyloid, Prions, and Other Protein Aggregates Edited by RONALD WETZEL VOLUME 310. Biofilms Edited by RON J. DOYLE VOLUME 311. Sphingolipid Metabolism and Cell Signaling (Part A) Edited by ALFRED H. MERRILL, JR., AND YUSUF A. HANNUN


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VOLUME 312. Sphingolipid Metabolism and Cell Signaling (Part B) Edited by ALFRED H. MERRILL, JR., AND YUSUF A. HANNUN VOLUME 313. Antisense Technology (Part A: General Methods, Methods of Delivery, and RNA Studies) Edited by M. IAN PHILLIPS VOLUME 314. Antisense Technology (Part B: Applications) Edited by M. IAN PHILLIPS VOLUME 315. Vertebrate Phototransduction and the Visual Cycle (Part A) Edited by KRZYSZTOF PALCZEWSKI VOLUME 316. Vertebrate Phototransduction and the Visual Cycle (Part B) Edited by KRZYSZTOF PALCZEWSKI VOLUME 317. RNA–Ligand Interactions (Part A: Structural Biology Methods) Edited by DANIEL W. CELANDER AND JOHN N. ABELSON VOLUME 318. RNA–Ligand Interactions (Part B: Molecular Biology Methods) Edited by DANIEL W. CELANDER AND JOHN N. ABELSON VOLUME 319. Singlet Oxygen, UV-A, and Ozone Edited by LESTER PACKER AND HELMUT SIES VOLUME 320. Cumulative Subject Index Volumes 290–319 VOLUME 321. Numerical Computer Methods (Part C) Edited by MICHAEL L. JOHNSON AND LUDWIG BRAND VOLUME 322. Apoptosis Edited by JOHN C. REED VOLUME 323. Energetics of Biological Macromolecules (Part C) Edited by MICHAEL L. JOHNSON AND GARY K. ACKERS VOLUME 324. Branched-Chain Amino Acids (Part B) Edited by ROBERT A. HARRIS AND JOHN R. SOKATCH VOLUME 325. Regulators and Effectors of Small GTPases (Part D: Rho Family) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 326. Applications of Chimeric Genes and Hybrid Proteins (Part A: Gene Expression and Protein Purification) Edited by JEREMY THORNER, SCOTT D. EMR, AND JOHN N. ABELSON VOLUME 327. Applications of Chimeric Genes and Hybrid Proteins (Part B: Cell Biology and Physiology) Edited by JEREMY THORNER, SCOTT D. EMR, AND JOHN N. ABELSON VOLUME 328. Applications of Chimeric Genes and Hybrid Proteins (Part C: Protein–Protein Interactions and Genomics) Edited by JEREMY THORNER, SCOTT D. EMR, AND JOHN N. ABELSON

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VOLUME 329. Regulators and Effectors of Small GTPases (Part E: GTPases Involved in Vesicular Traffic) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 330. Hyperthermophilic Enzymes (Part A) Edited by MICHAEL W. W. ADAMS AND ROBERT M. KELLY VOLUME 331. Hyperthermophilic Enzymes (Part B) Edited by MICHAEL W. W. ADAMS AND ROBERT M. KELLY VOLUME 332. Regulators and Effectors of Small GTPases (Part F: Ras Family I) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 333. Regulators and Effectors of Small GTPases (Part G: Ras Family II) Edited by W. E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 334. Hyperthermophilic Enzymes (Part C) Edited by MICHAEL W. W. ADAMS AND ROBERT M. KELLY VOLUME 335. Flavonoids and Other Polyphenols Edited by LESTER PACKER VOLUME 336. Microbial Growth in Biofilms (Part A: Developmental and Molecular Biological Aspects) Edited by RON J. DOYLE VOLUME 337. Microbial Growth in Biofilms (Part B: Special Environments and Physicochemical Aspects) Edited by RON J. DOYLE VOLUME 338. Nuclear Magnetic Resonance of Biological Macromolecules (Part A) Edited by THOMAS L. JAMES, VOLKER DO¨TSCH, AND ULI SCHMITZ VOLUME 339. Nuclear Magnetic Resonance of Biological Macromolecules (Part B) Edited by THOMAS L. JAMES, VOLKER DO¨TSCH, AND ULI SCHMITZ VOLUME 340. Drug–Nucleic Acid Interactions Edited by JONATHAN B. CHAIRES AND MICHAEL J. WARING VOLUME 341. Ribonucleases (Part A) Edited by ALLEN W. NICHOLSON VOLUME 342. Ribonucleases (Part B) Edited by ALLEN W. NICHOLSON VOLUME 343. G Protein Pathways (Part A: Receptors) Edited by RAVI IYENGAR AND JOHN D. HILDEBRANDT VOLUME 344. G Protein Pathways (Part B: G Proteins and Their Regulators) Edited by RAVI IYENGAR AND JOHN D. HILDEBRANDT VOLUME 345. G Protein Pathways (Part C: Effector Mechanisms) Edited by RAVI IYENGAR AND JOHN D. HILDEBRANDT


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VOLUME 346. Gene Therapy Methods Edited by M. IAN PHILLIPS VOLUME 347. Protein Sensors and Reactive Oxygen Species (Part A: Selenoproteins and Thioredoxin) Edited by HELMUT SIES AND LESTER PACKER VOLUME 348. Protein Sensors and Reactive Oxygen Species (Part B: Thiol Enzymes and Proteins) Edited by HELMUT SIES AND LESTER PACKER VOLUME 349. Superoxide Dismutase Edited by LESTER PACKER VOLUME 350. Guide to Yeast Genetics and Molecular and Cell Biology (Part B) Edited by CHRISTINE GUTHRIE AND GERALD R. FINK VOLUME 351. Guide to Yeast Genetics and Molecular and Cell Biology (Part C) Edited by CHRISTINE GUTHRIE AND GERALD R. FINK VOLUME 352. Redox Cell Biology and Genetics (Part A) Edited by CHANDAN K. SEN AND LESTER PACKER VOLUME 353. Redox Cell Biology and Genetics (Part B) Edited by CHANDAN K. SEN AND LESTER PACKER VOLUME 354. Enzyme Kinetics and Mechanisms (Part F: Detection and Characterization of Enzyme Reaction Intermediates) Edited by DANIEL L. PURICH VOLUME 355. Cumulative Subject Index Volumes 321–354 VOLUME 356. Laser Capture Microscopy and Microdissection Edited by P. MICHAEL CONN VOLUME 357. Cytochrome P450, Part C Edited by ERIC F. JOHNSON AND MICHAEL R. WATERMAN VOLUME 358. Bacterial Pathogenesis (Part C: Identification, Regulation, and Function of Virulence Factors) Edited by VIRGINIA L. CLARK AND PATRIK M. BAVOIL VOLUME 359. Nitric Oxide (Part D) Edited by ENRIQUE CADENAS AND LESTER PACKER VOLUME 360. Biophotonics (Part A) Edited by GERARD MARRIOTT AND IAN PARKER VOLUME 361. Biophotonics (Part B) Edited by GERARD MARRIOTT AND IAN PARKER VOLUME 362. Recognition of Carbohydrates in Biological Systems (Part A) Edited by YUAN C. LEE AND REIKO T. LEE

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VOLUME 363. Recognition of Carbohydrates in Biological Systems (Part B) Edited by YUAN C. LEE AND REIKO T. LEE VOLUME 364. Nuclear Receptors Edited by DAVID W. RUSSELL AND DAVID J. MANGELSDORF VOLUME 365. Differentiation of Embryonic Stem Cells Edited by PAUL M. WASSAUMAN AND GORDON M. KELLER VOLUME 366. Protein Phosphatases Edited by SUSANNE KLUMPP AND JOSEF KRIEGLSTEIN VOLUME 367. Liposomes (Part A) Edited by NEJAT DU¨ZGU¨NES VOLUME 368. Macromolecular Crystallography (Part C) Edited by CHARLES W. CARTER, JR., AND ROBERT M. SWEET VOLUME 369. Combinational Chemistry (Part B) Edited by GUILLERMO A. MORALES AND BARRY A. BUNIN VOLUME 370. RNA Polymerases and Associated Factors (Part C) Edited by SANKAR L. ADHYA AND SUSAN GARGES VOLUME 371. RNA Polymerases and Associated Factors (Part D) Edited by SANKAR L. ADHYA AND SUSAN GARGES VOLUME 372. Liposomes (Part B) Edited by NEJAT DU¨ZGU¨NES VOLUME 373. Liposomes (Part C) Edited by NEJAT DU¨ZGU¨NES VOLUME 374. Macromolecular Crystallography (Part D) Edited by CHARLES W. CARTER, JR., AND ROBERT W. SWEET VOLUME 375. Chromatin and Chromatin Remodeling Enzymes (Part A) Edited by C. DAVID ALLIS AND CARL WU VOLUME 376. Chromatin and Chromatin Remodeling Enzymes (Part B) Edited by C. DAVID ALLIS AND CARL WU VOLUME 377. Chromatin and Chromatin Remodeling Enzymes (Part C) Edited by C. DAVID ALLIS AND CARL WU VOLUME 378. Quinones and Quinone Enzymes (Part A) Edited by HELMUT SIES AND LESTER PACKER VOLUME 379. Energetics of Biological Macromolecules (Part D) Edited by JO M. HOLT, MICHAEL L. JOHNSON, AND GARY K. ACKERS VOLUME 380. Energetics of Biological Macromolecules (Part E) Edited by JO M. HOLT, MICHAEL L. JOHNSON, AND GARY K. ACKERS VOLUME 381. Oxygen Sensing Edited by CHANDAN K. SEN AND GREGG L. SEMENZA


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VOLUME 382. Quinones and Quinone Enzymes (Part B) Edited by HELMUT SIES AND LESTER PACKER VOLUME 383. Numerical Computer Methods (Part D) Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 384. Numerical Computer Methods (Part E) Edited by LUDWIG BRAND AND MICHAEL L. JOHNSON VOLUME 385. Imaging in Biological Research (Part A) Edited by P. MICHAEL CONN VOLUME 386. Imaging in Biological Research (Part B) Edited by P. MICHAEL CONN VOLUME 387. Liposomes (Part D) Edited by NEJAT DU¨ZGU¨NES VOLUME 388. Protein Engineering Edited by DAN E. ROBERTSON AND JOSEPH P. NOEL VOLUME 389. Regulators of G-Protein Signaling (Part A) Edited by DAVID P. SIDEROVSKI VOLUME 390. Regulators of G-Protein Signaling (Part B) Edited by DAVID P. SIDEROVSKI VOLUME 391. Liposomes (Part E) Edited by NEJAT DU¨ZGU¨NES VOLUME 392. RNA Interference Edited by ENGELKE ROSSI VOLUME 393. Circadian Rhythms Edited by MICHAEL W. YOUNG VOLUME 394. Nuclear Magnetic Resonance of Biological Macromolecules (Part C) Edited by THOMAS L. JAMES VOLUME 395. Producing the Biochemical Data (Part B) Edited by ELIZABETH A. ZIMMER AND ERIC H. ROALSON VOLUME 396. Nitric Oxide (Part E) Edited by LESTER PACKER AND ENRIQUE CADENAS VOLUME 397. Environmental Microbiology Edited by JARED R. LEADBETTER VOLUME 398. Ubiquitin and Protein Degradation (Part A) Edited by RAYMOND J. DESHAIES VOLUME 399. Ubiquitin and Protein Degradation (Part B) Edited by RAYMOND J. DESHAIES VOLUME 400. Phase II Conjugation Enzymes and Transport Systems Edited by HELMUT SIES AND LESTER PACKER

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VOLUME 401. Glutathione Transferases and Gamma Glutamyl Transpeptidases Edited by HELMUT SIES AND LESTER PACKER VOLUME 402. Biological Mass Spectrometry Edited by A. L. BURLINGAME VOLUME 403. GTPases Regulating Membrane Targeting and Fusion Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 404. GTPases Regulating Membrane Dynamics Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 405. Mass Spectrometry: Modified Proteins and Glycoconjugates Edited by A. L. BURLINGAME VOLUME 406. Regulators and Effectors of Small GTPases: Rho Family Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 407. Regulators and Effectors of Small GTPases: Ras Family Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL VOLUME 408. DNA Repair (Part A) Edited by JUDITH L. CAMPBELL AND PAUL MODRICH VOLUME 409. DNA Repair (Part B) Edited by JUDITH L. CAMPBELL AND PAUL MODRICH VOLUME 410. DNA Microarrays (Part A: Array Platforms and Web-Bench Protocols) Edited by ALAN KIMMEL AND BRIAN OLIVER VOLUME 411. DNA Microarrays (Part B: Databases and Statistics) Edited by ALAN KIMMEL AND BRIAN OLIVER VOLUME 412. Amyloid, Prions, and Other Protein Aggregates (Part B) Edited by INDU KHETERPAL AND RONALD WETZEL VOLUME 413. Amyloid, Prions, and Other Protein Aggregates (Part C) Edited by INDU KHETERPAL AND RONALD WETZEL VOLUME 414. Measuring Biological Responses with Automated Microscopy Edited by JAMES INGLESE VOLUME 415. Glycobiology Edited by MINORU FUKUDA VOLUME 416. Glycomics Edited by MINORU FUKUDA VOLUME 417. Functional Glycomics Edited by MINORU FUKUDA VOLUME 418. Embryonic Stem Cells Edited by IRINA KLIMANSKAYA AND ROBERT LANZA


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VOLUME 419. Adult Stem Cells Edited by IRINA KLIMANSKAYA AND ROBERT LANZA VOLUME 420. Stem Cell Tools and Other Experimental Protocols Edited by IRINA KLIMANSKAYA AND ROBERT LANZA VOLUME 421. Advanced Bacterial Genetics: Use of Transposons and Phage for Genomic Engineering Edited by KELLY T. HUGHES VOLUME 422. Two-Component Signaling Systems, Part A Edited by MELVIN I. SIMON, BRIAN R. CRANE, AND ALEXANDRINE CRANE VOLUME 423. Two-Component Signaling Systems, Part B Edited by MELVIN I. SIMON, BRIAN R. CRANE, AND ALEXANDRINE CRANE VOLUME 424. RNA Editing Edited by JONATHA M. GOTT VOLUME 425. RNA Modification Edited by JONATHA M. GOTT VOLUME 426. Integrins Edited by DAVID CHERESH VOLUME 427. MicroRNA Methods Edited by JOHN J. ROSSI VOLUME 428. Osmosensing and Osmosignaling Edited by HELMUT SIES AND DIETER HAUSSINGER VOLUME 429. Translation Initiation: Extract Systems and Molecular Genetics Edited by JON LORSCH VOLUME 430. Translation Initiation: Reconstituted Systems and Biophysical Methods Edited by JON LORSCH VOLUME 431. Translation Initiation: Cell Biology, High-Throughput and Chemical-Based Approaches Edited by JON LORSCH VOLUME 432. Lipidomics and Bioactive Lipids: Mass-Spectrometry–Based Lipid Analysis Edited by H. ALEX BROWN VOLUME 433. Lipidomics and Bioactive Lipids: Specialized Analytical Methods and Lipids in Disease Edited by H. ALEX BROWN VOLUME 434. Lipidomics and Bioactive Lipids: Lipids and Cell Signaling Edited by H. ALEX BROWN

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VOLUME 435. Oxygen Biology and Hypoxia Edited by HELMUT SIES AND BERNHARD BRU¨NE VOLUME 436. Globins and Other Nitric Oxide-Reactive Protiens (Part A) Edited by ROBERT K. POOLE VOLUME 437. Globins and Other Nitric Oxide-Reactive Protiens (Part B) Edited by ROBERT K. POOLE VOLUME 438. Small GTPases in Disease, Part A (in preparation) Edited by WILLIAM E. BALCH, CHANNING J. DER, AND ALAN HALL

S E C T I O N

O N E

NITRIC OXIDE-METABOLISING AND DETOXIFYING ENZYMES


C H A P T E R

O N E

Structural Studies on Flavodiiron Proteins Joaõ B. Vicente, Maria Armeńia Carrondo, Miguel Teixeira, and Carlos Frazaõ Contents 4 4

1. Introduction 2. Crystallization of Flavodiiron Proteins 3. Diffraction Data Collection, Structure Determination, and Refinement 4. Overall Description of Structures 4.1. Metallo-b-lactamase-like domain of flavodiiron proteins 4.2. Flavodoxin domain of flavodiiron proteins 4.3. Features of the nonheme diiron center 4.4. Features of the flavin mononucleotide moiety 5. Conclusion References

7 8 9 11 11 14 16 17

Abstract Crystallographic studies on flavodiiron proteins (FDPs) have revealed that the common sequence core (400 residues) that defines this protein family comprises two structural domains. The N-terminal domain (of approximately 250 residues) displays a metallo-b-lactamase-like-fold, being indeed structurally homologous to b-lactamases and glyoxalases, despite the poor sequence similarity. Whereas b-lactamases have mono- or dizinc sites and glyoxalases a mixed iron–zinc site, the lactamase domain of FDPs harbors a nonheme diiron center with carboxylate and histidine residues as ligands, assigned as the active site of NO and/or O2 reduction. The C-terminal domain of FDPs is characterized by a flavodoxin-like fold, homologous to short-chain flavodoxins, and harbors a flavin mononucleotide moiety, stabilized by van der Waals interactions and a number of hydrogen bonds. Structures of FDPs obtained in different conditions and oxidation states display some heterogeneities, mostly at the diiron site, but still fail to provide unequivocal evidence for some pending questions regarding the substrate activation mechanism of FDPs, namely the preference for either Instituto de Tecnologia Quı´mica e Biolo´gica, Universidade Nova de Lisboa, Oeiras, Portugal Methods in Enzymology, Volume 437 ISSN 0076-6879, DOI: 10.1016/S0076-6879(07)37001-8

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2008 Elsevier Inc. All rights reserved.

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4

Joaõ B. Vicente et al.

substrate (NO or oxygen) observed in different members of this protein family. More structural studies are therefore required to achieve a deeper understanding on these matters.

1. Introduction Flavodiiron proteins (FDPs) are widespread in Archaea and Bacteria (Wasserfallen et al., 1998), mostly anaerobic organisms, extending also to the Eukarya, namely in a few pathogenic protozoa (Andersson et al., 2003, 2006; Sarti et al., 2004). Currently, accumulated data suggest alternative or complementary roles in the detoxification of NO and/or oxygen (toxic for anaerobes). Initial studies on Desulfovibrio gigas rubredoxin:oxygen oxidoreductase (Dg_ROO, the designation given to the FDP of this organism) indicated a role in the protection against oxygen toxicity to this strict anaerobe (Chen et al., 1993), as D. gigas was considered at the time. Subsequent reports focusing on Escherichia coli FDP (named flavorubredoxin, FlRd, NorV) have demonstrated a function of this protein in the anaerobic protection against NO-derived stress (Gardner et al., 2002; Gomes et al., 2002b; Justino et al., 2005). Several members of this protein family were later shown to be endowed with both NO and oxygen reductase activities, with different substrate specificities, as judged by the comparative affinities (Rodrigues et al., 2006; Silaghi-Dumitrescu et al., 2003, 2005b). More recent reports on FDPs from methanogenic archaea suggest an exclusive role in oxygen reduction (Seedorf et al., 2004, 2007), accepting electrons directly from the organic cofactor F420H2 (ubiquitous in methanogens). Although preliminary studies on several FDPs revealed important features common to the protein family, it was not until the first crystallographic structure of an FDP was solved that the nature of the active site was unraveled (Frazaõ et al., 2000). Indeed, the structure of Dg_ROO revealed several aspects (described in detail later), which shed some light on the functional properties of FDPs. Although the subsequent FDP structures confirmed many of the Dg_ROO structural features, some heterogeneities at the active site are noteworthy. Several structural studies also aimed at understanding the structural conditionals that confer FDPs a selectivity for either substrate (NO or oxygen).

2. Crystallization of Flavodiiron Proteins The flavodiiron proteins for which X-ray crystallographic structures have been solved were isolated as described in Vicente et al. (2007), except the FDP from Thermotoga maritima (Tm_FDP). Whereas D. gigas rubredoxin:oxygen oxidoreductase (Dg_ROO) was isolated from its source

Structural Studies on Flavodiiron Proteins

5

organism (Chen et al., 1993), the remainder of FDPs were overexpressed heterologously in E. coli (Seedorf et al., 2007; Silaghi-Dumitrescu et al., 2005a). Tm_FDP was overexpressed in E. coli and purified by a highthroughput automated method (DiDonato et al., 2004). Flavodiiron proteins were crystallized essentially by two methods: (i) the batch method, using Zinc acetate as precipitant, or (ii) the vapor diffusion method, using alcohol type precipitants (PEGs or MPD). When crystal cryostabilization was necessary for data collection, it was performed by incremental additions of the stabilizing agent. Dg_ROO crystals, obtained with PEG 1 to 6K as precipitating agents in a wide range of pH (5–9) (Frazaõ et al., 1999), appeared within 1 day at room temperature in large numbers, amidst a gelatinous precipitate. To prevent the formation of this gel and to control the number of crystals, crystallization trials were performed at 277K, and also adding the detergent SB12 (N-dodecyl-N,N-dimethyl-3-ammonio-1-propansulfonate). Crystals obtained at both temperatures had similar parallelepiped shapes, although their diffraction quality changed significantly. Whereas the diffraction of crystals obtained at 277K exhibited twinning statistics, those obtained at room temperature developed as single crystals. Crystals that allowed solving the Dg_ROO structure were typically obtained as follows. A sitting drop composed by 3 ml of 10 mg/ml protein solution and an equal volume of precipitant solution [PEG 6K 10% (v/v), 100 mM Tris-maleic acid, pH 6.0] was equilibrated against 500 ml of the same solution. Orange-brown parallelepiped-shaped crystals grew within approximately 3 weeks up to 0.2 to 0.4 mm in length and were cryostabilized by adding to 20-ml drops of precipitant solution containing the crystals, small amounts of precipitant solution (initially 0.2 ml each) complemented with glycerol (25%, v/v), in the cold room, up to a final glycerol concentration of 25% (v/v). After stabilization, crystals were flash frozen in a nitrogen stream and diffraction data collected. Crystals belonged to space group P21212 (cell dimensions ˚ , b ¼ 101.2 A ˚ , and c ¼ 90.8 A˚), diffracted to 2.5 A ˚ resolution a ¼ 98.2 A and contained two molecules (one dimer) in the asymmetric unit. The flavodiiron protein from Moorella thermoacetica (Mt_FDP) was crystallized (Silaghi-Dumitrescu et al., 2005a) by the batch method in melting point capillaries (at room temperature) by mixing 10 ml of 1 mM (42 mg/ml) oxidized (as-isolated) (Mt_FDP_ox) solution to an equal volume of precipitant solution. Two slightly different precipitant solutions yielded crystals, 200 mM zinc acetate, 5% 2-propanol, 50 mM sodium cacodylate, pH 6.5, and 200 mM zinc acetate, 10% PEG 3000, 100 mM sodium acetate, pH 4.5. Relatively large crystals (0.2 0.2 0.5 mm) formed within 7 to 10 days, endowed with diffraction quality, were cryostabilized by soaking the crystals in mother liquor [1:1 (v/v) 25 mM MOPS (pH 7.3):precipitant] containing step increments of 5% ethylene glycol up to 20%, with each step taking 20 min. Crystals of

6


reduced Mt_FDP (Mt_FDP_red) were obtained in an anaerobic glove box by the addition of sodium dithionite powder to as-isolated Mt_FDP crystals [in 200 ml of mother liquor containing 20% (v/v) ethylene glycol and the pH 6.5 precipitant], resulting in crystals changing from orange-brown to colorless within a few minutes. Crystals of Mt_FDP reoxidized with NO (Mt_FDP_NO) were obtained starting from the reduced crystals prepared as described earlier. These crystals were back-soaked into the same mother liquor (though lacking sodium dithionite) and further incubated with the NO-releasing compound DEA-NONOate (in powder), leading to a fast color change back to the original orange-brown, attesting the Mt_FDP reoxidation by NO. The crystals obtained in three states (oxidized, reduced, and reduced þ NO reacted) were flash frozen in liquid nitrogen; all belonged to space group P43212. The cell dimensions and resolution of ˚ , c ¼ 276.7 A ˚ , and 3.0 A ˚ resolution for diffraction data were a ¼ b ¼ 159.6 A ˚ ˚ crystals of Mt_FDP_ox; a ¼ b ¼ 159.6 A, c ¼ 278.1 A, and 2.8 A˚ resolution ˚ , c ¼ 279.1 A ˚ , and 2.8 A˚ for Mt_FDP_red crystals; and a ¼ b ¼ 159.6 A resolution for Mt_FDP_NO crystals. Moreover, all the crystals were shown to contain four molecules (an assembly of two homodimers) in the asymmetric unit. Crystals of T. maritima FDP were obtained as part of a high-throughput automated method for protein production, crystallization, and structure determination, focusing on the proteome of this organism (DiDonato et al., 2004). Because the structure of Tm_FDP was deposited in the PDB but not published, information on the crystallization procedure is still scarce (PDB entry 1VME). Crystals obtained at 277 K in sitting nanodrops, by the vapor diffusion method, using 35.0% MPD, 0.1 M acetate (pH 4.5), belonged to space group P21 with cell dimensions a ¼ 55.24 A˚, b ¼ ˚ , c ¼ 90.13, and b ¼ 95.43 , diffracted to 1.8 A˚ and contained 95.83 A two molecules (one dimer) in the asymmetric unit. Contrary to other FDPs for which the crystallographic structure was solved, the FprA from Methanothermobacter marburgensis (Mm_FprA) was purified and crystallized only in anaerobic conditions (in an anaerobic glove box) (Seedorf et al., 2007). Different crystallization conditions yielded three crystal forms, grown in drops up to 20 ml using 20 mg/ml Mm_FprA solution and precipitant solution, all three of them obtained by the hanging drop vapor diffusion method, at 283 K, and in the presence of 1 mM dithiothreitol. Crystals obtained with 0.2 M ammonium sulfate, 0.1 M MES/KOH, pH 6.5, and 16–22% PEG MME 5000 displayed two different monoclinic P21 crystal forms, one containing eight molecules in the asym˚ , b ¼ 123.1 A˚, c ¼ 135.9 A ˚ , and metric unit, with cell dimensions a ¼ 97.8 A ˚ b¼ 103.4 , diffracting to 2.25 A resolution; and the second P21 form containing four molecules in the asymmetric unit, with cell dimensions ˚ , b ¼ 120.9 A ˚ , c ¼ 92.7 A ˚ , and b¼ 110.4 , diffracting to 1.7 A ˚ a ¼ 73.7 A resolution. The third crystal form resulted from a lower concentration of


7

PEG MME 5000 (8–16%), compensated with 15% glycerol, yielding tetragonal crystals in space group P43212, displaying four molecules in the ˚ , c ¼ 450.4 A ˚ , that asymmetric unit, with cell dimensions a ¼ b ¼ 88.7 A ˚ diffracted to 2.25 A resolution. Differences in data sets also resulted from the distinct ways the crystals were manipulated. The first crystal form (P21 with eight molecules in the a.u.) was measured after flash cooling the crystals inside the anaerobic chamber (grown in the presence of F420H2, the electron donor for Mm_FprA) in liquid nitrogen, corresponding to reduced Mm_FprA. The second and third crystal forms (P21 with four molecules in the a.u., and P43212) were flash frozen in a nitrogen gas stream after being exposed to air for minutes at 191 K, yielding presumably oxidized forms of Mm_Fpra.

3. Diffraction Data Collection, Structure Determination, and Refinement Two diffraction data sets of Dg_ROO were collected at cryogenic temperatures, a four-wavelength MAD data set around the Fe absorption ˚ resolution at beam line BM14 of ESRF (Frazaõ et al., 2000) edge to 2.7 A ˚ resolution at beam line X11 of DESY, and a single wavelength set to 2.5 A EMBL Hamburg outstation (Frazaõ et al., 1999). Diffraction intensities were measured with DENZO and scaled together with SCALEPACK of the HKL suite (Otwinowski and Minor, 1997). MADSYS (Hendrickson and Ogata, 1997) extracted F A values from MAD data which were used in SHELXS-97 (Sheldrick, 1990) to locate three out of the four Fe sites. The sites were further refined with SHARP (De La Fortelle and Bricogne, 1997), which found the fourth site from residual maps and included it in the phase improvement process using SOLOMON (Solomon et al., 2000), to ˚ . The polypeptide chain was modeled in the an overall f.o.m. of 0.95 at 2.7 A experimental maps followed by residues assignment using XTALVIEW (McRee, 1992). The refinement to 2.5 A˚ of the two molecules in the a.u. proceeded using SHELXL (Sheldrick and Schneider, 1997), restraining to their common values homologous 1–4 dihedral distances and atomic displacement parameters (a.d.p.s) of homologous atoms, and of waters with equivalent H-bonds to each monomer. The diiron centers and their ligating residues were restrained to a common geometry without target values. A residual lobe of electron density found close to the diiron site in different Fourier maps was initially assigned to a water molecule, but its refinement led to too low a.d.p.s. Taking into account the crystallization conditions it was then tentatively assigned to a dioxygen molecule. Tm_FDP diffraction data were obtained at 100 K at SSRL on beam line 9–2. Intensities were measured with MOSFLM (Leslie, 2006) and scaled together using SCALA (Evans, 2006) of the CCP4 suite (Collaborative

8


Computational Project Number 4, 1994). The phase problem was solved by MAD using a selenium-derivatized protein crystal. The structure was refined using the maximum likelihood target in REFMAC with six TLS groups ( Winn et al., 2003). Mt_FDP diffraction data were acquired at 98 K at the Advanced Light Source, Berkeley, on beam line 5.0.2. The molecular replacement solution by CNS (Brunger et al., 1998), using the Dg_ROO structure truncated to poly-Ala as the search model, was corroborated by the SAD solution of the iron sites using highly redundant (760 degrees) diffraction data, collected at the University of Georgia on a Rigaku Ru-200 rotating anode equipped with Osmic focusing mirrors and a R-axisIIc imaging plate detector. The graphical software O ( Jones et al., 1991) was used for model edition and completion and CNS (Brunger et al., 1998) for structure refinement, using the four-fold noncrystallographic symmetry operators to improve the quality of the electron density maps. Mm_FprA diffraction data were collected at cryo-temperature using beam line X10SA of the Swiss-Light-Source. Diffraction data were processed and scaled using the HKL (Otwinowski and Minor, 1997) and XDS (Kabsch, 1993) packages. The structure of active oxidized Mm_FprA was solved by molecular replacement with EPMR (Kissinger et al., 1999) using the Mt_FDP structure as the search model (the Dg_ROO model gave less reliable results). Phases for the two other crystal forms of Mm_FprA were obtained by molecular replacement using the active oxidized model. Refinement and model edition were performed with CNS (Brunger et al., 1998) and O ( Jones et al., 1991), making use of the noncrystallographic symmetry relationships. Refinement was completed with REFMAC (Murshudov et al., 1997) using the TLS option (each monomer treated as a separate TLS group), maximum likelihood minimization, and isotropic a.d.p.s refinement.

4. Overall Description of Structures Flavodiiron proteins have been isolated as functional homodimers and homotetramers (Vicente et al., 2007), which have been confirmed by the available X-ray structures (Frazaõ et al., 2000; Seedorf et al., 2007; SilaghiDumitrescu et al., 2005a). The homodimeric structure (Fig. 1.1) results from a ‘‘head-to-tail’’ arrangement of two monomers where approximately 12% of the possible solvent-accessible surface becomes occluded (Frazaõ et al., 2000; Seedorf et al., 2007). The tetrameric arrangement is composed by a loose dimer of two such dimers, where a further 9.5% of otherwise solvent-accessible surface becomes occluded (Seedorf et al., 2007). The available structures show similar three-dimensional arrangements with


9

Figure 1.1 Three-dimensional structure of Desulfovibrio gigas rubredoxin:oxygen oxidoreductase. Solvent-accessible transparent surface and ribbon representation of the ‘‘head-to-tail’’ homodimeric arrangement of D. gigas rubredoxin:oxygen oxidoreductase (Dg__ROO, PDB code 1E5D), the first flavodiiron protein whose crystallographic structure was solved. Each monomer is composed of two structural domains: the N-terminal domain with a metallo-b-lactamase-like fold is represented in lighter colors, whereas the C-terminal domain with a flavodoxin-like fold is represented in darker colors. The head-to-tail homodimeric structure places the nonheme diiron center active site (golden spheres) of each monomer in close proximity with the FMN moiety (yellow sticks) of the other monomer.

˚ upon superposition of the overall monomer Ca r.m.s.d.s within 1.1 to 1.8 A structures from four different species. The FDP monomers are composed of two different structural domains: an N-terminal domain with a metallo-blactamase-like fold and a C-terminal domain showing a flavodoxin-like fold.

4.1. Metallo-b-lactamase-like domain of flavodiiron proteins The lactamase N-terminal domain (up to residue 249, using the Dg_ROO numbering) folds in a abba sandwich, with the two inner b sheets being surrounded by two sets of three solvent-exposed a helices. Despite the low sequence identity between the lactamase domain of FDPs and other lactamase-like proteins (Gomes et al., 2002a), its overall fold (Fig. 1.2) is structurally very similar to the folds found in both class B Zn-b-lactamases (Carfi et al., 1995) (that hydrolyze penicillin) and human glyoxalase (Cameron et al., 1999) (that convert 2-oxoaldehydes into the corresponding 2-hydroxycarboxylic acids). Another striking difference between the lactamase domain of FDPs and other lactamase-like proteins concerns the nature of the metal active site. FDPs were found to harbor a nonheme diiron center (with histidine and carboxylate residues as ligands), whereas lactamases contain a mono- or a dizinc center and glyoxalases bind a mixed iron–zinc

10


A N

C B N

C C

N

C

Figure 1.2 Topology diagrams of the N-terminal domain of flavodiiron proteins and other lactamase-like folds.Topology diagrams of the structurally homologous lactamase domains found in (A) glyoxalases (first domain of human glyoxalase II, PDB entry 1QH5) (Cameron et al., 1999); (B) Zn-binding metallo-b-lactamases (Stenotrophomonas maltophilia, PDB entry 1SML) (Ullah et al., 1998); and (C) the N-terminal domain of Desulfovibrio gigas rubredoxin:oxygen oxidoreductase, PDB entry 1E5D) (Frazaõ et al., 2000). Circles depict helices, triangles depict b chains, and N and C represent the peptide chain direction. The simplest common secondary structure, found in glyoxalase, is represented in dark gray, light gray displays additional secondary structure elements present in metallo-b-lactamases, and white depicts those unique to FDPs.

binuclear center (Cameron et al., 1999; Zang et al., 2001). In FDPs the diiron site is located within a shallow groove in a cavity at the interface of the two sheets and is surrounded by ab loops and the C-terminal domain from the other monomer (which is made possible by the ‘‘head-to-tail’’ dimeric arrangement). Whereas the substrates of lactamases and glyoxalase are bulky and require large cavities near the binuclear metal centers, the FDPs substrates (NO and/or O2) are much smaller and can reach the diiron site, despite being covered by an additional two-stranded b sheet.


11

4.2. Flavodoxin domain of flavodiiron proteins The C-terminal domain of FDPs (from residue 250, Dg_ROO numbering) displays the typical aba flavodoxin fold, where a flavin mononucleotide (FMN) moiety is embedded that allegedly acts as the electron donor to the diiron site, where reduction of NO and/or oxygen takes place. Within a ˚ away from the diiron center, too far single monomer, the flavin is 35 A to be considered for effective electron transfer between the two redox sites. This observation highlights the relevance of the quaternary structure common to FDPs, in which the minimal operative unit is thought to be the ‘‘head-to-tail’’ homodimer. This arrangement places the diiron ˚ ) with the FMN moiety center of each monomer in close contact (6 A from the opposing monomer, yielding two independent catalytic sites per homodimer. Despite the significant structural similarity between the flavodoxin domain of FDPs and short-chain flavodoxins [comparable to the homology among flavodoxins themselves (Frazaõ et al., 2000)], an important difference is that the FMN reduction potentials in FDPs are higher than those in flavodoxins (see e.g., Gomes et al., 1997; Ludwig et al., 1997; Silaghi-Dumitrescu et al., 2003; Vicente and Teixeira, 2005), an observation that has been assigned to the relative excess of basic versus acidic residues surrounding the FMN isoalloxazine ring in FDPs.

4.3. Features of the nonheme diiron center Since the nonheme diiron center has been assigned as the active site of NO/O2 reduction in flavodiiron proteins (Gomes et al., 2002b; Silaghi-Dumitrescu et al., 2003), special emphasis has been given to analysis of the structural and functional features of this metal center. One obvious aspect is the pursuit of an understanding of the molecular basis for the selectivity of each studied FDP for either substrate (NO and/or O2). Although a definitive answer is far from being attained, some heterogeneities at the FDPs diiron centers (whose overall structure is globally conserved) are noteworthy. Whereas in Dg_ROO, Mt_FDP, and Tm_FDP the diiron center geometry is retained within the independent molecules in the a.u.s for the various available oxidation states, in Mm_FprA the diiron center differs significantly not only among the different oxidation states, but also between independent molecules of the same asymmetric units (by alternative conformations within a monomer). As a common feature in most structures, each iron is coordinated by two imidazole nitrogens, two oxygens from carboxylate residues, and a bridging solvent molecule, assigned as a putative m-oxo-, m-hydroxyl-, or m-aquo-ligand, since resolutions of the structures do not allow determination of its protonation state. However, in Mm_FprA, only 1 out of 16 independent structures deposited reports a bridging m-oxygen atom in the structure at the highest resolution. Interestingly, the bridging moiety appears to be redox insensitive, as the

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reduced structure of Mt_FDP still holds it, unlike other nonheme diiron centers that lose and regain the bridge throughout the oxygen-activating reaction cycle (Solomon et al., 2000). Considering the proximal iron the one closer to FMN (denoting the other as distal iron) and using Dg_ROO numbering (also throughout the text), the proximal iron is coordinated by residues His79, His146, and Glu81, and the distal iron is coordinated by His84, His226, and Asp83, with Asp165 acting as a bridging ligand for both irons. The ligands herein described are almost strictly conserved in this protein family, with few exceptions observed in FDPs encoded in the genomes of some cyanobacteria, which encode other copies of FDPs within the same genome, that retain the conserved ligands. A fifth coordination position remains open in each iron (somewhat trans to His79 and His84, parallel to each other), where a substrate is prone to bind. The exception to the general coordination observed in most FDPs is precisely found in the structure of Dg_ROO (Frazaõ et al., 2000), where the His84 ligand is displaced from the center, being replaced as a ligand by a solvent water molecule. This is because of the unique side chain conformation of His84, which makes a trans chi-1 angle, instead of the gauche(-) conformation found in other FDPs, in lactamases (Ullah et al., 1998), and in human glyoxalase (Cameron et al., 1999). A closer look at the structures allowed finding a structural feature that accounts for this heterogeneity: one H-bond between the His84 imidazole ND1 atom and the neighboring Asp225 carboxylate. The equivalent position to Asp225 in other FDP sequences (survey of 150 sequences) is not conserved, but varies among Ser (50%), Gly (34%), Asp (8%), and Ala (6%). From the available crystal structures, only Dg_ROO has an Asp in this position, which can account for its unique coordination, contrasting with the other FDPs, where the replacing residues are not able to make the same H-bond that ‘‘pulls’’ His84 away from the center in Dg_ROO. A Gly residue is in the equivalent position in Tm_FDP and a Ser in Mt_FDP, whose OG atom is too far to make the H-bond. Although these observations present a structural justification for the ligand heterogeneity between Dg_ROO and the other FDPs, there are still no clues for the structural properties that determine the specificity of FDPs for either substrate. To infer functional properties from the structural studies, it is important to attempt trapping reaction intermediates; therefore, structures of FDPs were obtained in different oxidation states, except Dg_ROO and Tm_FDP, which were reported only in the as-isolated oxidized state. Whereas Mt_FDP was crystallized in the oxidized form and then the crystals were reduced and reoxidized with an NO releaser, the Mm_FprA was crystallized anaerobically in the reduced form and oxidized upon O2 exposure before the crystal flash-cooling procedure.

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FMN

E81

FMN

E81

D83

D83 H226

D228 H146

H146 H79

H84 D165

D225

H79

H84

D225

D165

Figure 1.3 Structural features of the nonheme diiron active site of flavodiiron proteins. Stereo representation (cross-eyes) of the superposition of the diiron center from Desulfovibrio gigas rubredoxin:oxygen oxidoreductase (Dg__ROO) is shown in black (with residues numbering), whereas that of Thermotoga maritima flavodiiron protein (Tm_FDP) is represented in light gray.The two selected structures depict the heterogeneity of the active sites in the structures of FDPs, related with a conserved histidine residue (His84 in Dg__ROO), which is a ligand of the distal iron in most FDPs, but in Dg__ROO is replaced by a water molecule and appears displaced from the center, making a H-bond with Asp225 (dark gray). InTm_FDP, position 225 is a Gly residue (white symbol).

One evident observation (and possibly the most significant one) upon comparing the oxidized and reduced structures of Mt_FDP is that the latter has a larger active site cavity, being even able to harbor one ethylene glycol molecule ‘‘above’’ the diiron center, where an oxygen molecule is found in the oxidized structure, and one solvent water molecule in the NO-reoxidized structure. The aforementioned molecules located above the diiron center were refined at noncovalent bound distances to the iron atoms, on average at 2.8 A˚ among the four independent structures, to be compared with the typical 1.84 A˚ for Fe-O2 distances in high-resolution structures in the Cambridge Structural Database (Frazaõ et al., 2000). Contrasting with the low degree of structural variations in the different oxidation states of Mt_FDP, Mm_FprA structures display large differences among the reported crystal forms. Upon oxidation by oxygen exposure of the crystals obtained in anaerobic conditions, the diiron center becomes disrupted. Whereas the distal iron maintains its coordination sphere, the proximal iron is significantly displaced and its former ligands are displaced from the original conformation. Particularly, Glu81 is rotated away from the proximal position and forms a new remote metal-binding site, along ˚ from the distal with His26 and His267 (Mm_FprA numbering), at 7 A iron. Moreover, a third putative metal-binding site at the protein surface is reported, coordinated by His151, Asp320 (Mm_FprA residues numbering), and a water molecule. This displacement of the proximal iron is accompanied by a large conformational arrangement of the loop between residues 148–151, designated as ‘‘switch loop,’’ increasing the accessibility of

14


the redox centers. The resulting structure was therefore called the ‘‘inactive oxidized state.’’ As an intermediate situation between the as-isolated ‘‘active reduced’’ Mm_FprA structure and the ‘‘inactive oxidized state,’’ oxidation of the monoclinic crystals yielded an integral diiron center, although displaying a mixed conformation of the switch loop: 0.6 occupancy of the closed conformation found in the ‘‘reduced state’’ (and all the other FDP structures) and 0.4 occupancy of the open conformation observed in the ‘‘inactive oxidized state.’’ This ‘‘intermediate’’ structure was designated by the ‘‘active oxidized state.’’ Another observation regarding the oxidized structures is that the inactive open loop conformation swaps into trans the otherwise cis peptide bond Leu145-His146 (a nonproline cis bond also present in Dg_ROO and Tm_FDP, but not reported in Mt_FDP), which is necessary to project the imidazolyl ring toward the proximal iron. The four independent molecules in the asymmetric unit of the active oxidized state display a heterogeneously populated catalytic cavity, harboring either a diatomic molecule or a sulfate together with a monoatomic m-ligand to the diiron. The three different redox and conformational states of Mm_FprA may represent reaction intermediates with mechanistic implications. In the absence of reducing equivalents to feed the catalytic cycle of oxygen reduction by Mm_FprA, it appears that the enzyme may become inactivated upon prolonged oxygen exposure, still being unclear whether this inactivation is reversible. Notably, although the proximal iron is displaced away from its original location, it may still be bound to the protein in the alternative putative metal-binding sites aforementioned. Because the other structures were obtained in aerobic conditions with integer diiron centers, it is presently uncertain whether the observations regarding Mm_FprA are relevant to other FDPs.

4.4. Features of the flavin mononucleotide moiety Despite the poor sequence similarity between the C-terminal domain of flavodiiron proteins and short-chain flavodoxins, their structural homology is considerable, and both have been shown to bind a flavin mononucleotide moiety. The notable exception is Tm_FDP, whose deposited structure lacks the FMN moiety, an observation that can be accounted for by at least two nonmutually exclusive facts. On the one hand, as described by Vicente and co-workers (2007), the flavin content of heterologously overexpressed FDPs was improved by growing the expression cells and performing the purifications under specifically controlled conditions, which contrast with the experimental approach undertaken by high-throughput protocols. On the other hand, the structural motifs that bind the FMN in Dg_ROO, Mt_FDP, and Mm_FprA are conserved regions in the FDP family, but are less conserved in Tm_FDP. Therefore, the absence of FMN from the Tm_FDP structure can result from accumulation of the inherent


15

lower ability of Tm_FDP to stabilize the FMN (inferred from the sequence) and the automated high-throughput overexpression protocol. The absence of FMN in Tm_FDP is accompanied by two alternative conformations of the main chain segment composed by residues 312–320: one conformer overlaps with the protein backbone of the remaining FDPs, while the other partially occupies the region otherwise occupied by the missing FMN moiety. The crystallographic structures of FDPs provide structural clues for some of the spectroscopic and functional properties determined for studied FDPs. As mentioned earlier, the prevalence of basic over acidic residues in the flavin pocket of FDPs accounts for the higher potential of the second reduction step of FMN, with respect to canonical flavodoxins. The nature of the flavin semiquinone (one-electron reduced) that is stabilized also differs between the two protein families, which are related with the accessibility for protonation of the N5 position in the flavin isoalloxazine ring system. The FDP polypeptide establishes interactions with the isoalloxazine ring of FMN by both van der Waals contacts and H-bonds between main-chain amino groups and ring N atoms (at positions 1 and 5), or ring carbonyl O atoms (at positions 2 and 4). The H-bond with N5 precludes the protonation of this site, in contrast to what happens with canonical flavodoxins where N5 is accessible for protonation. Therefore, in the one-electron reduced state, the FMN in FDPs is deprotonated at N5 and stabilizes the red anionic semiquinone, whereas the FMN in canonical flavodoxins is protonated in the same position, therefore stabilizing the neutral blue form. The designation of ‘‘red’’ and ‘‘blue’’ for the anionic and neutral semiquinones (respectively) is associated with the visible absorption spectral features that identify either form (Ghisla and Edmondson, 2001). The structure of the FMN moiety also accounts for the ability of Mm_FprA to accept electrons directly from an organic cofactor (F420H2), contrasting with other FDPs that accept electrons from other redox proteins (typically rubredoxins). The FMN is located on the edge of the flavodoxindomain internal b sheet and, although its dimethyl benzene edge protrudes from the domain surface, it becomes occluded from solvent exposure upon dimerization. Comparing the FMN moiety among structures containing flavin, it is observed that the indole group of a tryptophan residue in Dg_ROO and Mt_FDP (Trp347 in Dg_ROO) is coplanar with the isoalloxazine plane, stacked at 3.5 A˚ distance in a ‘‘sandwich’’-like conformation, shielding the isoalloxazine Si face from the solvent. This residue is almost strictly conserved in the FDP family, being replaced by a glycine in some sequences, such as in Mm_FprA, where no homologous aromatic residue is found at this position. Moreover, the presence of this aromatic ‘‘sandwich’’ above the flavin ring may be related with the visible absorption spectral heterogeneity of the flavin moiety in FDPs (Vicente et al., 2007). The absence of this Trp residue is particularly relevant in the oxidized

16


structure of Mm_FprA, where the open conformation of the ‘‘switch loop’’ leaves enough space to accommodate parallel docking of the deazaisoalloxazine ring of F420H2 in front of the Si face of FMN (Seedorf et al., 2007), as required to closely approach the C5 of F420H2 and N5 of FMN to allow electron transfer, as observed in various other systems (Pejchal et al., 2005; Warkentin et al., 2001). The presence of the Trp residue in the majority of the FDP sequences (and possibly the same localization above the flavin ring system) precludes most FDPs from accepting electrons directly from organic cofactors (as F420H2). The absence of this Trp residue and its replacement by a Gly residue is found exclusively in methanogens (where F420H2 is an abundant cytoplasmic cofactor) and FDPs from cyanobacteria, which have an extra C-terminal flavin-binding domain, homologous to NAD(P)H: flavin oxidoreductases. It may be envisaged that the flavin from the C-terminal domain in these diflavin FDPs may have the same structural accessibility to dock in front of the flavodoxin-domain FMN prior to electron transfer. Currently, it is not known how this ‘‘sandwich’’ Trp residue affects the mechanism by which most FDPs accept electrons at the flavin moiety from the respective redox protein partners.

5. Conclusion Structural studies on flavodiiron proteins were remarkably important to deepen the knowledge of this protein family. Altogether, the solved crystallographic structures indicated a ‘‘head-to-tail’’ homodimeric structure as the minimal operative unit of this protein family. Each monomer is composed of an N-terminal metallo-b-lactamase-like domain and a C-terminal flavodoxin-like fold. Moreover, crystallographic structures provided the first and unequivocal evidence for the active site of NO/O2 reduction, a nonheme diiron center with histidine and carboxylate residues, embedded in the lactamase-like domain. The flavodoxin-like domain in turn binds a flavin mononucleotide moiety, which is located within one ˚ away from the diiron center. This observation attests the monomer at 35 A functional relevance of the ‘‘head-to-tail’’ homodimeric arrangement, as it brings the FMN cofactor of one monomer in close contact with the diiron active site from the opposing monomer. This quaternary structure thus allows FDPs to accept electrons at the flavin moiety and transfer them rapidly to the diiron center, where NO and/or O2 reduction occurs. Despite some heterogeneities found at the diiron coordination sphere among the solved structures, the accumulated body of structural data failed thus far to provide a definite answer on what determines the specificity of the various FDPs for either substrate. Therefore, more available structures are still required, ideally involving also FDPs with extra C-terminal redox-active domains.


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REFERENCES Andersson, J. O., Hirt, R. P., Foster, P. G., and Roger, A. J. (2006). Evolution of four gene families with patchy phylogenetic distributions: Influx of genes into protist genomes. BMC Evol. Biol. 6, 27. Andersson, J. O., Sjogren, A. M., Davis, L. A., Embley, T. M., and Roger, A. J. (2003). Phylogenetic analyses of diplomonad genes reveal frequent lateral gene transfers affecting eukaryotes. Curr. Biol. 13, 94–104. Brunger, A. T., Adams, P. D., Clore, G. M., DeLano, W. L., Gros, P., GrosseKunstleve, R. W., Jiang, J. S., Kuszewski, J., Nilges, M., Pannu, N. S., Read, R. J., Rice, L. M., et al. (1998). Crystallography and NMR system: A new software suite for macromolecular structure determination. Acta Crystallogr. D Biol. Crystallogr. 54, 905–921. Cameron, A. D., Ridderstrom, M., Olin, B., and Mannervik, B. (1999). Crystal structure of human glyoxalase II and its complex with a glutathione thiolester substrate analogue. Structure 7, 1067–1078. Carfi, A., Pares, S., Duee, E., Galleni, M., Duez, C., Frere, J. M., and Dideberg, O. (1995). The 3-D structure of a zinc metallo-beta-lactamase from Bacillus cereus reveals a new type of protein fold. EMBO J. 14, 4914–4921. Chen, L., Liu, M.-Y., LeGall, J., Fareleira, P., Santos, H., and Xavier, A. V. (1993). Rubredoxin oxidase, a new flavo-hemo-protein, is the site of oxygen reduction to water by the ‘‘strict anaerobe’’ Desulfovibrio gigas. Biochem. Biophys. Res. Commun. 193, 100–105. Collaborative Computational Project Number 4 (1994). The CCP4 suite: Programs for protein crystallography. Acta Cryst. D 50, 760–763. De La Fortelle, E., and Bricogne, G. (1997). Maximum-likelihood heavy-atom parameter refinement for multiple isomorphous replacement and multiwavelength anomalous diffraction methods. Methods Enzymol. 276, 472–494. DiDonato, M., Deacon, A. M., Klock, H. E., McMullan, D., and Lesley, S. A. (2004). A scaleable and integrated crystallization pipeline applied to mining the Thermotoga maritima proteome. J. Struct. Funct. Genomics 5, 133–146. Evans, P. (2006). Scaling and assessment of data quality. Acta Crystallogr. D Biol. Crystallogr. 62, 72–82. Frazaõ, C., Sieker, L., Coelho, R., Morais, J., Pacheco, I., Chen, L., LeGall, J., Dauter, Z., Wilson, K., and Carrondo, M. A. (1999). Crystallization and preliminary diffraction data analysis of both single and pseudo-merohedrally twinned crystals of rubredoxin oxygen oxidoreductase from Desulfovibrio gigas. Acta Crystallogr. D Biol. Crystallogr. 55, 1465–1467. Frazaõ, C., Silva, G., Gomes, C. M., Matias, P., Coelho, R., Sieker, L., Macedo, S., Liu, M.-Y., Oliveira, S., Teixeira, M., Xavier, A. V., Rodrigues-Pousada, C., et al. (2000). Structure of a dioxygen reduction enzyme from Desulfovibrio gigas. Nat. Struct. Biol. 7, 1041–1045. Gardner, A. M., Helmick, R. A., and Gardner, P. R. (2002). Flavorubredoxin, an inducible catalyst for nitric oxide reduction and detoxification in Escherichia coli. J. Biol. Chem. 277, 8172–8177. Ghisla, S., and Edmondson, D. A. (2001). Flavin coenzymes. In ‘‘Encyclopedia of Life Sciences.’’ Nature Publishing Group. Gomes, C. M., Frazaõ, C., Xavier, A. V., Legall, J., and Teixeira, M. (2002a). Functional control of the binuclear metal site in the metallo-beta-lactamase-like fold by subtle amino acid replacements. Protein Sci. 11, 707–712. Gomes, C. M., Giuffre, A., Forte, E., Vicente, J. B., Saraiva, L. M., Brunori, M., and Teixeira, M. (2002b). A novel type of nitric-oxide reductase: Escherichia coli flavorubredoxin. J. Biol. Chem. 277, 25273–25276.

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Gomes, C. M., Silva, G., Oliveira, R., LeGall, J., Liu, M.-Y., Xavier, A. V., RodriguesPousada, C., and Teixeira, M. (1997). Studies on the redox centers of the terminal oxidase from Desulfovobrio gigas and evidence for its interaction with rubredoxin. J. Biol. Chem. 272, 22502–22508. Hendrickson, W. A., and Ogata, C. M. (1997). Phase determination from multiwavelength anomalous diffraction measurements. Methods Enzymol. 276, 494–523. Jones, T. A., Zou, J. Y., Cowan, S. W., and Kjeldgaard, M. (1991). Improved methods for building protein models in electron density maps and the location of errors in these models. Acta Crystallogr. A47, 110–119. Justino, M. C., Vicente, J. B., Teixeira, M., and Saraiva, L. M. (2005). New genes implicated in the protection of anaerobically grown Escherichia coli against nitric oxide. J. Biol. Chem. 280, 2636–2643. Kabsch, W. (1993). Automatic processing of rotation diffraction data from crystals of initially unknown symmetry and cell constants. J. Appl. Crystallogr. 26, 795–800. Kissinger, C. R., Gehlhaar, D. K., and Fogel, D. B. (1999). Rapid automated molecular replacement by evolutionary search. Acta Crystallogr. D Biol. Crystallogr. 55, 484–491. Leslie, A. G. (2006). The integration of macromolecular diffraction data. Acta Crystallogr. D Biol. Crystallogr. 62, 48–57. Ludwig, M. L., Pattridge, K. A., Metzger, A. L., Dixon, M. M., Eren, M., Feng, Y., and Swenson, R. P. (1997). Control of oxidation-reduction potentials in flavodoxin from Clostridium beijerinckii: The role of conformation changes. Biochemistry 36, 1259–1280. McRee, D. E. (1992). A visual protein crystallographic software system for X11/XView. J. Mol. Graphics 10, 44–46. Murshudov, G. N., Vagin, A. A., and Dodson, E. J. (1997). Refinement of macromolecular structures by the maximum-likelihood method. Acta Crystallogr. D Biol. Crystallogr. 53, 240–255. Otwinowski, Z., and Minor, W. (1997). Processing of X-ray diffraction data collected in oscillation mode. Methods Enzymol. 276, 307–326. Pejchal, R., Sargeant, R., and Ludwig, M. L. (2005). Structures of NADH and CH3-H4 folate complexes of Escherichia coli methylenetetrahydrofolate reductase reveal a spartan strategy for a ping-pong reaction. Biochemistry 44, 11447–11457. Rodrigues, R., Vicente, J. B., Felix, R., Oliveira, S., Teixeira, M., and RodriguesPousada, C. (2006). Desulfovibrio gigas flavodiiron protein affords protection against nitrosative stress in vivo. J. Bacteriol. 188, 2745–2751. Sarti, P., Fiori, P. L., Forte, E., Rappelli, P., Teixeira, M., Mastronicola, D., Sanciu, G., Giuffre, A., and Brunori, M. (2004). Trichomonas vaginalis degrades nitric oxide and expresses a flavorubredoxin-type protein: A new pathogenic mechanism? Cell. Mol. Life Sci. 61, 618–623. Seedorf, H., Dreisbach, A., Hedderich, R., Shima, S., and Thauer, R. K. (2004). F420H2 oxidase (FprA) from Methanobrevibacter arboriphilus, a coenzyme F420-dependent enzyme involved in O2 detoxification. Arch. Microbiol. 182, 126–137. Seedorf, H., Hagemeier, C. H., Shima, S., Thauer, R. K., Warketin, E., and Ermler, U. (2007). Structure of coenzyme F420H2 oxidase (FprA): A di-iron flavoprotein from methanogenic Archaea catalyzing the reduction of O2 to H2O. FEBS J. 274, 1588–1599. Sheldrick, G. M. (1990). Phase annealing in SHELX-90: Direct methods for larger structures. Acta Crystallogr. A46, 467–473. Sheldrick, G. M., and Schneider, T. R. (1997). SHELXL: High-resolution refinement. Methods Enzymol. 277, 319–343. Silaghi-Dumitrescu, R., Coulter, E. D., Das, A., Ljungdahl, L. G., Jameson, G. N., Huynh, B. H., and Kurtz, D. M., Jr. (2003). A flavodiiron protein and high molecular weight rubredoxin from Moorella thermoacetica with nitric oxide reductase activity. Biochemistry 42, 2806–2815.


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Silaghi-Dumitrescu, R., Kurtz, D. M., Jr., Ljungdahl, L. G., and Lanzilotta, W. N. (2005a). X-ray crystal structures of Moorella thermoacetica FprA: Novel diiron site structure and mechanistic insights into a scavenging nitric oxide reductase. Biochemistry 44, 6492–6501. Silaghi-Dumitrescu, R., Ng, K. Y., Viswanathan, R., and Kurtz, D. M., Jr. (2005b). A flavodiiron protein from Desulfovibrio vulgaris with oxidase and nitric oxide reductase activities: Evidence for an in vivo nitric oxide scavenging function. Biochemistry 44, 3572–3579. Solomon, E. I., Brunold, T. C., Davis, M. I., Kemsley, J. N., Lee, S.-K., Lehnert, N., Neese, F., Skulan, A. J., Yang, Y.-S., and Zhou, J. (2000). Geometric and electronic structure/function correlations in non-heme iron enzymes. Chem. Rev. 100, 235–350. Ullah, J. H., Walsh, T. R., Taylor, I. A., Emery, D. C., Verma, C. S., Gamblin, S. J., and Spencer, J. (1998). The crystal structure of the L1 metallo-beta-lactamase from Stenotrophomonas maltophilia at 1.7 A resolution. J. Mol. Biol. 284, 125–136. Vicente, J. B., Justino, M. C., Gonçalves, V. L., Saraiva, L. M., and Teixeira, M. (2007). Biochemical, spectroscopic, and thermodynamic properties of flavodiiron proteins. Methods Enzymol. 437(this volume). Vicente, J. B., and Teixeira, M. (2005). Redox and spectroscopic properties of the Escherichia coli nitric oxide-detoxifying system involving flavorubredoxin and its NADH-oxidizing redox partner. J. Biol. Chem. 280, 34599–34608. Warkentin, E., Mamat, B., Sordel-Klippert, M., Wicke, M., Thauer, R. K., Iwata, M., Iwata, S., Ermler, U., and Shima, S. (2001). Structures of F420H2:NADPþ oxidoreductase with and without its substrates bound. EMBO J. 20, 6561–6569. Wasserfallen, A., Ragettli, S., Jouanneau, Y., and Leisinger, T. (1998). A family of flavoproteins in the domains Archaea and Bacteria. Eur. J. Biochem. 254, 325–332. Winn, M. D., Murshudov, G. N., and Papiz, M. Z. (2003). Macromolecular TLS refinement in REFMAC at moderate resolutions. Methods Enzymol. 374, 300–321. Zang, T. M., Hollman, D. A., Crawford, P. A., Crowder, M. W., and Makaroff, C. A. (2001). Arabidopsis glyoxalase II contains a zinc/iron binuclear metal center that is essential for substrate binding and catalysis. J. Biol. Chem. 276, 4788–4795.


C H A P T E R

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Biochemical, Spectroscopic, and Thermodynamic Properties of Flavodiiron Proteins Joaõ B. Vicente, Marta C. Justino, Vera L. Gonçalves, Lı´gia M. Saraiva, and Miguel Teixeira Contents 1. Introduction 2. Cloning of Genes Encoding Flavodiiron Proteins and Their Truncated Domains 3. Production and Purification of Recombinant Flavodiiron Proteins 4. Biochemical Characterization of Flavodiiron Proteins 5. Spectroscopic Properties 6. Redox Properties 7. Conclusions 7.1. Functional properties Acknowledgments References

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Abstract The flavodiiron proteins (FDPs), present in Archaea, Bacteria, and some protozoan pathogens (mostly anaerobes or microaerophiles), have been proposed to afford protection to microbes against nitric oxide and/or oxygen (toxic for anaerobes). The structural prototype of this protein family is a homodimer assembled in a ‘‘head-to-tail’’ configuration, with each monomer being composed of two domains: an N-terminal metallo-b-lactamase module harboring a nonheme diiron center (active site of NO/O2 reduction) and a C-terminal flavodoxin module, where a flavin mononucleotide moiety is embedded. Several FDPs bear C-terminal extra domains, which influence the composition of the respective electron transfer chains that couple NAD(P)H oxidation to NO/O2 reduction. Herein are described methodologies employed to successfully produce, isolate, and characterize fully operative recombinant flavodiiron proteins. Spectroscopic techniques, namely absorption (visible and near-ultraviolet) and Instituto de Tecnologia Quı´mica e Biolo´gica, Universidade Nova de Lisboa, Oeiras, Portugal Methods in Enzymology, Volume 437 ISSN 0076-6879, DOI: 10.1016/S0076-6879(07)37002-X

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electron paramagnetic resonance spectroscopies, allowed redox-sensitive spectral fingerprints to be obtained, used further in the functional characterization of isolated flavodiiron proteins. Altogether, these studies on pure proteins contribute to understanding the molecular determinants that govern the in vivo function of the FDPs.

1. Introduction The first report on a flavodiiron protein (FDP) focused on Desulfovibrio gigas rubredoxin:oxygen oxidoreductase (Dg_ROO), the terminal component of a soluble electron transfer chain proposed to be involved in oxygen detoxification, affording this (then considered) ‘‘strict’’ anaerobe protection from an otherwise toxic dioxygen (Chen et al., 1993b). Dg_ROO, a flavinbinding homodimer of 43-kDa monomers, was proposed to fully reduce oxygen to water, using electron equivalents from NADH, shuttled by rubredoxin and a NADH:rubredoxin oxidoreductase (Chen et al., 1993a; Gomes et al., 1997). The flurry of complete genome sequences led to the discovery of several Dg_ROO homologues widespread in Bacteria and Archaea and to establishment of the family of A-type flavoproteins (the former designation of flavodiiron proteins) (Wasserfallen et al., 1998). It was proposed that there is a common sequence core of about 400 amino acids, where a putative flavodoxin-like domain could be identified at the C terminus, and it was noted that some members of the protein family had extra C-terminal extensions. These extensions were identified as possible redox active domains, namely a rubredoxin domain in the Escherichia coli protein and a NAD(P)H:flavin oxidoreductase domain in the Synechocystis one. It was not until the crystallographic structure of Dg_ROO was solved that further insights into its functional properties were attained (Frazaõ et al., 2000). This structure elucidated that the previously proposed core of this protein family is indeed composed of an N-terminal b-lactamase-like domain fused to the flavin mononucleotide (FMN)-binding flavodoxin domain and revealed the active site of oxygen reduction: a nonheme diiron center in the lactamase fold, with carboxylate and histidine residues in its coordination sphere. It is worth noting that the structure revealed that a ‘‘head-to-tail’’ homodimeric quaternary arrangement is required to place the FMN cofactor of one monomer in close contact with the diiron center from the other monomer, allowing otherwise impaired electron transfer (the two cofactors are 25 A˚ apart in each monomer)(Vicente et al., 2007a). A survey of available FDP sequences suggested four structural classes for this protein family [adding one class to a previous classification (Saraiva et al., 2004)], accounting for the C-terminal extensions (Fig. 2.1), whose nature reflects itself in the composition of the electron transfer chains that couple

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Properties of Flavodiiron Proteins

Class A

FMN Fe-Fe

Class B

Class C

Class D

FAD

Fe-S

FMN

Flv

FMN Fe-Fe

Fe-S

FMN Fe-Fe

Fe-Fe

Figure 2.1 Modular arrangements in the flavodiiron protein family and corresponding structural classes. FMN, flavodoxin-like module, binding FMN; Fe-Fe, metallo-b-lactamase module, harboring the nonheme diiron active site; Fe-S, rubredoxin-like module, harboring a Fe-Cys4 center; Flv, NAD(P)H:flavin oxidoreductase module, binding FAD or FMN; FAD, predicted NAD(P)H:rubredoxin oxidoreductase module.

NAD(P)H or F420H2 oxidation to nitric oxide (NO) or O2 reduction. Class A FDPs are the simplest, consisting solely of the bidomain structural core (400 residues), and represent the majority of the found sequences. Class B FDPs (480 residues) bear a C-terminal rubredoxin domain and are restricted, so far, to enterobacteria. Class C FDPs (600 residues), also found so far only in cyanobacteria, have a NAD(P)H:flavin oxidoreductase C-terminal domain, and often there are multiple genes encoding these FDPs within the same organism. The newly proposed class D (900 residues) comprises FDPs where two extra C-terminal domains are fused: a rubredoxin domain and a NADH:rubredoxin oxidoreductase domain (homologous to the cognate reductase of class B FDPs). Class D FDPs were found in the genome sequences of some Clostridiales and of the protozoan pathogen Trichomonas vaginalis. Phylogenetic analyses revealed two interesting observations: (i) FDPs bearing C-terminal extensions cluster together according to their class (Saraiva et al., 2004) and (ii) genes encoding FDPs are prone to be transferred via lateral gene transfer among coexisting organisms (Andersson et al., 2003). This observation accounts for the finding of FDP-encoding genes in pathogenic protozoa, so far the only known eukaryotic FDPs. It is envisaged that the complexity of the modular arrangement of FDPs contrasts with the number of components of the corresponding electron transfer chains, i.e., class C and D FDPs should accomplish coupling of NAD(P)H oxidation to substrate reduction within the same polypeptide chain. Class B FDPs require one extra redox protein, and class A may require as many as two more redox partners to accomplish the same. This idea has been challenged only recently by a class A FDP (from a methanogenic source) that oxidizes F420H2 directly (Seedorf et al., 2004), an abundant redox cofactor in methanogenic organisms, and thus dispenses the involvement of other redox proteins.

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The idea of flavodiiron proteins as oxygen reductases came to a halt with molecular genetics studies on the E. coli flavodiiron protein (a class B FDP named flavorubredoxin because of its C-terminal rubredoxin domain) that proposed a role for this protein in NO detoxification. It was demonstrated that expression of the norV gene—encoding flavorubredoxin—was induced by NO and that an E. coli norV mutant strain was more sensitive to NO than the wild-type strain (Gardner et al., 2002; Justino et al., 2005b), with deleterious effects to NO-sensitive metabolic enzymes and affecting cell survival (Gardner et al., 2002). A role in anaerobic NO detoxification was thus proposed for flavorubredoxin, acting as an NO reductase, an activity that was confirmed further in vitro (Gomes et al., 2002). Flavodiiron proteins are presently considered a prominent family of NO-detoxifying enzymes, in the line of flavohemoglobin, although some members of the protein family retain a preference for oxygen as their substrate (Rodrigues et al., 2006; Seedorf et al., 2007). Research efforts have been employed to clarify the ambiguity of the possible roles for flavodiiron proteins. A thorough biochemical characterization of each studied member (and the corresponding electron transfer chains) is essential to understanding the molecular basis for the substrate preference (NO vs O2). In parallel, molecular genetics studies provide clues to understanding the function and relative role of each FDP in (i) NO detoxification (e.g., as a subversive mechanism of pathogens to counteract the host immune response) and/or (ii) O2 detoxification in anaerobic organisms (to allow survival of transient exposure to toxic environments).

2. Cloning of Genes Encoding Flavodiiron Proteins and Their Truncated Domains The production of recombinant flavodiiron proteins from various microbial sources has been successfully achieved by overexpression in E. coli, with T7 promoter-based expression vectors (Gomes et al., 2000, 2002; Rodrigues et al., 2006; Seedorf et al., 2004; Silaghi-Dumitrescu et al., 2003, 2005; Vicente et al., 2002; Wasserfallen et al., 1998). In our laboratory, flavodiiron proteins from classes B and C have been isolated as recombinant proteins overexpressed in E. coli: the E. coli flavorubredoxin (FlRd) and its truncated rubredoxin and flavodiiron domains, the Synechocystis sp. PCC6803 SsATF573 (the original designation for the 573 amino acid FDP from this organism, encoded by gene sll0550), and its C-terminal domain (Gomes et al., 2000, 2002; Vicente et al., 2002). The coding regions were amplified by polymerase chain reaction from genomic DNA, using primers containing restriction sites that allow cloning into the T7 expression vectors pET24aþ or pT7–7 (for the rubredoxin


25

domain, Rd domain). Cloning of DNA fragments encoding the C-terminal domains required the introduction of a NdeI site in the sense primers that changed to initiation codon (ATG) the codons of residue 422 of E. coli FlRd (Gomes et al., 2002) and residue 402 of Synechocystis SsATF573. Sequencing of the recombinant plasmids confirmed the correct nucleotide sequences (Vicente et al., 2002).

3. Production and Purification of Recombinant Flavodiiron Proteins Overexpression of E. coli flavorubredoxin (and its truncated domains) (Gomes et al., 2000) and Synechocystis SsATF573 (and the C-terminal domain) (Vicente et al., 2002) is performed in BL21-Gold(DE3) cells (Stratagene) under conditions that have been progressively optimized. Initially, Luria-Bertani (LB) broth (supplemented with 10 mM ferrous sulfate) was used to attempt the overexpression of FDPs. However, an improvement of iron and flavin incorporation was achieved by decreasing the rate of protein synthesis. This was attained by changing the growth medium to minimal medium M9 (Gomes et al., 2002; Silaghi-Dumitrescu et al., 2003), reducing the air chamber, and decreasing the growth temperatures from 37 to 28 C. Under optimized conditions, freshly transformed cells are grown in M9 minimal medium with 10 mM glucose (Ausubel et al., 1995) supplemented with 10 mM FeSO47H2O, in flasks filled to 70% of the volume, at 28 C and 130 rpm. Induction of expression is made with 100 mM isopropyl-1-thio-b-D-galactopyranoside (IPTG) when the cultures reach OD600 ¼ 0.3–0.4, and the cells are harvested after 7 h by centrifugation (11,000g, 10 min, 4 C). Cells resuspended in 10 mM Tris-HCl, pH 7.6, are disrupted in a French press cell at 130 MPa, followed by a 2-h ultracentrifugation (100,000g, 4 C) to remove cell debris. The soluble extracts are dialyzed against 10 mM Tris-HCl, pH 7.6, containing 18% (v/v) glycerol (buffer A). Complementing the buffers with glycerol increases the stability of the enzymes, preventing the loss of flavin moieties throughout the purification. In the purifications of intact flavorubredoxin, 500 mM of the protease inhibitor phenylmethanesulfonyl fluoride is added to all buffers to prevent peptidic breakage between the structural modules of FlRd (Vicente and Teixeira, 2005). All purification steps are done at 4 C. Dialyzed soluble extracts are applied onto a Q-Sepharose column (Amersham) equilibrated previously with buffer A and, by applying a gradient up to 1 M NaCl, proteins are eluted at 400–450 mM NaCl (Gomes et al., 2000, 2002; Vicente et al., 2002). After desalting, fractions are introduced into a Fractogel EMD TMAE column (Merck), eluted at 250 mM NaCl, concentrated, and further applied onto a gel filtration column (Superdex S-75 or S-200, both from Amersham)

26


equilibrated with buffer A containing 150 mM NaCl. Regarding purification of the C-terminal domain of Synechocystis SsATF573, the protein is purified from the soluble extract, in two steps: a Q-Sepharose fast flow column (Amersham) equilibrated in 20 mM KP buffer at pH 6 (buffer B) and a SP-Sepharose column (Amersham). The truncated form of SsATF573 is eluted with 200 mM KCl (Vicente et al., 2002). Protein purity is evaluated throughout the purification steps by SDS-PAGE (Garfin, 1990). The purifications of recombinant FDPs from Moorella thermoacetica (Silaghi-Dumitrescu et al., 2003), Desulfovibrio vulgaris (Silaghi-Dumitrescu et al., 2005), and D. gigas (Rodrigues et al., 2006) are conducted in similar ways, although with minor differences on the expression conditions. The production and purification of Methanobrevibacter marburgensis FprA are significantly different from other FDPs, namely the fact that the protein is only successfully isolated under anaerobic conditions. M. marburgensis FprA is overexpressed in E. coli Rosetta(DE3)pRare cells are induced at OD600 0.8 by the addition of 1 mM IPTG. Harvested cells are disrupted by ultrasonication and heated at 60 for 20 min. FprA is isolated from the soluble extract (obtained after a 150,000g ultracentrifugation) in one purification step, using a DEAE-Sepharose fast flow column equilibrated with 50 mM Tris-HCl, pH 7.6. The protein is recovered in the 400 mM NaCl fraction (Seedorf et al., 2004).

4. Biochemical Characterization of Flavodiiron Proteins Flavodiiron proteins from three (of the aforementioned four) classes have been studied, the majority of which belong to class A (properties summarized in Table 2.1). The monomeric molecular masses are determined by SDS-PAGE (Garfin, 1990), and the measured molecular masses are in accordance with the expected values inferred from the peptide sequences, namely 43–48 kDa per monomer for class A FDPs (400 residues), 54 kDa for class B FDPs (480 residues), and 70 kDa for class C FDPs (600 residues). The quaternary structure of isolated FDPs is measured by analytical gel permeation chromatography using the appropriate molecular mass standards. FDPs alternate between homodimers and homotetramers, satisfying the prerequisite of a dimer as the minimal functional unit, to allow proximity between the FMN from one monomer and the diiron center from the other monomer (Frazaõ et al., 2000; Seedorf et al., 2007; Silaghi-Dumitrescu et al., 2005). The purified proteins have been assayed for their cofactor content, namely in terms of flavin and iron incorporation. For the FDPs (and

Table 2.1

Physical-chemical properties and cofactor content of flavodiiron proteins Monomer molecular mass*

Quaternary structure

Cofactor content{ per monomer

43 kDa (44.8)

Homodimer

2 Fe (XRC) 1 FMN (XRC)

399

45 kDa (44.3)

Homodimer

410

n. d. (47.1) 45 kDa (45.1)

Homodimer

1.9 0.5 Fe (PAEA) 0.85 0.1 FMN 2 Fe (XRC)

Protein

Microorganism

a.a length

Class A Rubredoxin:oxygen oxidoreductase (ROO)

Desulfovibrio gigas

402

Flavodiiron protein

Moorella thermoacetica

Flavoprotein (Tm0755) Flavodiiron protein

Thermotoga maritima Desulfovibrio vulgaris

Flavoprotein A (FprA) Flavoprotein A (FprA) Flavoprotein A (FprA)

Methanobrevibacter arboriphilus Methanothermobacter marburgensis Rhodobacter capsulatus

402

— 404 420

45 kDa (46.1) 43 kDa (45.3) 48 kDa (46.2)

n. d.

1.8 0.1 Fe (PAEA) 0.8 0.1 FMN 2 Fe 1 FMN

Homotetramer

2 Fe 1 FMN

Homodimer

0.9 FMN (AE-HPLC)

Homodimer

Ref.

(Chen et al., 1993; Frazao et al., 2000) (SilaghiDumitrescu et al., 2003)

(SilaghiDumitrescu et al., 2005) (Seedorf et al., 2004) (Seedorf et al., 2004) (Jouanneau et al., 2000; Wasserfallen et al., 1998) (continued)

Table 2.1 (continued) Monomer molecular mass*

Quaternary structure

Cofactor content{ per monomer

1.3 FMN (AE-HPLC) 1 mol Fe/mol FMN (N/C) 0.7 FMN (AE-HPLC)

(Nolling et al., 1995)

Ref.

Protein

Microorganism

a.a length

Flavoprotein A (FprA)

Methanobacterium thermoautotrophicum strain DH

409

45 kDa (46.0)

Homodimer

Flavoprotein A (FprA)

Methanobacterium thermoautotrophicum Marburg

404

43 kDa (45.7)

Homotetramer

Escherichia coli

479

54 kDa (54.2)

Homotetramer

2.9 0.5 Fe (TPTZ) 0.8 0.2 FMN (AE-UVS)

(Frazao et al., 2000; Wasserfallen et al., 1998)

Synechocystis

573

70 kDa (63.5)

Homodimer

1.9 Fe (TPTZ) 0.8 FMN (AE-HPLC)

(Gomes et al., 2002; Wasserfallen et al., 1998)

Class B Flavorubredoxin (FlRd)

Class C SsATF573

(Wasserfallen et al., 1995)

* between brackets, molecular mass estimated from the aminoacid (a.a.) sequence. { between brackets, experimental methodology by which the cofactor was identified and quantified: XRC, X-ray crystallography; PAEA, plasma atomic emission analysis; AE-UVS, acid extraction plus visible spectroscopy; AE-HPLC, acid extraction followed by HPLC analysis; N/C – colorimetric method, using neocuproine or ferrozine. n.d. not determined.


29

truncated domains) of E. coli and Synechocystis, the protein concentrations are measured by the 2-bicinchoninic acid protein assay (Pierce) (Walker, 1994). The iron content is determined by the 2,4,6-tripyridyl-1,3,5-triazine method (Fischer and Price, 1964). Flavin quantification (adapted from Susin et al., 1993) is performed by acid extraction with TCA (10%) followed by centrifugation and supernatant neutralization with 1 M NH4CH3COO, pH 7. The nature of the flavin cofactors (FMN or FAD) is determined, when needed (Wasserfallen et al., 1998), by reversed-phase chromatography using a Nucleosil 100–5 C18 column equilibrated with 10 mM ammonium formate, pH 6.4 (containing 12% methanol), and performing a three-step gradient of increasing methanol concentration. The appropriate commercial flavin standards (FAD and FMN from Fluka) are treated and measured identically to flavins extracted from the protein samples. The extracted flavins are quantified spectrophotometrically using the following molar absorption coefficients: EFMN (l445) ¼ 12,200 M1 cm1; EFAD (l450) ¼ 11,300 M1 cm1; and EFMN þ FAD (l447) ¼ 11,750 M1 cm1 (Sober and Harte, 1968). The cofactor content of studied FDPs (Table 2.1) yields 1–2 Fe and 0.7–1 FMN per monomer. E. coli flavorubredoxin binds instead three iron ions per monomer, one in the rubredoxin domain and two in the diiron site. By replacing Fe2þ with Zn2þ in the growth medium of E. coli cells overexpressing M. thermoacetica FDP (Silaghi-Dumitrescu et al., 2003), the isolated FDP comes with a binuclear zinc site in place of the diiron site. This promiscuity is explained by equivalently high affinities of the center for Fe and Zn (Schilling et al., 2005).

5. Spectroscopic Properties To probe the functional properties of isolated flavodiiron proteins, spectroscopic methods proved to be essential, namely in characterization of the redox-active cofactors. Whereas visible spectroscopy was used mainly to characterize the flavins, electron paramagnetic resonance (EPR) spectroscopy allowed characterization of the diiron center. Visible and near-ultraviolet absorption spectra of as-isolated flavodiiron proteins are mostly dominated by the contribution of their flavin moieties. Nonheme diiron centers (Solomon et al., 2000) have much lower extinction coefficients than flavins (free or protein bound) (Ghisla and Edmonson, 2001), and therefore spectra of class A (Silaghi-Dumitrescu et al., 2003) and class C (Vicente et al., 2002) FDPs (which have only flavin and diiron cofactors) have features almost solely attributable to the flavin moieties (Fig. 2.2A and B). It is noteworthy that visible spectra of FDPs are slightly heterogeneous among different members of the protein family, in the sense that the band centered at 450 nm is broad and smooth in some cases

30


A FMN Fe-Fe

Class A

B Flv

FMN Fe-Fe

Classes A and C

C Fe-S

FMN Fe-Fe

Class B

300

400

500 600 Wavelength (nm)

700

Figure 2.2 Visible spectra of flavodiiron proteins. (A) Flavodiiron domain of Escherichia coli flavorubredoxin (i.e., with the rubredoxin domain truncated); (B) flavodiiron protein from Synechocystis sp. PCC6803, named SsATF573; and (C) E. coli flavorubredoxin. All spectra in 20 mM Tris-HCl,18% glycerol, pH 7.6, at 25.

( Jouanneau et al., 2000; Silaghi-Dumitrescu et al., 2003; Wasserfallen et al., 1998) and has two shoulders in others (Nolling et al., 1995; Vicente et al., 2002; Wasserfallen et al., 1995). This band is commonly assigned to charge transfer transitions within the isoalloxazine core, from the xylene ring to the pyrimidine ring. To understand this spectral heterogeneity, an inspection of the flavin pocket in the Dg_ROO structure was undertaken and compared with structural models generated with that structure as the template. In the structure of Dg_ROO, a tryptophan residue is coplanar with the FMN isoalloxazine ring (Trp347 in Dg_ROO). This Trp residue is conserved in FDPs where the same broad spectrum (Fig. 2.2A) is observed and appears in the same position in the modeled structures of other FDPs (Saraiva et al., 2004). However, in FDPs where this Trp is lacking, the spectral band centered at 450 nm has two shoulders (Fig. 2.2B). Therefore, it has been


31

proposed that this Trp residue may account for the spectral heterogeneity (Saraiva et al., 2004) by interacting with the FMN moiety (Vicente et al., 2008a). On top of the flavin absorption spectrum, the E. coli flavorubredoxin (class B FDP) has the contribution of the [Fe-Cys4] center from the rubredoxin domain (Fig. 2.2C) (Gomes et al., 2000; Vicente and Teixeira, 2005). Although the spectrum of FlRd overlaps in almost the entire visible region, above 560 nm, the observed broad band is almost exclusively because of the rubredoxin domain. This observation is of great value, namely in deconvoluting the functional behavior of the different cofactors. Electron paramagnetic resonance spectroscopy has proved to be a valuable tool in characterizing the cofactors of flavodiiron proteins, providing in fact the first direct spectroscopic evidence for the diiron center for a member of this protein family. Initially, EPR was used to characterize the flavin cofactor in D. gigas ROO (Gomes et al., 1997), where a signal at g 2.0 obtained under reductive conditions was attributed to the one electron-reduced semiquinone state of the flavin, proposed to correspond to the red anionic radical, based on the 1.6-mT line width (which concurred with visible spectroscopy data) (Gomes et al., 1997). Electron paramagnetic resonance spectroscopy is essential in studying the diiron site, which has very low absorptivity in the visible region. In oxidized states, only FDPs containing a rubredoxin core are EPR active, with the characteristic g 4.3 resonance typical of high-spin (S ¼ 5/2) ferric iron (Gomes et al., 2000; Vicente and Teixeira, 2005). Upon reduction, because of the spin change to S ¼ 2, the rubredoxin resonance vanishes. The diiron center is EPR silent in the oxidized state, as the two ferric ions are coupled antiferromagnetically [as confirmed by Mo¨ssbauer spectroscopy for the M. thermoacetica FDP (Silaghi-Dumitrescu et al., 2003)]. For this reason, the diiron center is only clearly detected by EPR spectroscopy in its one electron-reduced, mixed-valence (FeIII-FeII) state, displaying a rhombic signal with g values at g < 2.0 (Fig. 2.3), which has its maximal intensity at 7K (Vicente and Teixeira, 2005). An interesting observation is that obtained spectra differed in their shape and g values according to the way by which the mixed-valence state was obtained, i.e., in the presence (line 1 in Fig. 2.3) or absence (line 2 in Fig. 2.3) of redox mediators (in both cases, reduction was achieved by the addition of sodium dithionite). Nevertheless, the relaxation properties do not appear to be affected by the different shape, as their corresponding temperature dependences are practically identical (not shown). Full reduction of the diiron center to the FeII-FeII state results in the disappearance of this signal, leaving as sole EPR evidence for the diferrous state a g 11 signal in parallel-mode EPR, indicating an S ¼ 4 spin state. Hence, the achievement of a spectroscopic signature for the diiron center in E. coli FlRd allowed characterization of its thermodynamic

32


1.95

Fe-Fe

1.80

Intensity (A.U.)

1.74

1

1.93 1.88 2

1.82

300

350

400

450

Magnetic field (mT)

Figure 2.3 EPR spectra of the mixed-valence nonheme diiron center in flavorubredoxin. EPR spectra of the flavodiiron structural core (FDP-domain) of Escherichia coli flavorubredoxin, obtained in the course of a redox titration (line1) and by mild chemical reduction (line 2) with sodium dithionite. Spectra focus on the g > k2); and (iv) oxidized FlRd-reductase is rereduced by NADH (k4 ¼ 255 s1). As an oversimplification, the intramolecular eT within the complex (A* B ! A B*) was assumed to be an irreversible process, based on the information that the reduction of RdT by FlRd-reductase is largely favored thermodynamically, according to the redox potentials determined by Vicente and Teixeira (2005). Simulations of the model just described were carried out using the software FACSIMILE (AEA Technology, United Kingdom). Using this approach, experimental rates were suitably fitted when assuming the following set of rate constants: k1 ¼ 1.3 107 M1 s1, k1

13 s1, k2 ¼ 300 s1, and k3 5000 s1 (see Fig. 3.4, bottom).

3.3. Reduction of FlRd by FlRd-reductase Using stopped-flow spectroscopy, these experiments aim at measuring the kinetics of eT from FlRd-reductase to the whole, intact FlRd enzyme. These measurements are quite challenging because absorption spectra of the Fe-Cys4 and FMN centers bound to FlRd largely overlap in the UV/visible range. The optical contribution of the sole Fe-Cys4 center of FlRd can be dissected using the truncated Rd domain of the enzyme (RdT); by its subtraction from the overall spectrum of the native enzyme, the spectrum of the flavodiiron domain, largely dominated by the absorption of FMN, can be obtained (Fig. 3.5A). The validity of this experimental approach can be confirmed by expressing and purifying the isolated flavodiiron domain of FlRd, this time genetically truncated from the Rd domain, that indeed displays an absorption spectrum (not shown) very similar to that in Fig. 3.5A

56


400

k (s−1)

300

200

100

0

0

5

A* + B

10 15 20 [FIRd-reductase] (mM)

k1 k−1

A*− B

k2

25

30

A− B*

k3 k4

A + B* C

Figure 3.4 Reduction of the Rd domain of FlRd by FlRd-reductase. (Top) Rate constants observed upon mixing RdT in the oxidized state with FlRd-reductase at increasing concentrations, prereduced by NADH. Concentrations after mixing: RdT ¼ 7.7 mM; NADH ¼ 375 mM. Buffer: 50 mM Tris-HCl, 18% glycerol, pH 8.0. T ¼ 5 C. (Bottom) Model used to fit data in the top panel. A ¼ FlRd-reductase; B ¼ RdT; C ¼ NADH. The asterisk denotes the reduced state of the proteins. Oxidized RdT and reduced FlRd-reductase form a reversible complex (A*^ B; k1, k1). Afterward, intracomplex eToccurs (k2), followed by dissociation of the partner proteins (k3 k2) and rereduction of oxidized FlRd-reductase by NADH at k4 ¼ 255 s1, as determined independently. Data in the top panel are fitted by assuming k1 1.3 107 M1 s1; k1 ¼13 s1; k2 ¼ 300 s1; k3 5000 s1.

and labeled as ‘‘FDP.’’ Thus, by taking advantage of the expression of the isolated Rd and flavodiiron domains of FlRd, information is acquired on the relative optical contributions of the Fe-Cys4 and the FMN centers in FlRd, information that, as detailed later, proved to be crucial to the investigation of the eT properties of FlRd.

57

Kinetic Properties of E. coli Flavorubredoxin

A

Absorbance

0.12

FIRd

Fe-S

FMN

RdT

0.08

Fe-Fe

Fe-S

FMN

FDP

Fe-Fe

0.04

0.00

B

400

500 l (nm)

600

700

0.04

0.02 Fe-Fe

0.00 60 −0.02

k (s−1)

Δ Absorbance

FMN

−0.04

40 20 0

0

−0.06 400

500

5 10 15 20 25 [FIRd-reductase] (mM) 600

700

l (nm)

Figure 3.5 Flavorubredoxin reduction by FlRd-reductase. (A) Absorption spectrum of 10 mM oxidized flavorubredoxin (FlRd) and its individual domains (RdT and FDP). The spectrum of the flavodiiron (FDP) domain was obtained by subtracting the spectrum of RdT from the spectrum of the whole FlRd protein. (B) Absorption changes assigned to the FMN moiety of FlRd, after mixing the protein with FlRd-reductase prereduced by excess NADH. Concentrations after mixing: [FlRd] ¼ 10 mM; [FlRdreductase] ¼ 2.25 mM; [NADH] ¼ 375 mM. Buffer: 50 mM Tris-HCl,18% glycerol, pH 8. T ¼ 5 C. The optical contribution of the FMN center was obtained as detailed in the text. (Inset) Rate constant at which Fe-Cys4 (circles) and FMN (squares) centers of FlRd are reduced at increasing concentrations of FlRd-reductase.

The kinetics of eT from FlRd-reductase to oxidized FlRd was probed under pseudo-first-order conditions by mixing oxidized FlRd (20 mM) with FlRd-reductase (4.5 mM) prereduced by 750 mM NADH in the

58


stopped-flow instrument. Under these conditions, both the Fe-Cys4 and the FMN center in FlRd are reduced quickly by FlRd-reductase (1.15 are pooled and concentrated.

3. Purification of Recombinant NorBC Recombinant P. denitrificans NorBC is purified from cultures of E. coli JM109 that have been cotransformed with the pEC86 plasmid, which allows constitutive expression of the cytochrome c maturation genes required for NorC assembly (Arslan et al., 1998; Tho¨ny-Meyer et al., 1996) and the pNOREX plasmid (Butland et al., 2001). Methods of cell culture and purification are essentially as described by Butland et al. (2001) with some minor modifications. For large-scale preparations of recombinant NorBC, up to 20 cultures (800 ml of Terrific Broth in a 2-liter unbaffled conical flask) are used. Each culture is supplemented with the required antibiotics (100 mg ml1 ampicillin and 36 mg ml1 chloramphenicol) and inoculated with 15 ml of a 50-ml starter grown overnight in LB. After inoculation, the 800-ml cultures are grown at 37 on a rotary shaker (200 rpm) until A600 ¼ 0.4, at which point isopropyl-b-D-thiogalactoside is added to a final concentration of 1 mM in order to induce expression of the nor genes. After induction the cultures are grown according to the schedule outlined in Table 5.1. Cells are subsequently harvested (6900 g for 15 min at 4 ), frozen in liquid nitrogen, and stored at –80 until needed. Cells are thawed and resuspended in 100 mM Tris-HCl, 50 mM NaCl, and 1 mM EDTA, pH 7.6, and broken by two passages through a French pressure cell at 11,000 psi. Protease inhibitors (5 mM EDTA, 1 mM AEBSF 1 mM pepstatin, and 1 mM leupeptin) are added and membranes

Table 5.1 Batch culture growth conditions for E. coli expressing recombinant NorBC Time after induction (hours)

Shaking at 180 rpm

Temperature ( )

0–5 5–10 10–18

Yes No Yes

30 18 30

86

Sarah J. Field et al.

collected by ultracentrifugation (180,000 g, 1 h, 4 ). Membranes are resuspended in the same resuspension buffer and reharvested by ultracentrifugation (180,000 g, for 60 min at 4 ). The washed membranes are again resuspended in the same buffer, frozen rapidly in liquid nitrogen, and stored at –80 until needed. Membranes are thawed and solubilized using the same protocol described earlier for the solubilization of P. denitrificans membranes. Unsolubilized material is removed by ultracentrifugation (180,000 g, for 60 min at 4 ). The supernatant is diluted threefold in 50 mM Tris-HCl and 0.05% DDM, pH 7.6 (buffer A), and loaded onto a 100-ml Q-Sepharose column preequilibrated in buffer A. The column is then washed with buffer A until all unbound protein has washed through the column as judged by A280 of the eluent returning to zero. Other unwanted proteins are removed by exposing the column to the following gradient: 0–10% 50 mM Tris-HCl, 1 M NaCl, and 0.05% DDM (buffer B) over 200 ml. The column is then washed with 10% buffer B until the A280 of the eluent returns to zero. Finally, NorBC is eluted with a gradient of 10–55% buffer B over 1000 ml. Recombinant NorBC elutes at approximately 450 mM NaCl. A flow rate of 5 ml min1 is used throughout, and 10-ml fractions are collected routinely. This method also provides the basis for the study of wild-type and mutant forms of NorBC that have been engineered in a proton-conducting E-channel that leads from the periplasm to the active site of the enzyme (Butland et al., 2001; Flock et al., 2005; Thorndycroft et al., 2007).

4. Amperometric Assays of NO Consumption The consumption of NO by both membrane vesicles and purified NorBC can be measured under anaerobic conditions using a Clark-type oxygen electrode (Oxytherm, Hansatech Instruments, Kings Lynn, UK) that has been modified to allow the platinum cathode to be polarized at –0.8 V with respect to the Ag:AgCl reference electrode, which increases its sensitivity toward NO. The electrolyte is 1 M KCl, and a polytetrafluoroethylene membrane (12.5 mM thickness) is used as a gas-permeable barrier to protect electrodes from the reactants in the chamber. Stock solutions of NO are prepared by the addition of 8 ml NO gas to 3 ml, nitrogen-sparged, water, pH 3.0. The concentration of NO in the stock solution can be checked readily by titrating it into a solution of reduced myoglobin of known concentration and monitoring the absorption changes in the Soret region (e421 ¼ 114 mM1 cm1). The reaction chamber of the electrode is filled with 2 ml of 20 mM phosphate buffer, pH 7.6, warmed to 30 while stirring, and allowed to equilibrate for 5 min. Oxygen is removed from the chamber by the glucose

87

Bacterial NorBC

(16 mM )/glucose oxidase (4 U/ml)/catalase (20 U/ml) system (Fig. 5.2). Once anaerobic, 25 ml of a saturated NO solution (45 nmol) is added to the chamber. The electrode is allowed to equilibrate and three further aliquots are added. The response of the electrode should be directly related to the amount of NO, and this calibration should be carried out each time the electrode is used. If the response is too small, then the NO solution should be discarded and a fresh solution made. This calibration routine is especially important with the Oxytherm electrode, which is interfaced to a software program that converts the electrode current into the amount of oxygen present in solution. The present version of the software does not allow any other output. Consequently, when the electrode responds to the addition of NO under anaerobic conditions, the output given is in ‘‘nmol O2.’’ The calibration routine allows derivation of a conversion factor so that the amount of NO in solution can be calculated readily from the apparent amount of oxygen reported by the software.

75 65

DM Glucose Glucose oxidase

Paz

Electrode response (arbitary units)

55

25ul saturated NO

NOR

45 Background rate

35

Total rate

25 15 5 −5

Catalase Ascorbic Acid

−15 −25 0.0

6

12

Time (min)

18

21

Min 30.

Figure 5.2 NO reduction by purified NorBC. The assay was carried out in 20 mM phosphate buffer, 50 mM NaCl, and 0.02% DDM, pH 7.0, in a stirred reaction chamber thermostated at 30.The vessel was made anaerobic by the addition of glucose (16 mM ), glucose oxidase (4 U ml1), and catalase (20 U ml1). Subsequent additions to the reaction mixture were as follow: ascorbic acid (5 mM), saturated NO solution (4 45 nmol), and pseudoazurin (20 mM). The reaction was initiated by the addition of purified NorBC (10 nM ). Rates of NO consumption were determined by the time taken for the NO signal at the start of the enzyme-catalyzed reaction to reduce by 20%.

88


5. Pseudoazurin as an Electron Donor in Assays of NorBC One problem in assaying NOR is that although NO reductase activity can be measured easily using an amperometric steady-state assay, conditions reported in the literature have used different mediators either singly or in combination with the chemical electron donor ascorbate. For example, phenazine methosulfate (PMS) (Dermastia et al., 1991), N,N,N’,N’-tetramethyl-p-phenylenediamine (TMPD) (Hendriks et al., 1998b), and a combination of horse heart cytochrome c plus TMPD (Matsuda et al., 2002) or 2,3,5,6-tetramethylphenylenediamine (Carr and Ferguson, 1990) have all been used to mediate electron transfer to NOR. No single definitive method for assaying NOR using a physiologically relevant electron donor has been reported, which led us to devise a reliable steady-state assay of NOR activity using a physiologically relevant species, pseudoazurin, as the immediate electron donor. Figure 5.3 shows a representative trace of the consumption of NO by resuspended P. denitrificans membrane vesicles. The vesicles (100 ml containing

30 20

NO

40

NO

50

Succinate

Arbitary units

60

NO

70

Pseudoazurin

P.denitrificans membranes

80

10 0 0

2

4

6 8 Time (min)

10

12

Figure 5.3 Succinate-dependent NO consumption by P. denitrificans membrane vesicles. The reaction was carried out using 100 ml of membrane vesicles in nitrate-free succinate medium, succinate (30 mM ), Na2HPO4 (55 mM ), KH2PO4 (11 mM ), NH4Cl (6 mM ), MgSO4 (0.4 mM ), and Vishniac and Sante’s trace element solution (2 ml l1), pH 7.6, in a stirred reaction chamber thermostated to 30.The vessel was made anaerobic by the addition of glucose (16 mM ), glucose oxidase (4 U ml1), and catalase (20 U ml1). The reaction was initiated by the addition of saturated NO solution (90 mM) in the absence or presence of additional pseudoazurin (60 mM ).

Bacterial NorBC

89

approximately 0.2 mg total protein) are added to 2 ml of nitrate-free succinate medium, succinate (30 mM ), Na2HPO4 (55 mM ), KH2PO4 (11 mM ), NH4Cl (6 mM ), MgSO4 (0.4 mM ), and Vishniac and Sante’s trace element solution (2 ml liter1), pH 7.6, in a stirred reaction chamber thermostated at 30 . The lid is secured, and oxygen is removed from the chamber by the glucose (16 mM)/glucose oxidase (4 U ml1)/catalase (20 U ml1) system. Once the chamber becomes anaerobic, 90 mM NO is added. There is rapid consumption of NO because residual pseudoazurin and cytochrome c550 remain associated with the membrane after cell breakage and are able to shuttle electrons, initially donated to the electron transfer chain by the oxidation of succinate, from the cytochrome bc1 complex to NorC. The addition of nitrogen-sparged pseudoazurin (60 mM) increases the rate of NO reduction significantly, implying both that the amount of electron carriers remaining after membrane preparation are not sufficient to maintain optimal rates for NO-dependent respiration and that pseudoazurin has a physiological role in donating electrons to NOR. These data are consistent with other experiments showing that the ability of NOR to reduce nitric oxide is attenuated in strains unable to express either pseudoazurin of cytochrome c550 (Pearson et al., 2003; Thorndycroft et al., 2007). Recombinant pseudoazurin is used routinely in our laboratory as the immediate electron donor in assays of both native and recombinant NorBC in membrane fractions and after purification. To obtain recombinant pseudoazurin for these experiments, the gene encoding pseudoazurin in Paracoccus pantotrophus, pazS, was amplified and cloned into the pET-24d expression vector exactly as described by Pauleta and colleagues (2004). The resulting vector (pET-psaz) is used to transform E. coli BL21(DE3), which is grown aerobically at 37 in LB medium supplemented with kanamycin (30 mg ml1) and CuSO4 (0.5 mM ), for 24 h, without induction. Cells are harvested by sedimentation (5500 g for 20 minutes) and used immediately for the purification of pseudoazurin using a previously published method (Leung et al., 1997). Cells from 12 1-liter cultures cells are resuspended in a volume of 100 mM Tris-HCl, pH 7.3, equivalent to eight times their wet weight before being broken by five cycles of freezing, in a dry-ice ethanol bath, and thawing. Broken cells are sedimented by centrifugation (10,000 g, 30 min, at 4 ) and the supernatant, which contains the periplasmic proteins, including pseudoazurin, is recovered. A few crystals of potassium ferricyanide are added to the periplasmic fraction to oxidize the sample before it is applied directly to a 750-ml DEAE-Sepharose fast-flow column equilibrated with 50 mM Tris-HCl, pH 7.3. The column is developed with a gradient of 0–400 mM NaCl over 5 column volumes. The blue fractions that contain pseudoazurin are pooled, and ammonium sulfate is added to 80% saturation (561 g liter1). Precipitated proteins are removed by centrifugation, and the blue supernatant is applied to a 25-ml 15PHE-Sepharose

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column equilibrated with 80% NH4SO4 in 50 mM Tris-HCl, pH 8.0. The column is first washed with 3 column volumes of the same buffer before the column is developed using a gradient of 80–0% NH4SO4 over 10 column volumes at a flow rate of 2.0 ml min1. Fractions containing pseudoazurin (which elutes at about 30% NH4SO4) are pooled and exchanged to 20 mM potassium phosphate, pH 8.0, and the protein is concentrated. The concentration of the pseudoazurin solution can be determined using an E ¼ 1300 M1 cm1 at 590 nm. Pseudoazurin is also used as an electron mediator in the amperometric assay of purified NorBC, which is carried out in anaerobic potassium phosphate buffer supplemented with 0.02% DDM in a stirred reaction chamber thermostated at 30 . After the chamber has been made anaerobic with glucose/glucose oxidase/catalase as described earlier, ascorbic acid is added to a final concentration of 5 mM (see Fig. 5.2). Ascorbic acid is able to reduce pseudoazurin rapidly and hence acts as a source of electrons for the reaction. The trace is allowed to stabilize before the NO is added to a final concentration of 90 mM. Once the trace becomes stable, pseudoazurin is added and a small reduction in the NO signal is seen due to minute quantities of oxygen present in the pseudoazurin solution reacting with the NO (see Fig. 5.2). Once a background rate of NO consumption is established, the reaction is initiated by the addition of purified NorBC. The representative progress curve describing the NorBC catalyzed consumption of NO shown in Fig. 5.2 shows how the rate of NO consumption increases toward the end of the curve as a consequence of the relief of substrate inhibition (Girsch and de Vries, 1997). To compare the rates of reaction between samples of NOR at a constant concentration of NO, the rate of reaction is calculated by the time taken for 20% of the initial NO present in the system to be consumed. Using this approach, we have determined that for native NorBC the optimum pH for the reaction is pH 6 with an apparent Vmax of 904 min1 and an apparent Km for pseudoazurin of 20 mM (Thorndycroft et al., 2007). This assay also allows us to measure the activity of recombinant NorBC expressed in membranes of E. coli, which allows site-directed mutants to be screened rapidly and reliably for changes in catalytic activity (Thorndycroft et al., 2007). This is possible because E. coli lacks a cytochrome bc1 complex so that there is no input of electrons to NorBC from the E. coli respiratory chain and the rate of NO consumption measured depends solely on the exogenous electron donors. This system has been used to show that mutations in two conserved glutamate residues, E122 and E125, which lie in a periplasmic loop between helix III and helix IV of NorB, lead to changes in steady-state activity that are consistent with them forming the entrance to a channel that might couple the movement of protons from the periplasm to the active site with internal electron transfer during turnover (Thorndycroft et al., 2007).

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6. Preparation of NOR for Spectroscopic Investigation NorBC has four distinct redox centers and, in principle, can be studied at five different levels of reduction. In the fully oxidized state, the UVvisible spectrum of NorBC, as isolated, is characterized by a ligand-to-metal charge transfer (LMCT) absorption band at 595 nm. This distinct feature is thought to arise from the active site ferric heme b3, which is linked to FeB by a m-oxo bridge. At present, it is not known to which redox state of the enzyme nitric oxide binds during the catalytic cycle. In order to address this problem, we have developed methods for preparing NorBC in different redox states, which are described next.

6.1. Fully oxidized NorBC NorBC has been studied in three different redox states by both UV/visible and electron paramagnetic resonance (EPR) spectroscopies. The spectrum of the fully oxidized, as prepared form of the native enzyme from P. denitrificans, is shown in Fig. 5.1. The spectrum is typical of a hemoprotein and is characterized by expected bands in the a,b (550 and 523 nm) and Soret (410 nm) regions of the spectrum. In addition, there is a weak LMCT band at 595 nm, which is diagnostic of the m-oxo bridge that forms between heme b3 and FeB in the active site (Field et al., 2002; Gro¨nberg et al., 1999). The form of the UV/visible spectrum of fully oxidized NorBC as isolated is independent of pH, and the presence of the m-oxo bridge prevents the binding of small anionic ligands such as cyanide and fluoride ions to the ferric heme b3 (Gro¨nberg et al., 2004).

6.2. Fully reduced NorBC Figures 5.4A and 5.4B show the visible spectrum of samples of fully reduced NorBC prepared for EPR spectroscopy. This technique requires samples in the concentration range of 80–150 mM and consequently only the a,b region of the spectrum lies within the dynamic range of a typical UV/visible spectrophotometer. Because of the inherent oxidase activity of NorBC, fully and partially reduced samples must be prepared under highly anaerobic conditions, which are achieved by placing the enzyme solution, cuvettes, EPR tubes, and syringes in an anaerobic glove box ( Belle Technologies) for at least 30 min before beginning an experiment. Reduced and partially reduced samples are made routinely by the titration of NOR with europium dichloride (EuCl2). EuCl2 has a midpoint potential of approximately –400 mV depending on conditions; when complexed 1:1 with EGTA,

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A

B 65

65

60

60 OH2

55

Fe

e mM−1 cm−1

50

FeB

Fe

FeB

50

45

45

40

40

35

35

30

30

25

25

20

20

15

15

10

10

5

5

0 500

OH

55

0 550

600

650

700

500

550

600

650

700

Wavelength (nm)

Figure 5.4 Visible region electronic absorption spectra of fully and partially reduced forms of native NorBC from P. denitrificans. Electronic absorption spectra in the 500- to 700-nm region were recorded for NorBC in fully oxidized (solid line),‘‘three electronreduced’’ (dashed line), and fully reduced (dotted line) states. Samples were prepared in 20 mM Bis-Tris propane, 50 mM NaCl, and 0.02% DDM, pH 6 (A), and pH 8.5 (B). Levels of reduction indicated were achieved by slow titration of EuCl2:EGTA. (Inset) The proximal ligand to the active site heme b3 at each pH.

this drops to approx –1 V, allowing stochiometric reduction of proteins ( Vincent et al., 2003). EuCl2 is extremely oxygen sensitive and stock 100 mM solutions are prepared in anaerobic water inside an anaerobic glove box. Solutions (1 ml) of EuCl2 are prepared at the desired concentration by mixing with anaerobic EGTA (450 mM stock in 1 M NaOH) and dilution. Fresh EuCl2/EGTA solutions must be made every few hours. Protein samples for EPR are prepared in 1-mm path length cuvettes at a concentration of 80–150 mM. The EuCl2/EGTA is added to the enzyme solution in sealed cuvettes inside the glove box, and the extent of reduction of NorBC is monitored by UV/visible spectroscopy until the required partially or fully reduced state is reached. The UV/visible spectrum of fully reduced NorBC is essentially the same irrespective of pH and exhibits intense features at 525 and 550 nm that are characteristic of the a,b bands of reduced heme c in NorC with a shoulder at 560 nm arising from the reduced low-spin heme b in NorB.

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Carbon monoxide (CO) has been used extensively in the study of HCuOs because of the photolability of the bond between ferrous active site heme and CO. The recombination of photolyzed CO to the heme iron is influenced by a number of factors; therefore, this technique can give information about the effective pKa of the proximal ligand to the heme, the polarity of the heme distal pocket, and steric factors influencing binding of the substrate. Fully reduced NorBC is prepared by the addition of sodium ascorbate (5 mM ) and phenazine methosulfate (1 mM) (ascorbate/PMS) to yield a spectrum similar to that seen in Fig. 5.4. The effect of adding 1 mM CO to NorBC prepared in this way is shown in Fig. 5.5. While there is no change in wavelength maximum of the Soret band (420 nm) in response to the addition of CO, it does intensify and become narrower: e420 ffi 480 mM1 cm1 for the CO-bound form compared to e420 ffi 370 mM1 cm1 for the ascorbate/PMS-reduced form. This form of the enzyme has been used successfully to investigate the dynamics of CO recombination to the active site (Hendriks et al., 2001) and also as the starting point for ‘‘flow-flash’’ experiments to examine proton-coupled electron transfer monitored either electrochemically (Hendriks et al., 2002) or optically (Flock et al., 2005).

500 450 400 e mM−1 cm−1

350 300 250 200 150 100 50 0 400


700

Figure 5.5 CO binding to fully reduced native P. denitrificans NorBC. The electronic absorption spectrum (400^700 nm) was recorded of the fully oxidized form NorBC as prepared (solid line). Reduction of NorBC was achieved by the addition of sodium ascorbate (5 mM ) and phenazine methosulfate (1 mM) (dashed line). Sufficient saturated CO solution (1 mM ) was added anaerobically to the sample of reduced NorBC to give a final concentration of 0.1 mM CO in the cuvette (dotted line). All samples were prepared anaerobically in 20 mM BTP, 50 mM NaCl, and 0.02% DDM, pH 7.6.

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6.3. Partially reduced forms of NorBC Visible region electronic absorption spectra of the partially reduced form of NorBC referred to previously as ‘‘three electron-reduced’’ species are shown in Fig. 5.4. It can be seen that the low-spin hemes b and c are reduced substantially, as indicated by the increased intensity of the bands at 550 and 560 nm compared to the fully oxidized enzyme. In addition, the LMCT band seen at 595 nm in the fully oxidized enzyme shifts, although the position and the intensity of the shifted band are critically dependent on the pH of the buffer (Field et al., 2002). At pH 8.5, the distal ligand to heme b3 is a hydroxide, which gives rise to a LMCT band at 605 nm (see Fig. 5.4B). Decreasing the pH leads to protonation of the hydroxide and a shift in the LMCT band to 630 nm, as shown in Fig. 5.4A. The initial shift in the LMCT band from 595 nm associated with partial reduction is because of the breaking of the m-oxo bridge at the active site, which opens it to the binding of exogenous ligands to heme b3 (Gro¨nberg et al., 2004). The trigger for this breaking of the m-oxo bridge was thought to be because of the reduction of the nonheme iron FeB (Gro¨nberg et al., 1999). However, more recent work in our laboratory suggests that the trigger for breaking the m-oxo bridge may, in fact, be reduction of the low-spin heme b (S. J. Field, unpublished data). Visible region electronic absorption spectra also suggest that there is an increase in the amount of reduced heme in the fully reduced form of the enzyme compared to the ‘‘three electron-reduced’’ samples (see Fig. 5.4). A trivial explanation might be that when NorBC is poised in a partially reduced form, some of the low-spin heme b remains oxidized. However, this would also have to be true for the low-spin heme c, as the intensity of the 550-nm band also increases as the protein becomes fully reduced (see Fig. 5.4). Because the increase in absorbance associated with complete reduction affects both 550- and 560-nm bands, it is most likely that it is due to an underlying and poorly resolved feature, which is likely to be the broad transition associated with reduction of the low-potential heme b3 (see Figs. 5.4A and 5.4B). These characteristic spectroscopic changes have been exploited to determine the redox potential of the metal centers in NorBC in mediated potentiometic titration using sodium dithionite as a reductant (Gro¨nberg et al., 1999). As reductant is introduced into the system, the low-spin b- and c-type hemes start to reduce, as indicated by the increase in absorbance at 550 and 560 nm. There is also a shift in the position of the CT band from 595 nm described earlier (Fig. 5.6). Reduction potentials of low-spin heme c and low-spin heme b have been estimated to be 310 and 345 mV, respectively (Gro¨nberg et al., 1999). A plot of change of absorbance at 595 nm against potential gives rise to a curve that can fitted to two independent 0 0 n ¼ 1 oxidation/reduction events (Em ¼ þ320 mV and Em ¼ þ60 mV).

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0.55 0.50 0.45

Absorbance

0.40 0.35 0.30 0.25 0.20 0.15 0.10 0.05 0.00 500

600 650 550 Wavelength (nm)

700

Figure 5.6 Representative spectra recorded during a mediated potentiometric titration of native P. denitrificans NorBC. NOR (10 mM) in 20 mM Tris-HCl, 0.02% DDM, 340 mM NaCl, and 0.5 mM EDTA, pH 7.6, was reduced by additions of dithionite in the presence of 20 mM of each of the following mediators: phenazine methosulfate (þ80 mV), phenazine ethosulfate (þ55 mV), diaminodurene (þ250 mV), juglone (þ30 mV), 5-anthroquinone-2-sulfonate (^225 mV), 6-anthroquinone-2,6-disulfonate (^185 mV), and benzyl viologen (^311 mV). Spectra were recorded at the following potentials: E0m¼ þ400 mV (solid trace), þ360 mV (dotted trace), þ300 mV (dashed trace), and þ140 mV (dot^dash trace). 0

The second phase of reduction (Em ¼ þ60 mV) is associated with disappearance of the red-shifted CT band and can confidently be assigned to the reduction of heme b3 (Gro¨nberg et al., 1999). The relatively low potential of the catalytic heme in NorBC is somewhat surprising and has led to the proposal that the fully reduced form of the enzyme is not the catalytically relevant species. The apparent separation of the redox potential of heme b3 and the redox potentials of heme c and heme b can be exploited to poise NorBC, either chemically (as described earlier) or electrochemically (Field et al., 2002; Gro¨nberg et al., 1999). These partially reduced forms of the enzyme are excellent subjects for further interrogation of the environments of the heme centers by advanced spectroscopic methods. This approach to the study of NOR is illustrated by the EPR experiments described in the next section.

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7. Electron Paramagnetic Resonance Spectroscopy Originally the distribution of electrons in the ‘‘three electron-reduced’’ from of NorBC seemed clear; both low-spin heme c and low heme b are fully reduced and the third electron resides on the active site FeB (Field et al., 2002; Gro¨nberg et al., 1999). However, the electronic absorption spectra shown in Fig. 5.4 suggest that the situation is rather more complicated, as the molar extinction coefficient of the ‘‘three electron-reduced’’ form appears to be dependent on the pH of the sample. The extent to which both low-spin hemes are reduced is less at pH 8.5 (see Fig. 5.4B) than at pH 6.0 (see Fig. 5.4A). This may be accounted for by a different distribution of the added electrons among the four redox centers, which is also indicated by low-temperature EPR spectra of the same samples (Fig. 5.7). Figure 5.7A shows X-band EPR spectra of NorBC in three different oxidation states recorded at pH 6.0. The spectrum of the oxidized form of the enzyme is dominated by features originating from the low-spin hemes. A rhombic trio with g values of 3, 2.25, and 1.45 has been assigned to the B

A

Fully oxidised

Fully oxidised

3 e− reduced 3 e− reduced

Fully reduced

1000 2000 3000 4000 5000 6000 Field (gauss)

Fully reduced

1000 2000 3000 4000 5000 6000 Field (gauss)

Figure 5.7 X-band EPR spectra of NorBC. The fully oxidized, ‘‘three electronreduced’’and fully reduced states buffered in 20 mM BTP, 50 mM NaCl, and 0.02% DM, pH 6.0 (A), or 20 mM BTP, 50 mM NaCl, and 0.02% DM, pH 8.5 (B). All spectra were recorded at 10 K, with a modulation amplitude of 1 mT and a microwave power of 2 mW.

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ferric low-spin heme b in NorB (Cheesman et al., 1998; Gro¨nberg et al., 1999). The ramp-shaped signal at g ¼ 3.5 is a ‘‘high gmax signal’’ and represents the gz component of a rhombic trio in which values of the gy and gx components are less than 2. This signal is assigned to the ferric lowspin heme c in NorC (Cheesman et al., 1998; Gro¨nberg et al., 1999). Although the ferric high-spin heme b3 and ferric FeB both contain unpaired electrons and so in principle should be visible by EPR spectroscopy, no signals arising from the active site are present in the spectrum of the oxidized enzyme. This is because the high-spin heme b3 is coupled magnetically to the FeB to form an integer spin and hence EPR silent species (Cheesman et al., 1998). The signals seen at g ¼ 4.3 and g ¼ 6 may be because of small amounts of uncoupled active site components that represent less than 2% of the sample. The spectrum of oxidized NorBC is essentially the same at pH 8.5 (see Fig. 5.7B). Electron paramagnetic resonance spectra of the ‘‘three electronreduced’’ form of NorBC recorded at pH 6.0 (see Fig. 5.7A) and pH 8.5 (see Fig. 5.7B) are somewhat different. In both spectra the high gmax signal is no longer present, indicating that the low-spin heme c is fully reduced. Both spectra apparently show a small amount of the gz component (g ¼ 3.01) of the rhombic trio seen in oxidized spectra. The most obvious explanation is that this is a consequence of some residual oxidized low-spin heme b, although this is not consistent with the reported reduction potentials of heme b and heme c (Gro¨nberg et al., 1999). Careful inspection of the EPR signal at g ¼ 3.01 in the ‘‘three electron-reduced’’ EPR spectrum reveals it to have a slightly different line shape to the feature seen in the oxidized spectrum. This leads us to believe that it could be because of a small amount of uncoupled low-spin heme b3 at the active site, which becomes visible in the EPR experiment as the nonheme FeB becomes reduced. At pH 8.5, a signal at g ¼ 4.3 appears in the EPR spectrum of the ‘‘three electron-reduced’’ sample (see Fig. 5.7B) that is attributed to ferric nonheme FeB. Electrochemical poising experiments monitored by EPR spectroscopy have suggested that at this pH the reduction potentials of the centers at the active site are much closer than those determined from spectropotentiometric titrations recorded at pH 7.6 (S. J. Field, unpublished data). Earlier analyses of spectropotentiometric titrations placed the reduction potential of the nonheme FeB at around 320 mV (Gro¨nberg et al., 1999). If this is correct, then it would be expected that, in the ‘‘three electron-reduced’’ form, the nonheme FeB, along with both low-spin hemes, would be reduced and that the EPR spectrum would show a significant signal at g ¼ 6 arising from uncoupled ferric high-spin heme b3 (E0m ¼ þ60 mV). Figure 5.7 shows this not to be the case; instead, only small increases in both the g ¼ 6 (ferric heme b3) and the g ¼ 4.3 (ferric nonheme FeB) signals are seen together with the g ¼ 3.01 feature mentioned previously.

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This observation, together with the pH dependence of the electronic absorption (see Fig. 5.4) and EPR spectra (see Fig. 5.7) of the ‘‘three electron-reduced’’ form of NorBC, leads us to believe that not only is the reduction potential of the nonheme FeB lower than we originally suggested, but also that it is pH and possibly temperature dependent. If the reduction potentials of heme b3 and nonheme FeB are sufficiently close when the protein is poised in the ‘‘three electron-reduced’’ state, the sample will in reality be a mixture of two, three, and four electron-reduced forms of NorBC. In the two electron-reduced form, the dinuclear center will be fully oxidized and hence EPR silent. The four electron-reduced (fully reduced) enzyme will also be EPR silent as both active site metal centers are reduced. In the remaining fraction of the enzyme that genuinely contains three electrons, the single electron in the active site can reside on either heme b3 or the nonheme FeB, giving a mixed population of EPR active species. In one of these subpopulations, heme b3 is oxidized and the nonheme FeB is reduced. This accounts for the small g ¼ 6 EPR signal from ferric highspin heme b3 and possibly the new rhombic signal from ferric low-spin b3 described earlier. The second subpopulation of the enzyme containing three electrons has a dinuclear center in which the ferrous heme b3 is EPR silent and ferric nonheme FeB gives rise to the g ¼ 4.3 feature, which is more noticeable at pH 8.5 (see Fig. 5.7B). Further redox titrations in this pH range, which are monitored by both optical and EPR spectroscopies, will be required to resolve this issue completely.

8. Concluding Remarks NorBC is a complex integral membrane protein containing four distinct redox centers, two of which form a dinuclear active site, which can be poised in a number of different redox states. A robust spectropotentiometric and kinetic description of the native enzyme has depended on development of the purification and assay procedures described in this chapter. These methods have in turn enabled the regular production of a highly purified enzyme that has allowed us to record a series of benchmark spectra that report on its structural integrity. Engineered forms of NorBC will be important for future structure–function studies on the enzyme, but it is important to remember that E. coli does not have its own norCB genes nor does not have any enzyme that has a nonheme iron center such as that present in the active site of NOR. Consequently, the successful synthesis of recombinant holo P. denitrificans NorCB in E. coli was a major challenge, and our success has opened the way for the production of characterization of a range of mutant forms of the enzyme deficient in key residues important in

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proton transfer and metal ion binding. This will underpin future structural biology programs that will provide molecular structures to interface with currently available spectropotentiometric and kinetic data.

ACKNOWLEDGMENTS We acknowledge the contributions to the NOR work presented in this chapter made by a number of colleagues, including Dr. Gareth Butland, Dr. Karin Gro¨nberg, Dr. Louise Prior, and Dr. Lola Roldan. Dr. Myles Cheesman and Dr. Julea Butt provided much helpful advice on the application of EPR spectroscopy and electrochemical methods to the study of NOR. FHT was funded by a U.K. Medical Research Council Priority Area Studentship. Recent work in our laboratory was supported by grants from the John and Pamela Salter Trust to NJW and the U.K. Biotechnology and Biological Sciences Research Council to DJR (B19851) and NJW/DJR (BBC0077191).

REFERENCES Arslan, E., Schulz, H., Zufferey, R., Kunzler, P., and Tho¨ny-Meyer, L. (1998). Overproduction of the Bradyrhizobium japonicum c-type cytochrome subunits of the cbb3 oxidase in E. coli. Biochem. Biophys. Res. Comm. 251, 744–747. Bell, L. C., Richardson, D. J., and Ferguson, S. J. (1992). Identification of nitric oxide reductase activity in Rhodobacter capsulatus: The electron transport pathway can either use or bypass both cytochrome c2 and the cytochrome bc1 complex. J. Gen. Microbiol. 138, 437–443. Busch, A., Friedrich, B., and Cramm, R. (2002). Characterization of the norB gene, encoding nitric oxide reductase, in the nondenitrifying cyanobacterium Synechocystis sp. strain PCC6803. Appl. Environ. Microbiol. 68, 668–672. Butland, G., Spiro, S., Watmough, N. J., and Richardson, D. J. (2001). Two conserved glutamates in the bacterial nitric oxide reductase are essential for activity but not assembly of the enzyme. J. Bacteriol. 183, 189–199. Carr, G. J., and Ferguson, S. J. (1990). The nitric oxide reductase of P. denitrificans. Biochem. J. 269, 423–429. Cheesman, M. R., Zumft, W. G., and Thomson, A. J. (1998). The MCD and EPR of the heme centers of nitric oxide reductase from Pseudomonas stutzeri: Evidence that the enzyme is structurally related to the heme-copper oxidases. Biochem. 37, 3994–4000. Cramm, R., Pohlmann, A., and Friedrich, B. (1999). Purification and characterization of the single-component nitric oxide reductase from Ralstonia eutropha H16. FEBS Lett. 460, 6–10. de Boer, A. P., van der Oost, J., Reijnders, W. N., Westerhoff, H. V., Stouthamer, A. H., and van Spanning, R. J. (1996). Mutational analysis of the nor gene cluster which encodes nitric-oxide reductase from P. denitrificans. Eur. J. Biochem. 242, 592–600. de Vries, S., Strampraad, M. J., Lu, S., Moenne-Loccoz, P., and Schroder, I. (2003). Purification and characterization of the MQH2:NO oxidoreductase from the hyperthermophilic archaeon Pyrobaculum aerophilum. J. Biol. Chem. 278, 35861–35868. Dermastia, M., Turk, T., and Hollocher, T. C. (1991). Nitric oxide reductase: Purification from P. denitrificans with use of a single column and some characteristics. J. Biol. Chem. 266, 10899–10905.

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Field, S. J., Prior, L., Roldan, M. D., Cheesman, M. R., Thomson, A. J., Spiro, S., Butt, J. N., Watmough, N. J., and Richardson, D. J. (2002). Spectral properties of bacterial nitric-oxide reductase: Resolution of pH-dependent forms of the active site heme b3. J. Biol. Chem. 277, 20146–20150. Flock, U., Watmough, N. J., and A¨delroth, P. (2005). Electron/proton coupling in bacterial nitric oxide reductase during reduction of oxygen. Biochemistry 44, 10711–10719. Gardner, A. M., Helmick, R. A., and Gardner, P. R. (2002). Flavorubredoxin, an inducible catalyst for nitric oxide reduction and detoxification in E. coli. J. Biol. Chem. 277, 8172–8177. Gill, S. R., Fouts, D. E., Archer, G. L., Mongodin, E. F., Deboy, R. T., Ravel, J., Paulsen, I. T., Kolonay, J. F., Brinkac, L., Beanan, M., Dodson, R. J., Daugherty, S. C., et al. (2005). Insights on evolution of virulence and resistance from the complete genome analysis of an early methicillin-resistant S. aureus strain and a biofilm-producing methicillinresistant S. epidermidis strain. J. Bacteriol. 187, 2426–2438. Girsch, P., and de Vries, S. (1997). Purification and initial kinetic and spectroscopic characterization of NO reductase from Paracoccus denitrificans. Biochim. Biophys. Acta 1318, 202–216. Gro¨nberg, K. L., Watmough, N. J., Thomson, A. J., Richardson, D. J., and Field, S. J. (2004). Redox-dependent open and closed forms of the active site of the bacterial respiratory nitric oxide reductase revealed by cyanide binding studies. J. Biol. Chem. 279, 17120–17125. Gro¨nberg, K. L. C., Roldań, M. D., Prior, L., Butland, G., Cheesman, M. R., Richardson, D. J., Spiro, S., Thomson, A. J., and Watmough, N. J. (1999). A lowredox potential heme in the dinuclear center of bacterial nitric oxide reductase: Implications for the evolution of energy-conserving heme-copper oxidases. Biochem. 38, 13780–13786. Hendriks, J., Gohlke, U., and Saraste, M. (1998a). From NO to OO: Nitric oxide and dioxygen in bacterial respiration. J. Bioenerg. Biomembr. 30, 15–24. Hendriks, J., Warne, A., Gohlke, U., Haltia, T., Ludovici, C., Lubben, M., and Saraste, M. (1998b). The active site of the bacterial nitric oxide reductase is a dinuclear iron center. Biochemistry 37, 13102–13109. Hendriks, J. H., Jasaitis, A., Saraste, M., and Verkhovsky, M. I. (2002). Proton and electron pathways in the bacterial nitric oxide reductase. Biochemistry 41, 2331–2340. Hendriks, J. H., Prior, L., Baker, A. R., Thomson, A. J., Saraste, M., and Watmough, N. J. (2001). Reaction of carbon monoxide with the reduced active site of bacterial nitric oxide reductase. Biochemistry 40, 13361–13369. Kastrau, D. H. W., Heiss, B., Kroneck, P. M. H., and Zumft, W. G. (1994). Nitric oxide reductase from Pseudomonas stutzeri: Phospholipid requirement, electron paramagnetic resonance and redox properties. Eur. J. Biochem. 222, 293–303. Leung, Y. C., Chan, C., Reader, J. S., Willis, A. C., van Spanning, R. J., Ferguson, S. J., and Radford, S. E. (1997). The pseudoazurin gene from Thiosphaera pantotropha: Analysis of upstream putative regulatory sequences and overexpression in E. coli. Biochem. J. 321, 699–705. Matsuda, Y., Inamori, K., Osaki, T., Eguchi, A., Watanabe, A., Kawabata, S., Iba, K., and Arata, H. (2002). Nitric oxide-reductase homologue that contains a copper atom and has cytochrome c-oxidase activity from an aerobic phototrophic bacterium Roseobacter denitrificans. J. Biochem. 131, 791–800. Nierman, W. C., DeShazer, D., Kim, H. S., Tettelin, H., Nelson, K. E., Feldblyum, T., Ulrich, R. L., Ronning, C. M., Brinkac, L. M., Daugherty, S. C., Davidsen, T. D., Deboy, R. T., et al. (2004). Structural flexibility in the Burkholderia mallei genome. Proc. Natl. Acad. Sci. USA 101, 14246–14251.

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Pauleta, S. R., Guerlesquin, F., Goodhew, C. F., Devreese, B., Van Beeumen, J., Pereira, A. S., Moura, I., and Pettigrew, G. W. (2004). Paracoccus pantotrophus pseudoazurin is an electron donor to cytochrome c peroxidase. Biochemistry 43, 11214–11225. Paulsen, I. T., Seshadri, R., Nelson, K. E., Eisen, J. A., Heidelberg, J. F., Read, T. D., Dodson, R. J., Umayam, L., Brinkac, L. M., Beanan, M. J., Daugherty, S. C., Deboy, R. T., et al. (2002). The Brucella suis genome reveals fundamental similarities between animal and plant pathogens and symbionts. Proc. Natl. Acad. Sci. USA 99, 13148–13153. Pearson, I. V., Page, M. D., van Spanning, R. J., and Ferguson, S. J. (2003). A mutant of Paracoccus denitrificans with disrupted genes coding for cytochrome c550 and pseudoazurin establishes these two proteins as the in vivo electron donors to cytochrome cd1 nitrite reductase. J. Bacteriol. 185, 6308–6315. Poock, S. R., Leach, E. R., Moir, J. W., Cole, J. A., and Richardson, D. J. (2002). Respiratory detoxification of nitric oxide by the cytochrome c nitrite reductase of E. coli. J. Biol. Chem. 277, 23664–23669. Richardson, D. J., and Watmough, N. J. (1999). Inorganic nitrogen metabolism in bacteria. Curr. Opin. Chem. Biol. 3, 207–219. Sakurai, N., and Sakurai, T. (1997). Isolation and characterization of nitric oxide reductase from Paracoccus halodenitrificans. Biochemistry 36, 13809–13815. Stevanin, T. M., Poole, R. K., Demoncheaux, E. A., and Read, R. C. (2002). Flavohemoglobin Hmp protects Salmonella enterica serovar typhimurium from nitric oxide-related killing by human macrophages. Infect. Immun. 70, 4399–4405. Suharti, R. C., Strampraad, M. J., Schroder, I., and de Vries, S. (2001). A novel copper A containing menaquinol NO reductase from Bacillus azotoformans. Biochem. 40, 2632–2639. Takaya, N., Catalan-Sakairi, M. A., Sakaguchi, Y., Kato, I., Zhou, Z., and Shoun, H. (2003). Aerobic denitrifying bacteria that produce low levels of nitrous oxide. Appl. Environ. Microbiol. 69, 3152–3157. Tho¨ny-Meyer, L., Kunzler, P., and Hennecke, H. (1996). Requirements for maturation of Bradyrhizobium japonicum cytochrome c550 in E. coli. Eur. J. Biochemistry. 235, 754–761. Thorndycroft, F. H., Butland, G., Richardson, D. J., and Watmough, N. J. (2007). A new assay for nitric oxide reductase reveals two conserved glutamate residues form the entrance to a proton-conducting channel in the bacterial enzyme. Biochem. J. 401, 111–119. Tunbridge, A. J., Stevanin, T. M., Lee, M., Marriott, H. M., Moir, J. W., Read, R. C., and Dockrell, D. H. (2006). Inhibition of macrophage apoptosis by Neisseria meningitidis requires nitric oxide detoxification mechanisms. Infect. Immun. 74, 729–733. van der Oost, J., deBoer, A. P. N., deGier, J.-W. L., Zumft, W. G., Stouthamer, A. H., and van Spanning, R. J. M. (1994). The heme-copper oxidase family consists of three distinct types of terminal oxidases and is related to nitric oxide reductase. FEMS Microbiol. Lett. 121, 1–9. Vincent, K. A., Tilley, G. J., Quammie, N. C., Streeter, I., Burgess, B. K., Cheesman, M. R., and Armstrong, F. A. (2003). Instantaneous, stoichiometric generation of powerfully reducing states of protein active sites using Eu(II) and polyaminocarboxylate ligands. Chem. Commun. 259, 0–2591. Watmough, N. J., Butland, G., Cheesman, M. R., Moir, J. W., Richardson, D. J., and Spiro, S. (1999). Nitric oxide in bacteria: Synthesis and consumption. Biochem. Biophys. Acta 1411, 456–474. Zumft, W. G. (2005). Nitric oxide reductases of prokaryotes with emphasis on the respiratory, heme-copper oxidase type. J. Inorg. Biochem. 99, 194–215.


C H A P T E R

S I X

Redox-Controlled Dinitrosyl Formation at the Diiron-Oxo Center of NorA Rainer Cramm and Katja Strube Contents 1. Introduction 2. Genetic Context and Expression of the NorA Gene in R. eutropha 3. Purification of NorA 3.1. Overproduction 3.2. Isolation procedures 4. Disulfide Bridges in NorA 5. Iron Analysis and Preparation of Apo-NorA 6. Interconversion of Redox Forms of NorA 6.1. Preparation of diferrous NorA 6.2. Preparation of diferric NorA and oxyNorA 7. Generation of NorA-DNIC In Vitro 7.1. Procedure using NO-saturated buffer 7.2. Procedure using nitrite 8. Preparation of NorA-DNIC Formed In Vivo 9. Quantification of NO from NorA-DNIC 10. Outlook References

104 105 106 106 106 107 108 109 110 110 111 111 112 113 113 114 114

Abstract In the denitrifying bacterium Ralstonia eutropha H16, the NorA protein is coproduced with the respiratory nitric oxide (NO) reductase. NorA contains a diiron-oxo center, which can form stable adducts with dioxygen and NO. In contrast to other diiron proteins, the formation of NorA-NO requires both fully reduced protein and additional electrons. A minor fraction of in vitro NorANO represents a paramagnetic dinitrosyl iron complex (DNIC), while the major fraction is attributed to a DNIC of the structure {Fe(NO)2},10 which shows no electron paramagnetic resonance. The NorA-DNIC may be formed either upon Institut fu¨r Biologie/Mikrobiologie, Humboldt-Universita¨t zu Berlin, Berlin, Germany Methods in Enzymology, Volume 437 ISSN 0076-6879, DOI: 10.1016/S0076-6879(07)37006-7

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direct reaction of the protein with NO or upon incubation with nitrite due to an intrinsic nitrite reduction activity of NorA that liberates NO. NorA can be purified rapidly as a six histidine-tagged derivative from overproducing cells of Escherichia coli. This chapter describes procedures for the preparation of different redox forms of NorA for the formation of NorA adducts with NO, dioxygen, and azide, as well as for the quantification of NorA-bound NO.

1. Introduction Binding of nitric oxide (NO) to iron protein plays an important role in physiological systems. In addition to nitrosation of heme proteins, e.g., hemoglobin and soluble guanylate cyclase (Henry and Guissani, 1999), NO has also been shown to interact with many nonheme iron sites to form stable Fe-NO complexes (Butler and Megson, 2002). Of these, dinitrosyl iron complexes (DNICs) in which two NO molecules are bound to one iron atom are of particular interest, as they are supposed to represent a transport form of NO (Chiang and Darensbourg, 2006; Keese et al., 1997; Ueno and Yoshimura, 2000; Vanin, 1998) and to act as potent NO donors (Vanin et al., 1996). DNICs are generally described as {Fe(NO)2}n (with n being the sum of d-type electrons from the metal and unpaired electrons from the NO) system according to Enemark–Feltham formalism (Enemark and Feltham, 1974). Note that DNICs differ from diiron-dinitrosyls of the structure [FeNO]2 that have been described for some diiron proteins, such as R2 protein of aerobic ribonucleotide reductase and methane monooxygenase (Coufal et al., 1999; Haskin et al., 1995). In bacteria, protein-bound DNICs have been implicated in regulatory mechanisms. In Escherichia coli, the [4Fe-4S] cluster of the oxygen sensor Fnr (Cruz-Ramos et al., 2002) and the [2Fe-2S] cluster of the superoxide sensor SoxR (Ding and Demple, 2000) are converted to monomeric and dimeric thiolate-ligated DNICs. Furthermore, the iron uptake regulator Fur, which contains a monoiron center, can react with NO to a major paramagnetic {Fe(NO)2}9 DNIC, proposed to originate from a diamagnetic {Fe(NO)2}8 DNIC intermediate (D’Autreaux et al., 2004). A paramagnetic {Fe(NO)2}9 DNIC has also been detected upon binding of NO to the diiron protein NorA from the denitrifying bacterium Ralstonia eutropha H16 (Strube et al., 2007). However, this species represented a minor fraction (up to 20%), while the major species was an electron paramagnetic resonance (EPR)-silent dinitrosyl complex, most probably of the form {Fe(NO)2}10. Notably, and in contrast to Fur, DNIC formation at NorA is dependent on the supply of external electrons. It is unknown, however, whether the activity of NorA in vivo is controlled by a specialized redox system or if NorA is reduced by an unspecific electron donor.

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Dinitrosyl Formation at a Diiron-Oxo Center

2. Genetic Context and Expression of the NorA Gene in R. eutropha In R. eutropha, the norA structural gene is coexpressed with norB encoding the respiratory nitric oxide reductase (Bu¨sch et al., 2004; Pohlmann et al., 2000). The transcriptional activator NorR is encoded upstream of norA and is transcribed in the opposite direction (Fig. 6.1). In fact, the wild-type H16 contains a second set of the norRAB genes termed norR2A2B2 (Pohlmann et al., 2006). Products of the latter genes have not yet been characterized. Note that all physiological experiments related to NorR, NorA, or NorB were performed using a deletion mutant that lacks the norR2A2B2 set. NorR belongs to a subgroup of the NifA/NtrC family of regulators, which interact directly with their signal molecules (Studholme and Dixon, 2003). NorR orthologs are encoded in several bacterial genomes, including E. coli (Rodionov et al., 2005). E. coli NorR has been shown to bind NO by a monoiron center, which is coordinated by the N-terminal domain of the protein (D’Autreaux et al., 2005). Putative ligands of the iron have been identified for R. eutropha NorR by site-directed mutagenesis (Klink et al., 2007). In its active state, NorR activates expression of the norAB operon (Bu¨sch et al., 2004). The subsequent formation of nitric oxide reductase NorB is essential for denitrifying cells, probably because NO accumulates in

norR

− + norA

NO ON NO NorR

norB

N2O

NorA

NorB 2 NO

NO

Figure 6.1 Genetic context of the gene for NorA in R. eutropha. The regulator NorR activates expression of the norAB operon in response to NO. The norR gene is autoregulated negatively.The respiratory nitric oxide reductase NorB is instrumental in denitrification of R. eutropha.

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the cell to toxic concentrations if NorB is lacking (Cramm et al., 1997). In contrast, cells lacking NorA did not show a growth defect during denitrification (Pohlmann et al., 2000). Nevertheless, formation of the NorA-NO complex appears to be of physiological relevance. The reaction triggers a feedback loop, as NO-dependent activation of NorR (and hence expression of the norAB operon) is decreased when NO is scavenged by NorA (Strube et al., 2007).

3. Purification of NorA 3.1. Overproduction To allow isolation of NorA by a rapid one-step method (e.g., to purify the NO-bound form of NorA from cell cultures), NorA has been modified genetically to contain six extra histidine residues at its C terminus (His tag). R. eutropha NorA can be overproduced to high yields in E. coli BL21(DE3) or Rosetta(DE3) cells containing a pET22b-based overexpression plasmid. Fresh transformants should be used for each preparation. About 10 mg NorA can be easily prepared from a 1-liter culture grown in LB medium. Ten milliliters of an overnight culture is used to inoculate 1 liter of LB medium, freshly prepared in a baffled 5-liter Erlenmeyer flask. The culture is incubated at 30 with shaking (140 rpm) for 3 h to reach an optical density at 600 nm of 0.6 (3 h). Overexpression of norA is induced by the addition of isopropyl-b-D-thiogalactopyranoside to a final concentration of 1 mM, and incubation is continued for another 3 to 4 h at the aforementioned conditions. The culture is then chilled on ice, and cells are harvested by centrifugation at 5000 g for 15 min.

3.2. Isolation procedures Although NorA is formed during denitrification (and thus under conditions of reduced oxygen tension), the protein is fairly stable in the presence of air. Thus the isolation procedure may be carried out under aerobic conditions. Note, however, that during aerobic isolation, different redox states of the diiron center may form (see later) and an increased fraction of dimeric NorA will appear due to the formation of disulfide bridges. Therefore, it is advisable to perform the isolation of NorA in an anaerobic chamber. The pellet of a 1-liter culture is resuspended in 5 ml 50 mM sodium phosphate buffer, pH 8.0, 300 mM NaCl, and 20 mM imidazole, and cells are immediately broken by sonication on ice. Debris and unbroken cells are then removed by centrifugation at 12,000 g for 15 min, and the supernatant


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(soluble extract) is transferred to a clean tube. His-tagged NorA is purified from the soluble extract by affinity chromatography using commercial nickel-chelating resin (nitrilotriacetic acid) and standard buffers (50 mM sodium phosphate buffer, pH 8.0, 300 mM NaCl) containing the following concentrations of imidazole: equilibration, 20 mM; washing, 50 mM; and elution, 250 mM. Eluted fractions are combined and concentrated using a microfiltrator (Millipore, Microcon centrifugal filter device, cutoff size 10 kDa). Typically, this procedure will yield about 1 ml containing 300 to 400 nmol hexahistidine-tagged NorA. Such preparations are 95% pure, as judged by SDS-PAGE and subsequent staining with Coomassie brilliant blue. If necessary, further purification steps may be added: the sample is desalted using PD-10 columns (GE Healthcare) and applied to an anion-exchange column (ResourceQ, GE Healthcare) preequilibrated with 50 mM sodium phosphate buffer, pH 8.0, and 10 mM NaCl. NorA elutes at approximately 150 mM NaCl in a linear gradient ranging from 10 to 500 mM NaCl in 50 mM sodium phosphate buffer (pH 8.0). Purified NorA is used immediately for experiments. For storage, glycerol is added to a final concentration of 20% (v/v). Aliquots are then frozen in liquid nitrogen and stored at –80 .

4. Disulfide Bridges in NorA If isolated under aerobic conditions, preparations contain aggregated NorA species that are readily detected in nonreducing SDS-PAGE. The amount of a dimeric NorA species increases upon prolonged exposure of such preparations to air (Fig. 6.2). After overnight incubation at 4 , up to 20% of NorA is detectable as a homodimer. NorA contains four cysteine residues, three of which are conserved according to a multiple alignment of NorA orthologs from different bacteria (Strube et al., 2007). Two of these are part of the conspicuous CCGG sequence within the N-terminal domain of NorA. After anaerobic preparation of NorA from aerobically grown cells of E. coli, all of the four cysteines possess free thiol groups, according to a titration with 5,50 -dithiobis(2-nitrobenzoic acid) (DTNB) of the native protein. Therefore, all four cysteines are solvent accessible, and NorA does not form disulfide-bridged homodimers in the cell. Interestingly, only two free thiols can be detected by DTNB titration after prolonged incubation of NorA in air. Because only a monomeric and a dimeric form are resolved by SDS-PAGE, this result has to be explained by an additional intramolecular cysteine bridge, which, however, does not lead to an altered mobility of the protein in standard gels.

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A

S

1

2

B

S

1

2

83 62 47.5

32.5

Figure 6.2 Disulfide bridging in NorA. NorA (10 mg) aerobically purified (A) and after aerobic incubation overnight at 4 (B) in the absence (lane 1) or in the presence (lane 2) of 10 mM DTT, separated in a12% SDS-PAGE gel. Molecular weights of protein standard (lanes S) are shown on the left side.

5. Iron Analysis and Preparation of Apo-NorA Selection of an appropriate method for chemical iron determination is crucial in avoiding underestimation of the iron content of NorA. For example, the chelator ferene S turned out to be less suitable even if the incubation periods were extended considerably. Good results are obtained with a rapid and simple method using the iron (II) chelator 2,20 -bipyridyl. In an anaerobized rubber-sealed reaction tube, 100 ml NorA (0.01–0.3 mM ) is reduced with excess sodium dithionite for 5 min. Subsequently, 0.1 ml of a 20 mM 2,20 -bipyridyl stock solution is added and the sample is vortexed vigorously for 5 min. During this time, the red-pink bipyridyl-iron complex is formed. Protein is removed prior to iron quantification using a microfiltrator (Millipore, Microcon centrifugal filter device, cutoff size 10 kDa). Absorption of the filtrate is measured at 520 nm, and iron is quantified using an absorption coefficient of e ¼ 8000 M1 cm1. A similar method is used to prepare apo-NorA: 0.5 ml of 0.3 mM purified NorA is mixed with 10 mM ascorbate and 10 mM phenazine methosulfate (PMS) in a reaction tube placed in an anaerobic chamber. Iron is chelated by the addition of 0.1 ml of a 20 mM 2,20 -bipyridyl stock solution. After incubation for 1 h with gentle shaking, the solution is loaded onto a PD-10 desalting column (GE Healthcare) and eluted with 50 mM sodium phosphate buffer, pH 8.0, 300 mM NaCl. The eluate contains iron-free apo-NorA with no detectable absorption above 300 nm.

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6. Interconversion of Redox Forms of NorA Binuclear iron clusters may adopt three redox states: (i) fully oxidized, (ii) fully reduced, or (iii) mixed valent. A fully reduced diiron center is a prerequisite for the formation of dioxygen or nitric oxide adducts of NorA. Solutions of diferrous NorA are colorless, and the protein shows only weak (if any) signals in both EPR spectra and optical spectra above 300 nm (Fig. 6.3). As expected for a diferric diiron-(hydr)oxo center, fully oxidized NorA is EPR silent because of antiferromagnetic coupling. Solutions of diferric NorA are pale yellow. In the optical spectrum the protein shows a prominent signal at 353 nm (e 4500 M1 cm1) and a shoulder at 430 nm (see Fig. 6.3, inset trace a). Diferric NorA combines readily with azide, yielding a light orange product with characteristic signals at 327 and 453 and a shoulder at 490 nm (see Fig. 6.3, inset trace b). Frequently, in aged preparations of NorA, a minor fraction gives rise to an S ¼ 1/2 EPR signal with gz¼ 1.9665, gy¼ 1.9230, gx ¼ 1.8732 (below 30 K), which is because of a mixed-valence center (Strube et al., 2007). Mixed-valent NorA is not detectable in samples prepared freshly under anaerobic conditions.

0.4 Absorbtion (arbitrary units)

Absorbtion (arbitrary units)

0.8

0.6 b 0.4

0.2

0.3 0.2

b

0.1 a 0.0 300

400


700

800

a

c 0.0 300

400

500 Wavelength (nm)

600

Figure 6.3 Absorption spectrum of anaerobically purified NorA.Trace a:150 mM NorA isolated anaerobically; trace b: sample from trace a exposed to air for 60 min; trace c: addition of a few crystals of dithionite to sample of trace a. (Inset) Trace a, 70 mM air-exposed NorA incubated overnight; trace b, sample from trace a incubated with excess azide (10 mM ) for 1 min.

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6.1. Preparation of diferrous NorA Purification under anaerobic conditions yields mainly diferrous NorA. However, depending on the concentration of residual oxygen during the isolation procedure, such NorA preparations may also contain traces of oxy- and diferric NorA. A homogeneous sample of diferrous NorA is prepared by the addition of reductant (ascorbate/PMS or dithionite, cf. Fig. 6.3, trace c and Fig. 6.4, trace a) and subsequent recovery of NorA by dialysis or gel filtration. A convenient procedure is as follows: later. In an anaerobized cuvette sealed with a rubber stopper, 300 mM NorA is reduced by the addition of 10 mM ascorbate and 10 mM PMS. Bleaching of NorA absorbance may be monitored by optical spectroscopy (keep in mind the intrinsic absorbance of the reducing compounds). Subsequently, the solution is applied to a PD-10 column (GE Healthcare) placed in an anaerobic chamber. Note that after removal of the reductant, traces of oxygen present in the environment will immediately lead to formation of some oxyNorA.

6.2. Preparation of diferric NorA and oxyNorA Diferric NorA can be prepared easily from oxyNorA by a simple incubation step. To prepare oxyNorA, a sample of diferrous NorA (devoid of artificial reductant) is exposed to air at room temperature for 30 to 60 min.


0.14


0.5

0.4

0.3 b

0.2

0.12 0.10 0.08 0.06 0.04 0.02 0.00 400

500


800

0.1 a 0.0 400

500

600

700

800

900

Wavelength (nm)

Figure 6.4 Absorption spectrum of NorA-NO. NorA (150 mM) was incubated in a cuvette closed with a rubber septum and reduced with 10 mM ascorbate and 10 mM PMS (trace a). Subsequently, NO-saturated buffer (400 mM NO) was added. (Inset) NorA-NO (110 mM) as isolated from E. coli cell cultures treated with 10 mM NO.


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Only minimal amounts of dioxygen are required; formation of oxyNorA was observed even in an anaerobic cuvette closed with a Teflon stopper due to the presence of dioxygen adhered to the Teflon material. OxyNorA is yellow-red in solution and is characterized by a 520-nm band (e ¼ 1153 M1 cm1) in the optical spectrum (see Fig. 6.3, trace b). Because the region between 400 and 550 nm contains bands for ligand-to-iron charge transfer transitions with noninnocent ligands such as peroxide (Reem et al., 1989), it is feasible that oxygen is bound as a hydroperoxo-Fe3þ adduct, similar to oxygen binding by hemerythrin (Stenkamp, 1994). Upon prolonged aerobic incubation on ice, oxyNorA autooxidizes probably by a mechanism similar to the formation of diferric hemerythrin from oxygenized hemerythrin by loss of the peroxide ligand (Stenkamp, 1994). After an overnight incubation at 4 , NorA is recovered quantitatively in the diferric state. Diferric NorA is readily distinguished from diferrous NorA by the addition of azide, as only the diferric form reacts to an orange adduct (see Fig. 6.3, inset, trace b).

7. Generation of NorA-DNIC In Vitro 7.1. Procedure using NO-saturated buffer In general, the addition of NO-saturated buffer to stirred NorA solutions in rubber-sealed cuvettes is superior to the addition of NO gas. The latter procedure leads to high local concentrations of NO, which may result in the precipitation of protein. Aqueous solutions saturated with nitric oxide (1.9 mM ) can be prepared by bubbling nitric oxide gas through pure water under anaerobic conditions. An alkaline trap should be connected upstream of the water vial to remove traces of certain other N-oxides. If the system is not anaerobized properly, the NO solution will finally contain considerable amounts of nitrite. For experiments with NorA, this would be highly undesirable, as the protein also reacts with nitrite (see later). It is advisable to check NO solutions by optical spectroscopy. Pure NO solutions will show no absorption above 300 nm, while contaminations (nitrite or other oxidation products of NO) give additional absorption between 300 and 400 nm. For the reaction of NorA with nitric oxide, 200 ml of 150 mM diferrous NorA is transferred to an anaerobic microcuvette that contains a small stirrer bar and is sealed with a rubber septum. This step should be carried out in an anaerobic chamber to prevent oxidation of both reactants. Subsequently, 1 mM of ascorbate and catalytic amounts of PMS (1 mM) are added with a gas-tight syringe and the cuvette is placed on a stirring device, ideally integrated into the cuvette holder of a spectrophotometer. After an

112


incubation period of 1 min at room temperature, 20 ml of a saturated NO solution is injected with a gas-tight syringe. This procedure is repeated three times to provide an excess of NO. The resulting NorA-DNIC is greenish and is characterized by the formation of two specific signals at 420 and 750 nm in the optical spectrum (Fig. 6.4, trace b). The absorption coefficient of the 750-nm band is E ¼ 126 M1 cm1.

7.2. Procedure using nitrite

NorB

NO (nmol)

30

NO (nmol)

In the presence of reductant, NorA is able to catalyze a limited number of nitrite reduction cycles, yielding NO as the end product. After the reaction is finished, the optical spectrum of the sample is indistinguishable from NorA-DNIC as prepared with NO-saturated buffer. Thus the catalytic turnover of nitrite is probably product inhibited as a consequence of NorA-DNIC formation. Nevertheless, excess nitric oxide is formed during this reaction, which can be monitored by amperometric methods using NO-sensitive electrodes (Fig. 6.5). Nitrite reduction by NorA is slow, and high amounts of nitrite are required to follow this reaction as the Km of NorA for nitrite is around 7 mM. The reaction is best followed in phosphate buffer, pH 8.0. Lower pH values increase the chemical formation of NO from nitrous acid, which may cover the NorA-derived reaction at some point. Higher pH values decrease the activity of NorA.

30 15 0 0

20

4 8 12 Time (min)

10 0 0

2

4 6 8 Time (min)

10

Figure 6.5 Nitrite reduction to NO. NO formation by NorA (3 mM) was followed in a Clark electrode. The reaction buffer was 50 mM sodium phosphate buffer (pH 7.0), 20 mM D-glucose,10 units of glucose oxidase, 250 units of catalase,10 mM ascorbate, and 10 mM PMS.The reaction volume was 2 ml. Bold face arrows denote addition of 2.5 mM nitrite. After incubation for 5 min, 250 nM purified NO reductase NorB of R. eutropha was added (indicated by a thin arrow) to verify that the electrode signal was caused by NO. (Inset) An experiment without adding NO reductase.


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8. Preparation of NorA-DNIC Formed In Vivo NorA-DNIC is formed in vivo (see Fig. 6.4, inset) and can be isolated from NO-exposed cultures of E. coli. NorA is overproduced as described earlier except that cells are transferred to a sealed bottle 1.5 h after induction. NO-saturated buffer (1.9 mM NO) is added to the culture to a final concentration of 10 mM. Cells are exposed to NO for 10 min. NorA is isolated in an anaerobic chamber by a rapid procedure (10 min) using commercial nickel-chelate spin columns (Qiagen) equilibrated with phosphate buffer, pH 8.0, 300 mM NaCl, 20 mM imidazole. Spin steps are carried out at a low centrifugal force (2000 rpm) for 1 min. Soluble extracts are passed three times through the spin columns to increase the total yield. After extensive washing (phosphate buffer, pH 8.0, 300 mM NaCl, 20 mM imidazole), the columns are eluted with phosphate buffer, pH 8.0, 300 mM NaCl, 250 mM imidazole. To save time, further purification steps are omitted and the protein is transferred to a rubber-sealed cuvette. In the absence of reductant, NorA-DNIC is sensitive toward exposure to oxygen. Assuming that the initial amount of NorA-DNIC in the cell is 100%, a half-time of about 20 min is calculated for the decay of the DNIC in air.

9. Quantification of NO from NorA-DNIC The following procedure has to be carried out in an anaerobic chamber to avoid loss of NO by air/oxidation. NorA (50 to 200 mM) is incubated with three equivalents of NO, 1 mM ascorbate, and 10 mM PMS in a sealed vial without headspace for 15 min. Alternatively, a sealed cuvette may be used to monitor formation of the DNIC by optical spectroscopy. Using a gas-tight syringe, the solution is then transferred to a sealed microfiltrator (Millipore, Microcon centrifugal filter device, cutoff size 10 kDa) and centrifuged at 12,000g for 1 min to obtain some protein-free filtrate. NO is quantified in a modified Griess assay. Aliquots from the retenate and the filtrate are injected into rubber-sealed reaction tubes that contain 300 ml Griess reagent under an atmosphere of dioxygen. Under these conditions, NO is oxidized to nitrite, which is quantified in the Griess reaction by measuring the absorption at 550 nm. Note that the retenate (containing NorA-DNIC) has to be incubated for about 5 h for quantitative analysis, perhaps as a consequence of slow conversion of the DNIC into nitrite (Boese et al., 1995). In agreement with the initial NorA/NO ratio of 1/3 and DNIC formation by NorA, a ratio of 1.8 (0.15) molecules NO per diiron center versus 0.8 (0.1) molecules free NO is determined by this procedure.

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As a control, NO-exposed apo-NorA may be used in this assay to exclude contamination with unspecifically bound NO or nitrite.

10. Outlook Orthologs of NorA are encoded in many proteobacteria and firmicutes and have been termed YtfE, DnrN, ScdA, or NipC. In most cases where it has been studied, these proteins are formed in response to nitrosative stress ( Justino et al., 2005; Kim et al., 2003; Pohlmann et al., 2000; Sebbane et al., 2006; Vollack and Zumft, 2001). The physiological role of NorA and its orthologs is still enigmatic. In E. coli, YtfE has been shown to be involved in the response toward oxidative and nitrosative stress ( Justino et al., 2005, 2006). In particular, the activity of the [Fe-S] containing nitrate reductase NAR was decreased greatly in the YtfE mutant. It was demonstrated that YtfE facilitates the repair of damaged [4Fe-4S] centers, and a role of YtfE in iron insertion was suggested ( Justino et al., 2007). However, no molecular basis for this activity is yet available. In contrast, the growth of denitrifying (and thus NO-exposed) R. eutropha NorA mutant cells was not impaired significantly (Pohlmann et al., 2000; Strube et al., 2007), showing that the activity of the NAR enzyme in R. eutropha does not depend on NorA. Therefore, it has to be considered that, albeit both proteins are 50% identical in their amino acid sequence, NorA and YtfE serve different physiological functions.

REFERENCES Boese, M., Mordvintcev, P. I., Vanin, A. F., Busse, R., and Mulsch, A. (1995). S-nitrosation of serum albumin by dinitrosyl-iron complex. J. Biol. Chem. 270, 29244–29249. Bu¨sch, A., Pohlmann, A., Friedrich, B., and Cramm, R. (2004). A DNA region recognized by the nitric oxide-responsive transcriptional activator NorR is conserved in beta- and gamma-proteobacteria. J. Bacteriol. 186, 7980–7987. Butler, A. R., and Megson, I. L. (2002). Non-heme iron nitrosyls in biology. Chem. Rev. 102, 1155–1166. Chiang, C. Y., and Darensbourg, M. Y. (2006). Iron nitrosyl complexes as models for biological nitric oxide transfer reagents. J. Biol. Inorg. Chem. 11, 359–370. Coufal, D. E., Tavares, P., Pereira, A. S., Hyunh, B. H., and Lippard, S. J. (1999). Reactions of nitric oxide with the reduced non-heme diiron center of the soluble methane monooxygenase hydroxylase. Biochemistry 38, 4504–4513. Cramm, R., Siddiqui, R. A., and Friedrich, B. (1997). Two isofunctional nitric oxide reductases in Alcaligenes eutrophus H16. J. Bacteriol. 179, 6769–6777.


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Cruz-Ramos, H., Crack, J., Wu, G., Hughes, M. N., Scott, C., Thomson, A. J., Green, J., and Poole, R. K. (2002). NO sensing by FNR: Regulation of the E. coli NO-detoxifying flavohaemoglobin, Hmp. EMBO J. 21, 3235–3244. D’Autreaux, B., Horner, O., Oddou, J. L., Jeandey, C., Gambarelli, S., Berthomieu, C., Latour, J. M., and Michaud-Soret, I. (2004). Spectroscopic description of the two nitrosyl-iron complexes responsible for fur inhibition by nitric oxide. J. Am. Chem. Soc. 126, 6005–6016. D’Autreaux, B., Tucker, N. P., Dixon, R., and Spiro, S. (2005). A non-haem iron centre in the transcription factor NorR senses nitric oxide. Nature 437, 769–772. Ding, H., and Demple, B. (2000). Direct nitric oxide signal transduction via nitrosylation of iron-sulfur centers in the SoxR transcription activator. Proc. Natl. Acad. Sci. USA 97, 5146–5150. Enemark, J. H., and Feltham, R. D. (1974). Principles of structure, bonding and reactivity for metal nitrosyl complexes. Coord. Chem. Rev. 13, 339–406. Haskin, C. J., Ravi, N., Lynch, J. B., Munck, E., and Que, L., Jr. (1995). Reaction of NO with the reduced R2 protein of ribonucleotide reductase from E. coli. Biochemistry 34, 11090–11098. Henry, Y., and Guissani, A. (1999). Interactions of nitric oxide with hemoproteins: Roles of nitric oxide in mitochondria. Cell. Mol. Life Sci. 55, 1003–1014. Justino, M. C., Almeida, C. C., Goncalves, V. L., Teixeira, M., and Saraiva, L. M. (2006). Escherichia coli YtfE is a di-iron protein with an important function in assembly of ironsulphur clusters. FEMS Microbiol. Lett. 257, 278–284. Justino, M. C., Almeida, C. C., Teixeira, M., and Saraiva, L. M. (2007). Escherichia coli di-iron YtfE protein is necessary for the repair of stress-damaged iron-sulfur clusters. J. Biol. Chem. 282, 10352–10359. Justino, M. C., Vicente, J. B., Teixeira, M., and Saraiva, L. M. (2005). New genes implicated in the protection of anaerobically grown E. coli against nitric oxide. J. Biol. Chem. 280, 2636–2643. Keese, M. A., Bose, M., Mulsch, A., Schirmer, R. H., and Becker, K. (1997). Dinitrosyldithiol-iron complexes, nitric oxide (NO) carriers in vivo, as potent inhibitors of human glutathione reductase and glutathione-S-transferase. Biochem. Pharmacol. 54, 1307–1313. Kim, C. C., Monack, D., and Falkow, S. (2003). Modulation of virulence by two acidified nitrite-responsive loci of Salmonella enterica serovar typhimurium. Infect. Immun. 71, 3196–3205. Klink, A., Elsner, B., Strube, K., and Cramm, R. (2007). Characterization of the signaling domain of the NO-responsive regulator NorR from R. eutropha H16 by site-directed mutagenesis. J. Bacteriol. 189, 2743–2749. Pohlmann, A., Cramm, R., Schmelz, K., and Friedrich, B. (2000). A novel NO-responding regulator controls the reduction of nitric oxide in R. eutropha. Mol. Microbiol. 38, 626–638. Pohlmann, A., Fricke, W. F., Reinecke, F., Kusian, B., Liesegang, H., Cramm, R., Eitinger, T., Ewering, C., Potter, M., Schwartz, E., Strittmatter, A., Voss, I., et al. (2006). Genome sequence of the bioplastic-producing ‘‘Knallgas’’ bacterium R. eutropha H16. Nat. Biotechnol. 24, 1257–1262. Reem, R. C., McCormick, J. M., Richardson, D. E., Devlin, F. J., Stephens, P. J., Musselman, R. L., and Solomon, E. I. (1989). Spectroscopic studies of the coupled binuclear ferric active site in methemerythrins and oxyhemerythrin: The electronic structure of each iron center and the iron-oxo and iron-peroxide bonds. J. Am. Chem. Soc. 111, 4688–4704. Rodionov, D. A., Dubchak, I. L., Arkin, A. P., Alm, E. J., and Gelfand, M. S. (2005). Dissimilatory metabolism of nitrogen oxides in bacteria: Comparative reconstruction of transcriptional networks. PLoS Comput. Biol. 1, e55.

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Sebbane, F., Lemaitre, N., Sturdevant, D. E., Rebeil, R., Virtaneva, K., Porcella, S. F., and Hinnebusch, B. J. (2006). Adaptive response of Yersinia pestis to extracellular effectors of innate immunity during bubonic plague. Proc. Natl. Acad. Sci. USA 103, 11766–11771. Stenkamp, R. E. (1994). Dioxygen and hemerythrin. Chem. Rev. 94, 715–726. Strube, K., de Vries, S., and Cramm, R. (2007). Formation of a dinitrosyl iron complex by NorA, a nitric oxide binding di-iron protein from Ralstonia eutropha H16. J. Biol. Chem. 282, 20292–20300. Studholme, D. J., and Dixon, R. (2003). Domain architectures of s 54-dependent transcriptional activators. J. Bacteriol. 185, 1757–1767. Ueno, T., and Yoshimura, T. (2000). The physiological activity and in vivo distribution of dinitrosyl dithiolato iron complex. Jpn. J. Pharmacol. 82, 95–101. Vanin, A. F. (1998). Dinitrosyl iron complexes and S-nitrosothiols are two possible forms for stabilization and transport of nitric oxide in biological systems. Biochemistry (Mosc.) 63, 782–793. Vanin, A. F., Stukan, R. A., and Manukhina, E. B. (1996). Physical properties of dinitrosyl iron complexes with thiol-containing ligands in relation with their vasodilator activity. Biochem. Biophys. Acta 1295, 5–12. Vollack, K. U., and Zumft, W. G. (2001). Nitric oxide signaling and transcriptional control of denitrification genes in Pseudomonas stutzeri. J. Bacteriol. 183, 2516–2526.

C H A P T E R

S E V E N

Purification and Functional Analysis of Fungal Nitric Oxide Reductase Cytochrome P450nor Li Zhang* and Hirofumi Shoun† Contents 118 119 119 120 121 122 123 124 125 126 127 128 130 131 131 131 131

1. Introduction 2. Screening of P450nor Activity 3. Gas Analysis 4. Purification of P450nor 5. Nitric Oxide Reductase Activity Assay 6. Protein Sequencing 7. Isolation of cDNA 8. Subcellular Fractionation of T. cutaneum 9. Site-Directed Mutagenesis 10. Expression of Recombinant Proteins 11. Purification of Recombinant Proteins 12. Titration of NAD Analogs 13. Stopped-Flow Rapid Scan Analysis 14. Other Analysis 15. Conclusion Acknowledgments References

Abstract Cytochrome P450nor (P450nor) is a nitric oxide (NO) reductase involved in fungal denitrification. Denitrification is a biological process in which nitrate or nitrite is reduced to gaseous nitrogen, the reverse reaction of nitrogen fixation. It therefore plays an important role in maintaining global environmental homeostasis. The involvement of P450nor in fungal denitrification indicates that denitrification not only occurs in prokaryotic bacteria, but also in eukaryotic fungi. In addition, the reduction of NO to nitrous oxide catalyzed by P450nor has added new insight into the function of cytochrome P450s, which are usually

* {

Department of Biology, University of Kentucky, Lexington, Kentucky Department of Biotechnology, University of Tokyo, Bunkyo-ku, Tokyo, Japan

Methods in Enzymology, Volume 437 ISSN 0076-6879, DOI: 10.1016/S0076-6879(07)37007-9

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Li Zhang and Hirofumi Shoun

monooxygenases. Currently, five isozymes of P450nor have been isolated from the subdivisions of eumycota, and studies on the function and structure of P450nor have provided important information for both molecular mechanisms of P450 reactions and wastewater treatment. This chapter describes the screening of NO reductase activities, cloning, purification, and functional analysis of P450nor.

1. Introduction Fungal nitric oxide reductase cytochrome P450nor (P450nor) is a unique heme-thiolate protein involved in fungal denitrification by reducing nitric oxide (NO) to nitrous oxide (N2O) (for reviews, see Daiber et al., 2005; Shoun, 2006; Zhang et al., 2002a). Unlike other cytochrome P450s, P450nor is a reductase without monooxygenase activity, although it belongs to the P450 super family (CYP55A). It receives electrons directly from the distal pocket from NADH and/or NADPH without any help from a flavoprotein such as P450 reductase (Nakahara et al., 1993). The overall reaction is described in the following equation:

2NO þ NADðPÞH þ Hþ ! N2 O þ NADðPÞþ þH2 O P450nor was originally isolated from the filamentous fungus Fusarium oxysporum (Shoun and Tanimoto, 1991), indicating for the first time that denitrification is not restricted in prokaryotes. Denitrification is a reverse procedure of nitrogen fixation, which plays crucial roles in the nitrogen cycle of the ecosystem. As part of the whole denitrification procedure, fungal denitrification contains reduction steps of nitrogen sources from nitrate to nitrous oxide. P450nor plays a central role in the last step by catalyzing the reduction of NO to N2O. Knockout of P450nor from F. oxysporum results in the defect of denitrification activity, indicating that P450nor is necessary for fungal denitrification (Takaya and Shoun, 2000). We have isolated and cloned five isoforms of P450nor, which all belong to the CYP55A family: Fnor (CYP55A1) from F. oxysporum (Kizawa et al., 1991; Nakahara and Shoun, 1996), Cnor1 (CYP55A2) and Cnor2 (CYP55A3) from Cylindrocarpon tonkinense (Kudo et al., 1996; Usuda et al., 1995) (both fungi are the anamorph generation of ascomycetous fungi), Tnor (CYP55A4) from basidiomycetous yeast Trichosporon cutaneum (Zhang et al., 2001), and Anor (CYP55A5) from another ascomycetous fungus Aspergillus oryzae (Kaya et al., 2004). Therefore, P450nor activity is widely distributed in the subdivisions of eumycota.

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The protocols described herein have been used successfully to clone and purify fungal P450nor from yeast. This chapter also includes protocols for the functional analysis of this novel P450.

2. Screening of P450nor Activity Denitrification activities were screened successfully from soil fungi and yeasts (Shoun et al., 1992; Tsuruta et al., 1998). Yeast strains are obtained from the Japanese Collection of Microorganisms ( JCM, RIKEN, Japan). Potato dextrose (PD) broth (Difco; pH 7.4) is used in preculture. Each strain is inoculated into 120 ml of PD broth in a 500-ml Erlenmeyer flask sealed with a cotton plug for 3 days on a rotary shaker (120 rpm). Glycerol-peptone medium is used for detecting the denitrifying activity by anaerobic incubation at 30 or 25 (Candida sp. is incubated at 15 because of its isolation from Antarctica) for 4 days: 3% glycerol, 0.2% peptone, 10 mM sodium nitrite (or nitrate), a trace element solution (1 ml/liter), which contains the following chemicals (in 1 liter of distilled water): 0.2 g CoCl2H2O, 0.2 g FeSO47H2O, 1.0 g FeCl26H2O, 4.0 g CuSO45H2O, 0.2 g CaCl2, 8.6 mg Na2MoO42H2O, and 0.2 mg Na2SeO3. In brief, every 40 ml of preculture is inoculated into 120 ml of the aforementioned medium in a 500-ml Erlenmeyer flask with two side arms, which is sealed with a rubber stopper after replacing the headspace air with argon, and then incubated on a rotary shaker (120 rpm).

3. Gas Analysis An aliquot (usually 0.5 ml) of the upper-space gas in each flask is collected with a syringe through the side arm after incubation. The amount of gas (N2O) formed is determined by gas chromatography with a Shimadzu gas chromatograph GC 12A equipped with a Porapack Q column (3 2 mm, inner diameter) and by isotope mass spectrometry with a Finnigan Delta plus isotope mass spectrometer when necessary. Peaks are identified and quantified by comparison with standard gases (GL Sciences, Tokyo, Japan). [15N]nitrite (99 atom %) can be obtained from Shoko-Tsusho (Tokyo). A typical result is shown in the work of Tsuruta et al. (1998). Different from bacterial denitrification, which usually contains the whole process from nitrate to nitrogen (NO3 ! N2), most of the yeast strains that exhibit denitrifying activity use only the minimal process of denitrification from nitrite to nitrous oxide (NO2 ! N2O).

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4. Purification of P450nor Because P450nor is typically detectable in yeast T. cutaneum, it was used to purify P450nor (Zhang et al., 2001). Preculture is prepared by seeding the yeast in 300 ml of YPD medium (10 g/liter glucose, 10 g/liter peptone, and 5 g/liter yeast extract, pH 7.4) in a 500-ml flask and incubating at 30 for 24 h at 100–120 rpm. The whole preculture is then transferred to 3 liters of YPD medium in a 5-liter conical flask and incubated in the same conditions as the preculture. Wet cells of yeast are filter collected and kept at –80 until use. In contrast to Fnor in F. oxysporum, Tnor does not require any induction by nitrogen sources. Therefore, nitrite or nitrate can be omitted from the whole incubation. Wet cells are disrupted at 4 by grinding with aluminum oxide in 0.1 M potassium phosphate buffer (pH 7.15) containing 10% (v/v) glycerol, 0.1 mM EDTA, 0.1 mM dithiothreitol (DTT), and each of 0.25 mM protease inhibitors, phenylmethylsulfonyl fluoride (PMSF), leupeptin, and tosylphenylalanyl chloromethyl ketone (TPCK). The cell homogenate is subjected to sequential centrifugation at 1500g for 15 min, at 10,000g for 20 min, and at 105,000g for 1 h. The resulting supernatant (soluble fraction) is dialyzed overnight against buffer A (10 mM potassium phosphate buffer, pH 7.3, containing 10% glycerol, 0.1 mM EDTA, and 0.1 mM DTT), and then the dialyzed fraction is again subjected to 105,000g centrifugation for 1 h. The whole supernatant is loaded onto a DEAEcellulose (Whatman DE-52, England) open column equilibrated with the same buffer. After being washed with the same buffer, proteins are eluted with a linear gradient of 0–0.5 M KCl in the same buffer, and fractions containing P450nor are collected, concentrated, and dialyzed overnight against buffer B (50 mM potassium phosphate containing 10% glycerol, 0.1 mM EDTA, and 0.1 mM DTT, pH 7.5). The sample is then mixed with the same volume of 4 M ammonium sulfate in buffer B and precipitates formed are removed by centrifugation at 10,000g for 10 min. The resulting supernatant is applied to a phenyl-Superose HR5/5 column (Pharmacia) equilibrated with 2 M ammonium sulfate in buffer B. The column is then eluted with a linear gradient of ammonium sulfate in buffer B from 2 to 0 M. Fractions containing P450nor are collected and dialyzed overnight against buffer A and loaded onto a Mono Q HR5/5 column (Pharmacia) equilibrated with the same buffer. Protein is eluted with a linear gradient of 0–0.5 M KCl in buffer A. Fractions containing purified P450nor are dialyzed against buffer A and stored at –80 until use. An aliquot is always taken at each step for Nor activity assay and SDS-PAGE. All purification manipulations should be conducted below 4 or on ice with the exception of phenyl-Superose and Mono Q column chromatographies, which are

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conducted at room temperature. Tnor is very abundant in yeast T. cutaneum; 2 mg Tnor is purified from only 63 g wet cells using the aforementioned method. The recovery of Tnor is half of the total soluble P450 proteins. The molecular mass is around 43 kDa, which is similar to Fnor and Cnor (42– 46 kDa) (Fig. 7.1). In addition, absorption spectra of the purified protein also indicate the characteristic of cytochrome P450 (Zhang et al., 2001). Absorption spectra of the purified protein in the resting state are in an oxidized form (ferric state) with a mixture of high-spin (390 nm) and low-spin (412 nm) states.

5. Nitric Oxide Reductase Activity Assay Nor activity can be assayed by measuring N2O production (Nakahara et al., 1993; Usuda et al., 1995) or NADH consumption (Kaya et al., 2004; Shiro et al., 1995). Typically, Tnor activity is assayed anaerobically in a 20-ml tube sealed with a double butyl rubber stopper. The reaction mixture

M

Sup DE52 (NH4)2SO4

P

Q10 Q11

kDa 94 67

Tnor 43

30

Figure 7.1 SDS-PAGE analysis of purified P450nor fromT. cutaneum. Lanes from left to right were, respectively, performed using 1, marker proteins containing carbonic anhydrase (30 kDa), ovalbumin (43 kDa), bovine serum albumin (67 kDa), and phosphorylase b (94 kDa); 2, soluble fraction (222 mg); 3, after DEAE52-cellulose column (180 mg); 4, after precipitation with ammonium sulfate (227 mg); 5, after phenyl-Superose column (53 mg); 6 and 7, fractions 10 and 11 after Mono Q column (15.6 and 6.2 mg, respectively). The Tnor band is cut out for trypsin treatment, HPLC, N-terminal amino acid sequencing.

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(5 ml) contains 100 mM potassium phosphate buffer (pH 7.2), 10% glycerol, 0.1 mM EDTA, 0.1 mM DTT, 5 mM NADH or NADPH, and 4 nM purified Tnor. After replacing the gas phase with helium, the reaction mixture is prewarmed at 30 for 2 min. The reaction is initiated by adding 1.0 ml NO gas with a syringe. The mixture is vortexed frequently, and every 0.5-ml gas phase sample of the reaction is collected by a syringe at each time point for GC analysis. Figure 7.2 shows typical P450nor activity from purified Tnor using NADH as the electron donor. Turnover rates against NO (nmol NO/min/nmol P450nor) of purified Tnor using NADH and NADPH as electron donors are 1.27 and 1.05 104, respectively, which are almost the same as that of purified recombinant Fnor (1.28 104), Cnor1 (1.05 104), and Cnor2 (1.13 104) using NADH as the electron donor at 30 .

6. Protein Sequencing Purified protein (60 mg) is applied to SDS-PAGE and blotted onto a nitrocellulose membrane. The blot is stained with Ponceau S (Sigma), and the protein band is excised and digested partially with trypsin (Aebersold et al., 1987). The digested peptides are separated by HPLC on a reversedphase C18 column, and their N-terminal amino acid sequences are determined using a Perkin Elmer Precise 492 automated protein sequencer. Sequences of peptides show a very high identity to other P450nors, for example, to the Cnor1 amino acid sequence (Fig. 7.3).

N2O (nmol)

1200

800

400

0 0

5 10 Reaction time (min)

15

Figure 7.2 Nitric oxide reductase activity of P450nor fromT. cutaneum. Incubation was conducted anaerobically in a total volume of 5 ml with 5 mM NADH and 4 nM purified Tnor, and the total amounts of N2O were determined by GC with Porapak Q. , with NADH; ○, without NADH.


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STRAAPKFPFARASGMEPPA (N-terminus) MASEPPSFPFQRASGMEPPAEFARLRATDPVSKVKLFDGSLAWLVTKYKDVTFVATDERLS AAAQAQPTFVDMDAPDHMK (P8) KVRTRPGFPELNAGGKQAAKAKPTFVDMDAPDHMNQRGMVESLFTLEHVKKLQPYIQKTVD DLLAAMKKKGCANGPVDLVKEFALPVPSYIIYTILGVPFNDLDHLTNQNAIRTNGSSTARE ASAANQELLDYLASLVEKRLEEPKDDLISKLCTEQVKPGNIEKADAVQIAFLLLVAGNATM VNMIALGVVTLFQHPEQLAQLKANPSLAPQFVEELCRYHTASALAIKRTAKVDLEIGGKHI TFDKDPLAFGQGPHR (P13) VELTAVF KANEGIIASNQSANRDADIFENPDEFNMNRKWPAEDPLGYGFGPHRCIAEHLAKAELTTVF ETLYK (P21) DVGIVELPVT (P20) ATLFKEFPDLNIAVPFEKINFTPLGGDVGVVDLPVTF

Figure 7.3 Comparison of amino acid sequences of Tnor tryptic peptides with C. tonkinense Cnor1.

7. Isolation of cDNA Total DNA is prepared from T. cutaneum by the standard method (also see Tomura et al., 1994). In brief, about 5–10 g of yeast wet cells is freeze dried and disrupted by grinding in 8 ml of TE buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0). Then 4 ml lysis buffer (50 mM Tris-HCl, pH 8.0, 100 mM EDTA, 100 mM NaCl, 1% SDS) is added and incubated at 65 for 30 min with gentle mixing. After adding 1/10th volume of 5 M (pH 5.2) potassium acetate, an equal volume of phenol-chloroform (1:1) is added and the mixture is incubated for another 30 min. After centrifugation at 11,000 rpm for 20–30 min, the aqueous phase is added with two-fold 100% ethanol and put in an –80 freezer for 30 min before centrifugation for another 30 min at 11,000 rpm. The pellet is washed with 70% ethanol, dissolved in 4 ml of TE with RNase, and incubated at room temperature overnight. DNA is extracted with phenol-chloroform another two to three times to completely get rid of proteins. The DNA is precipitated by ethanol, washed with 70% ethanol, and finally dissolved in 4 ml of TE buffer. Four mixed primers are designed according to the amino acid sequences of the purified P450nor and the trypsin digests (see Fig. 7.3, underlined): P1, 50 -CCNAARTTYCCNTTYGC-30 P2, 50 -TCNACDATNCCNACRTC-30 P3, 50 -GAYATGGAYGCNCCNGAYCAYATG-30 P4, 50 -GGRTCYTTRTCRAANGT-30 The first polymerase chain reaction (PCR) is conducted using 400 pmol of P1 and P2 primers each and 0.5 mg total DNA as the template. Amplification is performed by the process of denaturation at 94 for 1 min followed

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by 30 cycles of incubations at 94 for 20 s, 50 for 1 min, and 72 for 1.5 min, and extension at 72 for 10 min. Second-round PCR is performed under the same conditions using the product of the first PCR as the template and P3 and P4 as the primers. The nucleotide sequence of the second PCR product is determined by the dideoxy chain-termination method (Sanger et al., 1977) using the automated DNA sequencer LONG READIR Model 4200 (LI-COR). The entire cDNA is cloned by the rapid amplification of cDNA ends (RACE) using the SMART RACE cDNA amplification kit (Clontech). Poly(A)þ mRNA is prepared from T. cutaneum cells using an mRNA purification kit (Pharmacia) according to the supplier’s instructions. Primers are designed based on the nucleotide sequence of the PCR product given earlier. The nucleotide sequences of the primers are GSP1, 50 -GGGTCCTTGTCAAACGTGCGGCGAATGT-30 NGSP1, 50 -CGCTGTAGTTACTCGCGATGATGCCCTC-30 (for 50 RACE) GSP2, 50 -CACATGAAGCAGCGCGGCCTCGTCGAGG-30 NGSP2, 50 -TCCAGAGCGTCATCGACGAGGCGCTCGA-30 (for 30 RACE) The RACE products are ligated to pGEM-T vectors (Promega) and their nucleotide sequences are determined. The deduced amino acid sequence from the sequence of cDNA contains the same sequences as those of the N terminus of native Tnor and its tryptic fragments (see Fig. 7.3). It contains 416 amino acids with a molecular mass of 45.2 kDa. The sequence shows overall similarities to those of other isoforms, with identities of 65.1, 65.3, and 64.8% to Fnor, Cnor1, and Cnor2, respectively.

8. Subcellular Fractionation of T. cutaneum By comparing the amino acid sequences of P450nor isoforms, it is interesting to note that the sequences of the N-terminal residues are not conserved among these proteins. The N-terminal portion of Fnor and Cnor1 contains hydrophobic and positively charged amino acid residues, which acts as the targeting signal for transportation to mitochondria; this portion is cleaved before the mature protein is formed. The purified Tnor protein also lacks the first 18 amino acid residues and therefore is considered to be a mitochondrial protein as well. To confirm its localization within mitochondria, yeast cells cultured under the denitrifying conditions described earlier are disrupted below 4 by grinding with a 2.5-fold (w/w) excess of quartz sand in 0.8 M sucrose buffer (isotonic conditions), pH 7.2, containing 0.1 mM EDTA, 10 mM


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Tris-HCl, and the protease inhibitors described previously. The quartz sand and undisrupted cells are removed by centrifugation at 1500g for 30 min. The cell homogenate is then subjected to centrifugation at 10,000g for 60 min. The resulting supernatant is separated into soluble (cytoplasm) and microsomal fractions by further centrifugation at 105,000g for 90 min. The precipitate resulting from the centrifugation at 10,000g described earlier (large particles) is resuspended in sucrose buffer and subjected to centrifugation at 105,000g for 180 min on a discontinuous sucrose density gradient (1.0, 1.26, and 1.46 M ). The resulting interface fractions are collected, and their activities of nitric oxide reductase and the mitochondrial marker enzyme (cytochrome c oxidase, COX) are measured. Most of the NADH- and NADPH-dependent Nor activities can be recovered together with COX activity, indicating that Tnor is localized in the mitochondria (Zhang et al., 2001).

9. Site-Directed Mutagenesis Of the five cloned CYP55A genes, CYP55A1 (Fnor) from F. oxysporum has been the most studied. Recombinant proteins of Fnor have been made in order to study the structure and function of P450nor (Kudo et al., 2001; Obayashi et al., 2000; Okamoto et al., 1997; Shimizu et al., 2000; Su et al., 2004; Umemura et al., 2004; Zhang et al., 2002b) (Table 7.1, primers used for mutagenesis can be found from the listed references). A study on the crystal structure of the P450nor double mutant GG (Gly73/Gly75) with substrate confirmed the importance of these amino acids in the P450nor reaction (Oshima et al., 2004). The P450nor isozymes isolated from fungi can be divided into two types, depending on their specificity for electron donors, NADH and NADPH. Fnor and Cnor1 are specific for NADH, whereas Cnor2, Tnor, and Anor can utilize both NADH and NADPH. Cnor2, Tnor, and Anor contain multiple Ala and Gly residues in the region 73SA(P)GGKAAA80. Using site-directed mutagenesis to modify the structure of Fnor, studies indicate that Ser73 and Ser75 in the B’-helix (74ASGKQA79) determine the specificity for NADH and NADPH (Zhang et al., 2002b). Site-directed mutagenesis is achieved by PCR using template pfp(450)20, which has the Fnor cDNA cloned in the pUC18 vector (Kizawa et al., 1991). Primers M13–47 and M13-RV (Takara, Otsu, Japan) are specific for pUC18. The following primers are designed to study the function of B’helix and its neighboring region (73SASGKQAA80 in Fnor) (mutated sites are underlined): 3A (74ASGKQA79 ! AAA): GAGCTTAGCGCCGCTGCAGCCAA GGCAAA

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Table 7.1.

Functional amino acids in P450nor from F. oxysporum

Amino acid position

Thr

243

Ser286, Asp393 Arg64, Arg174 Ser73, Ser75 Asp88 Glu71

Function

References

Binding to nicotinic acid ring

Okamoto et al. (1997) Obayashi et al. (2000) Shimizu et al. (2000) Kudo et al. (2001)

Proton delivery Interact with pyrophospate moiety Access 2’-phosphate group of NADPH Release of NADþ Help Arg64 for NADH binding

Zhang et al. (2002b) Umemura et al. (2004) Su et al. (2004)

SG (Ser73/Gly75): CTTAGCGCCGGTGGAAAGCAAGCA GG (Gly73/Gly75): TTCCCTGAGCTTGGCGCCGGTGGAAAG The first PCR (30 cycles) using M13–47 and mutant forward primers or M13-RV and mutant reverse primers is performed as follows: 1 min for 94 , 2 min for 55 , and 2 min for 72 . The PCR products are purified from agarose gel and mixed in PCR buffer and H2O and are subjected to the following treatment: 10 min for 95 , 30 min with a gradient decrease of temperature from 95 to 37 , and 30 min for 37 . After the addition of Taq polymerase and dNTP, the mixture is kept for 10 min at 72 , after which M13–47 and M13-RV primers are added into the reaction mixture and PCR (30 cycles) is performed again as described earlier. The mutated DNA products are digested with restriction nucleases BssHII and BglII, and the 1.1-kb fragments are purified and inserted into the expression vector for P450nor [pT7nor, which comprises pRSET C (Invitrogen) with the T7 promoter in which the N-terminal His tag is replaced with the full-length P450nor cDNA] in lieu of the corresponding wild-type restriction fragments. The presence of the desired mutation is confirmed by DNA sequencing, and segments corresponding to the inserted restriction fragments are then sequenced completely to exclude the acquisition of unwanted mutations during amplification or cloning.

10. Expression of Recombinant Proteins Using the aforementioned strategy, the DNAs of Fnor, its derivatives, and Tnor in pT7nor vector are introduced individually into Escherichia coli JM109(DE3). The transformed cells are precultured overnight at 37 in


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20 ml of LA broth (1% tryptone, 0.5% yeast extract, 0.5% NaCl, 25 mg/ml ampicillin) supplemented with 0.5% glucose. Transformant precultures are transferred to 2 liters of LA broth in 5-liter Erlenmeyer flasks, shaken for 5–6 h, and further shaken at 120 rpm overnight at 30 in the presence of 1 mM isopropyl-1-thio-b-D-galactoside.

11. Purification of Recombinant Proteins The transformed cells are harvested, suspended in Tris buffer [10 mM Tris-HCl, pH 8.0, 10% (v/v) glycerol, 0.1 mM DTT, 0.1 mM EDTA, 0.1 mM PMSF, 0.1 mM TPCK] and disrupted twice using a French pressure cell press (SimAminco, France) at 20,000 psi. The suspension is centrifuged at 1500g for 30 min to remove cell debris and then centrifuged at 20,000g for 1 h. The resulting supernatant is dialyzed overnight against Tris buffer. A Whatman DEAE-cellulose column is equilibrated with Tris buffer, and then the sample is charged and eluted with a 0 to 0.4 M KCl gradient. Fractions containing P450nor are collected and dialyzed against Tris buffer. The P450nor fraction is purified further by chromatography through either Mono Q HR 5/5 or Resource Q columns (Pharmacia Biotech) in the same manner as described previously for DEAE. Using this method, the absorbance ratio of mutant proteins is always over 1.6 at 413 nm compared with that at 280 nm. Recombinant proteins of Fnor, its mutants, and Tnor have almost the same high-spin ratio between 54 and 59% in 50 mM N-Tris (hydroxymethyl) methyl-2-aminoethanesulfonic acid (TES) buffer (pH 7.2), suggesting that they have similar structures and the mutations do not affect overall structural conformation. The content of the high-spin form is used as an approximate value in comparing P450 with its mutants. The percentage of high-spin heme of P450nor at the resting state is estimated from the ratio of the absorbance of the high-spin form (390 nm) to that of the lowspin form (414 nm) as described previously (Imai and Komori, 1992; Imai et al., 1997). From data provided by Dr. Yoshio Imai through personal communication, we simply calculate spin ratios (%) using YH ¼ (101XH – 39.9)/(43.6 þ 56.7XH) and YL ¼ (83.5XL – 44.3)/(56.7 þ 43.6XL) in which YH and YL represent high-spin (%) and low-spin (%) ratios, respectively, while XH ¼ A390/A414 and XL ¼ A414/A390, respectively. For example, the purified Tnor at ferric state gives absorbance of 0.454 and 0.47 at 390 and 414 nm, respectively (Zhang et al., 2001); therefore, XH ¼ 0.966. From the formula just given, the high-spin ratio YH ¼ 58.6%. In order to investigate the possible role of B’-helix in the access of NAD(P)H, mutant 3A in which the 74ASGKQA79 is replaced by three alanines to destroy the B’-helix can be utilized. NADH-dependent

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Nor activity of mutant 3A decreases to the same level as the NADPHdependent activity of wild-type Fnor, indicating its crucial role in conferring full Nor activity (Zhang et al., 2002b). Mutations at Ser73 and Ser75 improve NADPH-dependent Nor activity significantly without affecting NADH-dependent activity (Fig. 7.4), indicating that these amino acids are crucial for determining the specificity of P450nor for NADH and NADPH.

12. Titration of NAD Analogs Although the spectral perturbation of P450nor-bound heme by some ligands can be observed (Okamoto et al., 1998), it is difficult to detect spectral perturbation upon mixing P450nor and NADH, possibly because of the powerful absorbance of NADH that overwhelms the spectral perturbation of bound heme. Because SG and GG mutant proteins can enhance the rate of reduction of NO with NADPH, it is possible to screen NAD(P)H analogs to detect spectral perturbation using such mutants. NAD(P)H and NAD(P) analogs (all from Sigma) are used to investigate their binding to P450nor. The screening of NAD(P) analogs b-NADþ, b-NADPþ, and 500 WT_NADH WT_NADPH SG_NADPH GG_NADPH

N2O (nmol)

400

300

200

100

0 0

1

2

3

Min

Figure 7.4 B’-helix of Fnor modulate the interaction with NADPH. Nor activity of WT (Ser73/Ser75) with NADPH as electron donor is only 13% of that with NADH as electron donor. SG (Ser73/Gly75) and GG (Gly73/Gly75) mutant proteins with NADPH as electron donor exhibit 63 and 79% activities of WT with NADH. NAD(P)H, 1 mM; P450nor,4 nM.

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70

70

60

60

50

50

40

1/ΔA

1/ΔA

3-pyridinealdehyde adenine dinucleotide (PAAD) are found to cause a detectable spectral change in WT and SG mutant proteins. Therefore, they can be analyzed further by spectrophotometric titration using a Beckman DU 7500 spectrophotometer. The dissociation constant (Kd) is calculated from the difference in absorbance (DA) at 413 nm from that at 395 nm. Recombinant P450nor (5 mM) in 50 mM TES buffer at pH 7.2 is mixed with an equal volume (100 ml) of each substrate analog (dissolved in the same buffer) and the spectrum of the mixture (200 ml volume) is recorded. 3-Pyridinealdehyde adenine dinucleotide and b-NADþ are found to give a type I spectral change (the increase in absorbance around 390 nm because of the high-spin state heme with a concomitant decrease in absorbance at 413 nm because of the low-spin heme) in Fnor, SG, GG, and Tnor (Oshima et al., 2004; Zhang et al., 2002b). It is interesting to note that while b-NADPþ induces a reverse type I spectral change in Fnor and its mutants, it causes a type I spectral change in Tnor, which can utilize both NADH and NADPH as electron donors. Thus, the reverse type I spectral change appears to be caused by binding in an unfavorable state. It is obvious that these NAD(P)H analogs can bind to P450nor in two alternative states within the bound form that cause either a type I or a reverse type I spectral change. It is also evident from the effects of these mutations in the B’-helix region that these analogs bind to P450nor from the distal side. The spectral perturbation caused by NADH analogs shows saturation against the ligand concentration, suggesting a specific binding of each ligand to Fnor and its mutants. The dissociation constant (Kd) of each Fnor-ligand complex is determined by spectrophotometric titration (Fig. 7.5). Maximum spectral changes of Fnor WT and SG proteins by PAAD are the same (0.0627 and 0.0628, respectively), whereas Kd values are 6.11 and 1.65 mM,

WT Kd = 6.11 mM

30

SG Kd = 1.65 mM

30 20

20 y = 15.947 + 97.385x R2 = 0.985

10 0

40

0.0

y = 15.913 + 26.239x R2 = 0.991

10 0

0.1

0.2

0.3 0.4 1/PAAD

0.5

0.6

0

1

2

3

1/PAAD

Figure 7.5 Double-reciprocal plot of PAAD concentration and the absorbance change at 395 and 413 nm of FnorWTand mutant SG.

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respectively, showing much higher affinity of NADH analogs to mutants, which led to the success in obtaining the crystal structure for the PAAD and GG complex (Oshima et al., 2004).

13. Stopped-Flow Rapid Scan Analysis The overall reaction can be divided into three partial reactions (Schemes 1–3) (Shiro et al., 1995): Fe3þ þ NO ! Fe3þ-NO (Scheme 1) Fe3þ-NO þ NAD(P)H þ Hþ ! I þ NAD(P)þ (Scheme 2) I þ NO ! Fe3þ þ N2O þ H2O (Scheme 3) Except for the last step (Scheme 3), these partial reactions can be observed as an isolated reaction. The chemical nature of the specific intermediate (I ) with a Soret absorption peak at 444 nm, which is formed upon reduction of the ferric-NO complex (Fe3þ-NO) with NAD(P)H (Scheme 2), is still under debate (Daiber et al., 2002, 2005; de Groot et al., 2005; Lehnert et al., 2006; Silaghi-Dumitrescu, 2003). Because the mutations at Ser73 and/or Ser75 in Fnor modulate the interaction with NADPH as well as overall Nor activity, the intermediate (I ) can be observed as an isolated reaction by rapid scan analysis as reported (Shiro et al., 1995). The P450nor reducing half-reaction is analyzed by following the appearance of the intermediate (I ) at 444 nm upon reduction of the Fe3þ-NO complex with NAD(P)H at 10 using a Unisoku rapid scan analyzer (Osaka, Japan) where 50 mM TES buffer (pH 7.2) is used, instead of phosphate buffer or Tris-HCl buffer, to eliminate the effects of anions on Nor activity (Kudo et al., 2001). The P450nor enzyme (final, 5 mM) in Fe3þ-NO is mixed with an equal volume of NADH or NADPH (final concentration, 20 mM ) anaerobically and the spectral changes are recorded. The gate time is set to 1 ms and the rate of I formation (kobs) is calculated. Under controlled conditions, accumulation of the intermediate (I ) is detectable upon reduction of Fe3þ-NO with NADH in WT Fnor (Scheme 2), whereas it is not available with NADPH (Zhang et al., 2002b). This is possibly because reduction (I formation) is slower than its decomposition. However, the accumulation of I upon reduction with NADPH as well as with NADH can be seen in both SG and GG mutants, showing that the rate of the reduction with NADPH was enhanced by the mutations. The apparent rate constant for the reduction at a fixed NADH (or NADPH) concentration (kobs) is obtained with each mutant from the time-dependent spectral change during the process of I formation (Zhang et al., 2002b), which clearly demonstrates that NADHor NADPH-dependent kobs for each enzyme species is parallel with the overall activity (see Fig. 7.4), indicating that the enhanced overall NADPH-dependent Nor activities in SG or GG arise from the reduction step.


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14. Other Analysis Protein concentration is determined by the standard method. Cytochrome c oxidase activity is measured by the method of Orii and Okunuki (1965). The P450nor concentration is determined spectrophotometrically from the CO difference spectrum as reported (Omura and Sato, 1964) using an extinction coefficient of 86.3 mM1cm1 for the difference in absorbance between 448 and 490 nm (Nakahara et al., 1993). NADH is determined from the absorbance at 340 nm using a molar absorption coefficient of 6.22 mM1cm1.

15. Conclusion The P450nor product is available commercially (Gekkeikan, Japan), with the goal of lowering NO levels within the human body. The importance of NO biology on human health has been well documented. Additionally, plants use NO as a weapon against invading pathogens. This chapter described some basic protocols on screening, purification, cloning, and functional analysis of cytochrome P450nor, providing some information and methods that may be helpful in further NO research.

ACKNOWLEDGMENTS Thanks are given to Professor Yoshio Imai for his kind discussion and suggestion of spin ratio calculations. We also thank R.W. Cameron Dingle and Georgia Zeigler, Department of Molecular and Biomedical Pharmacology, University of Kentucky, for reading and suggestions on the manuscript.

REFERENCES Aebersold, R. H., Leavitt, J., Saavedra, R. A., Hood, L. E., and Kent, S. B. H. (1987). Internal amino acid sequence analysis of proteins separated by one- or two-dimensional gel electrophoresis after in situ protease digestion on nitrocellulose. Proc. Natl. Acad. Sci. USA 84, 6970–6974. Daiber, A., Nauser, T., Takaya, N., Kudo, T., Weber, P., Hultschig, C., Shoun, H., and Ullrich, V. (2002). Isotope effects and intermediates in the reduction of NO by P450NOR. J. Inorg. Biochem. 88, 343–352. Daiber, A., Shoun, H., and Ullrich, V. (2005). Nitric oxide reductase (P450nor) from Fusarium oxysporum. J. Inorg. Biochem. 99, 185–193. de Groot, M. T., Merkx, M., Wonders, A. H., and Koper, M. T. (2005). Electrochemical reduction of NO by hemin adsorbed at pyrolitic graphite. J. Am. Chem. Soc. 127, 7579–7586.

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Imai, Y., and Komori, M. (1992). In ‘‘Shin Seikagaku Jikken Koza,’’ Vol. 5, pp. 205–221. Japanese Biochemical Society, Tokyo-kagakudojin, Tokyo. Imai, Y., Okamoto, N., Nakahara, K., and Shoun, H. (1997). Absorption spectral studies on heme ligand interactions of P-450nor. Biochem. Biophys. Acta 1337, 66–74. Kaya, M., Matsumura, K., Higashida, K., Hata, Y., Kawato, A., Abe, Y., Akita, O., Takaya, N., and Shoun, H. (2004). Cloning and enhanced expression of the cytochrome P450nor gene (nicA; CYP55A5) encoding nitric oxide reductase from Aspergillus oryzae. Biosci. Biotechnol. Biochem. 68, 2040–2049. Kizawa, H., Tomura, D., Oda, M., Fukamizu, A., Hoshino, T., Gotoh, O., Yasui, T., and Shoun, H. (1991). Nucleotide sequence of the unique nitrate/nitrite-inducible cytochrome P-450 cDNA from F. oxysporum. J. Biol. Chem. 266, 10632–10637. Kudo, T., Takaya, N., Park, S.-Y., Shiro, Y., and Shoun, H. (2001). A positively charged cluster formed in the heme-distal pocket of cytochrome P450nor is essential for interaction with NADH. J. Biol. Chem. 276, 5020–5026. Kudo, T., Tomura, D., Liu, D., Dai, X., and Shoun, H. (1996). Two isozymes of P450nor of Cylindrocarpon tonkinense: Molecular cloning of the cDNAs and genes, expressions in the yeast, and the putative NAD(P)H-binding site. Biochimie 78, 792–799. Lehnert, N., Praneeth, V. K. K., and Paulat, F. (2006). Electronic structure of iron(II)– porphyrin nitroxyl complexes: Molecular mechanism of fungal nitric oxide reductase (P450nor). J. Comput. Chem. 27, 1338–1351. Nakahara, K., and Shoun, H. (1996). N-terminal processing and amino acid sequence of two isoforms of nitric oxide reductase cytochrome P450nor from F. oxysporum. J. Biochem. (Tokyo) 120, 1082–1087. Nakahara, K., Tanimoto, T., Hatano, K., Usuda, K., and Shoun, H. (1993). Cytochrome P-450 55A1 (P-450dNIR) acts as nitric oxide reductase employing NADH as the direct electron donor. J. Biol. Chem. 268, 8350–8355. Obayashi, E., Shimizu, H., Park, S.-Y., Shoun, H., and Shiro, Y. (2000). Mutation effects of a conserved threonine (Thr243) of cytochrome P450nor on its structure and function. J. Inorg. Biochem. 82, 103–111. Okamoto, N., Imai, Y., Shoun, H., and Shiro, Y. (1998). Site-directed mutagenesis of the conserved threonine (Thr243) of the distal helix of fungal cytochrome P450nor. Biochemistry 37, 8839–8847. Okamoto, N., Tsuruta, K., Imai, Y., Tomura, D., and Shoun, H. (1997). Fungal P450nor: Expression in Escherichia coli and site-directed mutagenesis at the putative distal region. Arch. Biochem. Biophys. 337, 338–344. Omura, T., and Sato, R. (1964). The carbon monoxide-binding pigment of liver microsomes. II. Solubilization, purification and properties. J. Biol. Chem. 239, 2379–2385. Orii, Y., and Okunuki, K. (1965). Studies on cytochrome a. XV. Cytochrome oxidase activity of the Okunuki preparation and its activation by heat, alkali and detergent treatments. J. Biochem. (Tokyo) 58, 561–568. Oshima, R., Fushinobu, S., Su, F., Zhang, L., Takaya, N., and Shoun, H. (2004). Structural evidence for direct hydride transfer from NADH to cytochrome P450nor. J. Mol. Biol. 342, 207–217. Sanger, F., Nicklen, S., and Coulson, A. R. (1977). DNA sequencing with chainterminating inhibitors. Proc. Natl. Acad. Sci. USA 74, 5463–5467. Shimizu, H., Obayashi, E., Gomi, Y., Arakawa, H., Park, S.-Y., Nakamura, H., Adachi, S., Shoun, H., and Shiro, Y. (2000). Proton delivery in NO reduction by fungal nitric-oxide reductase: Cryogenic crystallography, spectroscopy, and kinetics of ferric-NO complexes of wild-type and mutant enzymes. J. Biol. Chem. 275, 4816–4826. Shiro, Y., Fujii, M., Iizuka, T., Adachi, S., Tsukamoto, K., Nakahara, K., and Shoun, H. (1995). Spectroscopic and kinetic studies on reaction of cytochrome P450nor with nitric


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oxide: Implication for its nitric oxide reduction mechanism. J. Biol. Chem. 270, 1617–1623. Shoun, H. (2006). Denitrification and anaerobic energy producing mechanisms by fungi. Tanpakushitsu Kakusan Koso. 51, 419–429. Shoun, H., Kim, D. H., Uchiyama, H., and Sugiyama, J. (1992). Denitrification by fungi. FEMS Microbiol. Lett. 73, 277–281. Shoun, H., and Tanimoto, T. (1991). Denitrification by the fungus Fusarium oxysporum and involvement of cytochrome P-450 in the respiratory nitrite reduction. J. Biol. Chem. 266, 11078–11082. Silaghi-Dumitrescu, R. (2003). Nitric oxide reduction by heme-thiolate enzymes (P450nor): A reevaluation of the mechanism. Eur. J. Inorg. Chem. 2003, 1048–1052. Su, F., Fushinobu, S., Takaya, N., and Shoun, H. (2004). Involvement of a Glu71-Arg64 couple in the access channel for NADH in cytochrome p450nor. Biosci. Biotechnol. Biochem. 68, 1156–1159. Takaya, N., and Shoun, H. (2000). Nitric oxide reduction, the last step in denitrification by F. oxysporum, is obligatorily mediated by cytochrome P450nor. Mol. Gen. Genet. 263, 342–348. Tomura, D., Obika, K., Fukamizu, A., and Shoun, H. (1994). Nitric oxide reductase cytochrome P-450 gene, CYP55, of the fungus Fusarium oxysporum containing a potential binding-site for FNR, the transcription factor involved in the regulation of anaerobic growth of E. coli. J. Biochem. (Tokyo) 116, 88–94. Tsuruta, S., Takaya, N., Zhang, L., Shoun, H., Kimura, K., Hamamoto, M., and Nakase, T. (1998). Denitrification by yeasts and occurrence of cytochrome P450nor in Trichosporon cutaneum. FEMS Microbiol. Lett. 168, 105–110. Umemura, M., Su, F., Takaya, N., Shiro, Y., and Shoun, H. (2004). D88A mutant of cytochrome P450nor provides kinetic evidence for direct complex formation with electron donor NADH. Eur. J. Biochem. 271, 2887–2894. Usuda, K., Toritsuka, N., Matsuo, Y., Kim, D.-H., and Shoun, H. (1995). Denitrification by the fungus Cylindrocarpon tonkinense: Anaerobic cell growth and two isozyme forms of cytochrome P-450nor. Appl. Environ. Microbiol. 61, 883–889. Zhang, L., Kudo, T., Takaya, N., and Shoun, H. (2002a). Distribution, structure and function of fungal nitric oxide reductase P450nor: Recent advances. Int. Congr. Ser. 1233, 197–202. Zhang, L., Kudo, T., Takaya, N., and Shoun, H. (2002b). The B’ helix determines cytochrome P450nor specificity for the electron donors NADH and NADPH. J. Biol. Chem. 277, 33842–33847. Zhang, L., Takaya, N., Kitazume, T., Kondo, T., and Shoun, H. (2001). Purification and cDNA cloning of nitric oxide reductase cytochrome P450nor (CYP55A4) from Trichosporon cutaneum. Eur. J. Biochem. 268, 3198–3204.


C H A P T E R

E I G H T

A Quantitative Approach to Nitric Oxide Inhibition of Terminal Oxidases of the Respiratory Chain Maria G. Mason,* Rebecca S. Holladay,* Peter Nicholls,* Mark Shepherd,† and Chris E. Cooper* Contents 1. Introduction 2. Evaluation of Current Techniques for Measuring pNO, pO2, and KM (O2) 3. Nitric Oxide Donor Compounds 4. Nitric Oxide Kinetics 4.1. Measurement of IC50 (NO) 4.2. Factors that may influence IC50 values 4.3. Measurement of KD (NO) 4.4. Pitfalls of nitric oxide kinetic analysis 4.5. Comparison of dynamic and steady state IC50 NO measurements 4.6. Measurement of NOoff rates and estimation of NOon rates 5. Oxygen Kinetics 6. Optical Detection of Enzyme Intermediates in the Presence of Oxygen and NO 6.1. Spectro-electrode system 6.2. Identification of nitric oxide-bound intermediates Appendices Appendix A. Correction for electrode response time Appendix B. Instrumentation for measuring nitric oxide and oxygen kinetics Appendix C. Equipment suppliers Acknowledgments References

* {

136 137 138 139 139 141 141 144 146 146 149 151 152 153 153 153 153 155 156 156

Department of Biological Sciences, University of Essex, Colchester, United Kingdom University of Sheffield, Western Bank, Sheffield, United Kingdom


#


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136

Maria G. Mason et al.

Abstract Inhibition of terminal respiratory oxidases by nitric oxide (NO) plays important physiological roles in signaling and host defense. Using a bacterial quinol oxidase and mitochondrial cytochrome c oxidase, this chapter describes simple polarographic methods to quantify the kinetic characteristics of inhibition by NO. This chapter points out the inherent pitfalls of both experimental design and data analysis and compares alternative methods. Additionally, it describes a system designed to acquire polarographic and spectral data simultaneously to permit identification of spectral intermediates under defined conditions.

1. Introduction Nitric oxide (NO) inhibition of bacterial and mitochondrial terminal respiratory chain oxidases is increasingly being seen as playing a key physiological role in host defense (Fang, 1997; Pieters and Ploegh, 2003) and signaling (Carreras and Poderoso, 2007). NO can reversibly bind, irreversibly damage, or be metabolized by oxidases (Cooper, 2002). In mitochondria, nitric oxide inhibition can perturb electron transfer chain redox states at sites distinct from the oxidase without directly compromising energy metabolism (Quintero et al., 2006); thus it can play a role in cell signaling, possibly via modulating the production of reactive oxygen species (Palacios-Callender et al., 2004; Sarkela et al., 2001). Quantitative measurements are crucial to the interpretation of the importance of these interactions. At sufficiently high levels there are few biomolecules that do not react with NO. Therefore, understanding at what level NO may interact with an oxidase and measuring NO levels in vivo are both necessary. In addition to the importance for cell physiology (Cooper and Giulivi, 2007; Giulivi et al., 2006), analyzing the mechanism of inhibition can also inform an understanding of the chemistry of the interactions (Mason et al., 2006). However, quantitative measurements in this area are difficult. NO is an unstable compound that can react at multiple sites in cells (Cooper, 1999); indeed even individual oxidases can have multiple, and distinct, interactions with NO (Cooper, 2002). The purpose of this chapter is to describe methods for the measurement of the kinetic constants associated with nitric oxide inhibition of oxygen consumption by respiratory chain oxidases. It focuses on numbers derived primarily from steady state enzyme kinetics. It largely ignores the wealth of spectroscopic data describing NO/oxidase interactions, although it illustrates a system for measurement of the nature of optically detectable intermediates at defined levels of nitric oxide and oxygen. Because it is impossible to analyze nitric oxide inhibition of oxygen consumption without a parallel discussion of the methods used to measure

Nitric Oxide Inhibition of Oxidases

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the oxygen kinetics of respiration, optimum methods are described for measuring low levels of both substrate (O2) and inhibitor (NO). We use examples from the interaction of NO with a bacterial quinol oxidase from Escherichia coli and mitochondrial cytochrome c oxidase. However, the analysis methods are appropriate for any enzyme that consumes oxygen and is inhibited reversibly by nitric oxide, particularly if the ligands share the same site of interaction. In the case of quinol and cytochrome c oxidase, the oxygen reactive site is a high-spin ferrous heme center.

2. Evaluation of Current Techniques for Measuring pNO, pO2, and KM (O2) For a real time non-invasive measure of steady state nitric oxide partial pressure and/or concentration, polarography is currently the only viable technique. A number of sensors have been developed, including optical (Barker et al., 1999a,b), electrochemical (Wang et al., 2005), and dualmode amperometric/voltammetric (Malinski and Taha, 1992). However, these sensors are not yet available commercially. Optical (Larfars and Gyllenhammar, 1995) and paramagnetic ( James and Swartz, 2002) measurements can integrate NO accumulation over time, but generally do so by removing nitric oxide, thus perturbing the system in the process. However, for steady state oxygen measurements, a range of techniques are available (Swartz et al., 1997). Some of these directly measure pO2, e.g., most Clark-type polarographic electrodes and electron paramagnetic resonance (EPR) oximetry based on solid particles; others measure parameters that respond to the product of the diffusion rate and the concentration of oxygen, e.g., probes of absorbance (D’Mello et al., 1995), phosphorescence (Lo et al., 1996; Wilson et al., 1988), fluorescence ( Ji et al., 2002), and EPR probes based on soluble materials (Swartz et al., 1997). The distinction between pO2 and [O2] is important for in vivo experiments where the oxygen solubility and diffusion rate might be unknown, but is less critical for the in vitro measures of the type described here, where the measurements can be readily interconverted. The systems we study are always likely to produce biologically active molecules, e.g., nitrogen oxides, superoxide, and peroxide. Any measurement requires a range of controls to ensure that the probe is not being perturbed or causing a perturbation. Therefore, in general, we prefer using nonbiologically active systems such as oxygen electrodes or the current range of artificial phosphorescent and fluorescent probes rather than heme proteins such as myoglobin or hemoglobin. Most terminal oxidases are membrane proteins. Therefore, oxygen reacts with these oxidases in an active site separated from the bulk phase by the protein interior and a biological (cell membrane) or artificial

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(detergent micelle) membrane phase. As oxygen is measured in the external medium, it could be argued that oxygen diffusion and solubility in the membrane phase distort the oxygen kinetics of terminal membrane-bound oxidases. However, measurements of the interactions of hydrophobic gases (CO, O2, or NO) with oxidases seem, fortunately, insensitive to the nature of the aqueous/hydrophobic interface (Meunier and Rich, 1998). Extreme caution is required when measuring oxygen in the presence of NO, as it can perturb the measurement in two ways: either NO can act directly on the probe or the probe can affect the NO concentration. In the case of hemoglobin or myoglobin as an oxygen probe, NO can both alter the oxygen p50 ( Jia et al., 1996; Yonetani et al., 1998) and be scavenged by the oxygenated heme protein itself (Larfars and Gyllenhammar, 1995). Scavenging is likely to be less of an issue for nonbiological phosphorescent and fluorescent probes; although the probe signal depends on physical quenching by oxygen, it is unlikely that the intrinsic signal will be unduly affected by the presence of NO, as its concentration is likely to be at least 100 times less than that of oxygen in the types of experiments described here. However, potential artifacts should always be assessed by the use of suitable controls. Under certain conditions, similar problems can arise with polarographic electrodes, as cross talk can occur between dissolved oxygen and the nitric oxide probe, and vice versa; however, these conditions are unlikely to occur during steady state measurements of NO inhibition. Fluorescent or phosphorescent oxygen probes rely on oxygen perturbing a strong intrinsic signal. Therefore, they are most sensitive at low oxygen concentrations and run the risk of saturation at high pO2. In contrast, the sensitivity of polarographic electrodes is weakest at low oxygen tensions as the probe directly measures pO2; therefore, specialized, expensive highresolution oxygen electrode systems are needed for the low micromolar range (Gnaiger et al., 1995). The optimal choice of pO2 probe may therefore in some situations be dependent on the range of oxygen concentrations being studied.

3. Nitric Oxide Donor Compounds We commonly use nitric oxide donor compounds as an alternative to solutions saturated with NO gas. The principal advantages of using NO donors are as follow: stock solutions are relatively stable at high pH with sensible handling (i.e., use at 5 and store at –20 under N2); the concentration and any degradation of the donor compound can be checked readily by UV spectroscopy; freshly made stock solutions do not contain nitrate/ nitrite as a contaminant (which is difficult to remove completely from solutions of NO gas); and NO donor solutions can be prepared at much

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higher concentrations than those of the gas. A suitable donor for nitric oxide kinetics is the fast-releasing NO donor Proli NONOate (t1/2 ¼ 1.8 s at 37 in 0.1 M KPi at pH 7.4) (from Alexis Biochemicals, http://www.axxora.com).

4. Nitric Oxide Kinetics 4.1. Measurement of IC50 (NO) The most common method for determining the IC50 for NO is to measure the recovering oxygen respiration rate simultaneously with declining nitric oxide concentration, after inhibition by NO (Borutaite and Brown, 1996; Brown and Cooper, 1994; Griffiths and Garthwaite, 2001). An example of this type of experiment is shown in Fig. 8.1. Respiration by the oxidase is initiated by the addition of substrate, and the oxygen concentration begins to decrease. Once respiration reaches a steady rate, an aliquot of nitric oxide (or nitric oxide donor) solution is added to the chamber and respiration becomes inhibited. Substrate

3.0

250

B

NO donor

2.5

0.8

200

150 1.5 100 1.0 50

0.5

[O2] mM

[NO] mM

2.0

Respiration rate (mM O2 s−1)

A

0.6

0.4

0.2

0.0

0.0 0

100

200 300 Time (s)

400

0 500

0

0.5

1

[NO] mM

Figure 8.1 Measurement of IC50 (NO). (A) Time-dependent traces of oxygen (black line) and nitric oxide (grey triangles) concentration. Oxygen decreases as E. coli membranes respire on NADH until nitric oxide is added at approximately 100 s and respiration is inhibited. Respiration recovers as nitric oxide decays. Data shown within the boxed area are transformed into nitric oxide concentration-dependent respiration rates; (B) data points (þ) are fitted to the Hill equation ( ) to obtain values for IC50 and h (also see text). Membranes from E. coli strain RKP4544 (Stevanin et al., 2000) were grown (Gibson et al., 1977) and prepared (Poole and Haddock, 1974) as described previously. Experiments were performed using combined NO/O2 polarography (see Appendix B) in a light-impenetrable chamber at 35 in 0.1 M KPi, pH 7.4.

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Nitric oxide does not persist in aerobic solutions as it reacts with oxygen (Ford et al., 1993) (Eq. (8.1)) and other materials such as membrane lipids (O’Donnell et al., 1997) and reduced cytochrome c (Sharpe and Cooper, 1997) used commonly in experiments of this type.

4NO þ O2 þ 2H2 O ! 4NO2- þ 4Hþ

ð8:1Þ

Inhibition of cytochrome c oxidase by NO is generally reversible (Brown and Cooper, 1994) and continues only while NO remains in solution. Data obtained during the recovery of respiration (boxed area in Fig. 8.1) are transformed to give respiration rates as a function of [NO], shown plotted in Fig. 8.1B, together with the fit to an adapted form of the Hill equation (Eq. (8.2)). This fitting procedure gives values for IC50 and h. Traditionally, the Hill equation is used to determine the number of molecules bound; however, in this type of analysis, the value for h should not be interpreted in this way, as values greater than 1 are obtained from this procedure for cytochrome c oxidase, which requires only one molecule of NO to inhibit, and the value of h relates to the turnover number and the degree of buffering of inhibition (Mason et al., 2006). During partial inhibition the reductive pressure on the uninhibited population increases if the turnover is less than maximal (Chance et al., 1970) and consequently inhibition is buffered more at low turnover and h has a greater value.

v ¼V

V ½ih Kh0:5 þ ½ih

ð8:2Þ

where v is velocity, V is limiting velocity, [i] is inhibitor concentration, K0.5 is the inhibitor concentration ([i]) giving 50% V, and h is the Hill coefficient. Certain substrates, especially of a nonphysiological type, react directly with oxygen. This nonenzymatic autooxidation is evident when an enzyme cannot be completely inhibited. Equation (8.2) can be modified to account for this inherent autooxidation (see Eq. (8.3)). The oxygen concentrationdependent autooxidation rate is determined either by adding increasing amounts of NO until further decreases in respiration are not observed, (the residual rate is then the autooxidation rate) or by performing separate control experiments, in the absence of oxidase, at a number of oxygen concentrations (autooxidation is negligible below 40 mM O2 with ascorbate/ N,N,N 0 ,N 0 -tetramethyl-p-phenylenediamine).

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v ¼ Va

V ½ih Kh0:5 þ ½ih

ð8:3Þ

where Va is the sum of the autooxidation rate and the limiting velocity.

4.2. Factors that may influence IC50 values An important consideration when undertaking these types of experiments is to ensure that all factors are carefully controlled, such as temperature, incident light, and substrate concentration. This is because the IC50 values obtained depend on all these factors. Temperature is important because NO binding and, more importantly, dissociation rates are temperature-dependent (Sawicki and Gibson, 1977). Additionally, experiments are performed in a lightimpenetrable chamber because the bonds between nitric oxide and ferrous heme a3 and, to a lesser extent, cupric CuB are photosensitive (Borutaite et al., 2000; Sarti et al., 2000; Wever et al., 1985). Finally, if the experimental results are to be interpreted in a meaningful way, the enzyme turnover number must be known. This is for two reasons: (i) the predominance of the different pathways (competitive and noncompetitive with oxygen) of inhibition is determined by the electron flux and therefore turnover number of the enzyme and (ii) the competitive interaction is turnover number-dependent because the apparent affinity for oxygen (KM0 O2) depends on the turnover number. Consequently, the apparent IC50 for NO is also dependent on the turnover number (Antunes et al., 2004; Mason et al., 2006). To illustrate the effect that experimental conditions can have, literature values of IC50 (NO) for cytochrome c oxidase are shown in Fig. 8.2, plotted as a function of oxygen concentration. The data are so scattered is because values were obtained under different experimental conditions of enzyme turnover, temperature, and light.

4.3. Measurement of KD (NO) An example of experimental results obtained where the temperature and enzyme turnover were carefully controlled is shown in Fig. 8.3. Values for slope and intercept were obtained by regression analysis. Because of the relatively small error on the value of the slope (i.e., IC50 dependence on oxygen concentration), we can be confident that it is quite accurate. However, the large error on the value of the intercept, KD (NO), means that it is rather inaccurate and therefore not a reliable way of determining the KD. A more accurate way to determine the KD for a competitive

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1500

Apparent IC50 (NO) nM

1200

900

600

300

0 0

50

100 [O2] mM

150

200

Figure 8.2 Literature values of IC50 (NO) for cytochrome c oxidase. Oxygen concentration-dependent IC50 values for NO are shown from several literature sources (Bellamy et al., 2002; Borutaite and Brown,1996; Brookes et al., 2003; Brown and Cooper, 1994; Hollis et al., 2003; Koivisto et al., 1997; Lizasoain et al., 1996; Mason et al., 2006; Palacios-Callender et al., 2004; Torres et al., 1995). The distribution of values is scattered because it depends not only on oxygen concentration (Brown and Cooper, 1994), but also on other factors, such as enzyme turnover number (Antunes et al., 2004; Mason et al., 2006) and light (Borutaite et al., 2000; Sarti et al., 2000).Values are from experiments using purified enzyme, mitochondria, and whole cells.

inhibitor is from the relationship described in Eq. (8.4), between the apparent IC50 (NO), the substrate (oxygen) concentration, and the KM for oxygen (see later); it is essential that the KM (O2) is measured under the same experimental conditions as those used to obtain IC50 values.

IC50 0 ¼ KD þ KD ½S=KM

ð8:4Þ

(Cornish-Bowden, 2004). Once the KM (O2) has been determined, mean IC50 values can be plotted as a function of O2/KM (O2). Both the slope and the intercept are now dependent on KD, and the analysis is more robust. Figure 8.4 shows mean data from Fig. 8.3 replotted and fitted to Eq. (8.4). Although the error on the intercept is still large, the error on the slope is small and the value of KD is determined with greater precision. It is worth emphasizing that this type of analysis is for a purely competitive inhibitor and therefore if the interaction is not purely competitive the

143


450 400

apparent IC50 (nM NO)

350 300 250 200 150 100 50 0 0

20

40

60

80 [O2] mM

100

120

140

160

Figure 8.3 Oxygen concentration-dependent IC50 (NO) values in a controlled oxidase system. IC50 (NO) values for E. coli membranes, as a function of oxygen concentration, were obtained as described in Fig. 8.1 and using Eq. (8.1). Turnover number (244 e s16 SE) and temperature (35) were controlled carefully, and the reaction was performed in a light-impenetrable chamber. The solid line shows the fit from regression analysis, which gave values of 2.0 ( 0.24 SE) for the slope and 2.6 ( 25 SE) for the intercept. For other details, see legend to Fig. 8.1.

apparent IC50 (nM NO)

400

300

200

100

0 0

200

400 [O2] / KM(O2)

600

800

Figure 8.4 Measurement of KD (NO). IC50 (NO) values from Fig. 8.3 were binned into three different oxygen tension groups and the mean values of IC50 are plotted as a function of [O2]/KM (O2). Data were fitted to Eq. (8.4) before averaging.The solid line shows the fit from regression analysis, which gave values of 0.54 ( 0.06 SE) for the slope and 2.6 ( 25 SE) for the intercept. For other details, see legend to Fig. 8.3.

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fit may be poor. Mixed inhibition by nitric oxide has been demonstrated in mitochondrial cytochrome c oxidase (Mason et al., 2006); in addition to binding at the ferrous heme a3 of the binuclear center, in competition with oxygen (Brown and Cooper, 1994), there is also a noncompetitive inhibitory interaction that predominates when electron flux, and therefore enzyme turnover, is low and a high proportion of the enzyme is in the oxidized form (Mason et al., 2006; Sarti et al., 2000). Although mixed inhibition may also occur in other heme-copper oxidases, it has not yet been verified.

4.4. Pitfalls of nitric oxide kinetic analysis A number of pitfalls are associated with this type of analysis; an important consideration is the response time of the electrodes. Although oxygen electrode response times are relatively fast (t1–3 s) and are not usually corrected for in this type of experiment, this is not the case with nitric oxide electrodes. There are a number of commercially available NO electrodes with different characteristics, but generally the faster responding electrodes have greater noise and the slower responding electrodes have less noise. Figure 8.5 shows representative traces from similar inhibition experiments using NO electrodes with different response times; the NO trace in the lower panel is from a fast-responding electrode (t 4 s) with maximal baseline noise equivalent to 50 nM NO, whereas that in the upper panel is from a slow-responding NO electrode with no discernible noise (i.e., 1 nM NO resolution). In both traces the respiration rate returns to its maximal value just before 400 s (boxed areas). In the lower trace the recovery to maximal respiration corresponds to disappearance of NO, as expected, whereas in the upper trace (slow electrode response time) at the point of recovery to maximal respiration, the NO electrode records residual NO of approximately 800 nM. This is clearly an artifact of the response time of the electrode. The current challenge for nitric oxide electrode design and manufacture is for a fast-responding nitric oxide electrode with subnanomolar resolution, which is not perturbed by commonly used reductants such as ascorbate. Slow electrode response times can be corrected for (see Appendix A). However, the benefit of using a noise-free electrode is then lost by the noise introduced from the differential used in the correction process. Additionally, it is important to confirm that the response time of the electrode is the same upon a sudden increase in NO concentration as it is for a sudden decrease in NO concentration, as slow-response time electrodes do not always have this characteristic. The response time should also be NO concentration-independent.

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NO

1000

0.6

Substrate

[NO] nM

0.2 0

0

100

200

300

400

500

600

0.0 700 0.6

1000

0.4 500

Respiration rate ( mM O2 s−1)

0.4 500

0.2 0

0

100

200

300 400 Time (s)

500

600

0.0 700

Figure 8.5 Nitric oxide electrode response time is critical for accurate IC50 measurements. Experimental traces showing respiration rates (grey lines) of mitochondrial cytochrome c oxidase together with nitric oxide concentrations (black lines).The upper and lower panels compare results obtained using different NO electrodes with slow (t 14 s) and fast (t 4 s) response times.Traces obtained from the electrode with a fast response time show simultaneous disappearance of NO and recovery of full respiration rate (boxed region, lower panel); conversely, traces obtained from the electrode with a slow response time show that relief of inhibition (boxed region, upper panel) is not simultaneous with disappearance of NO.This apparent residual NO is an artifact of the electrode response time. Bovine cytochrome c oxidase, purified as described previously (Mason et al., 2006; Yonetani, 1960), was respiring on ascorbate plus cytochrome c (TN 10 e s1 of aa31) in 0.1 M KPi/0.1% lauryl maltoside, pH 7.4, at 35 in a light-impenetrable chamber using combined NO/O2 polarography (see Appendix B).

The only way to check if the electrode response time needs correcting or not is to correct and compare; if the portion of the trace required for analysis is different in the corrected and uncorrected traces, then correction is required; if they overlay, correction is not required. Although not recommended for this type of dynamic experiment, lownoise, slow-responding electrodes are useful for measurement of stable steady state NO concentrations. The creation of true NO steady states, and measurement of the respective enzyme inhibition, has only been attempted by the group of Garthwaite (Bellamy et al., 2002; Griffiths and Garthwaite, 2001; Griffiths et al., 2003). It has the advantage that it creates a situation analogous to ‘‘normal’’ enzyme kinetic measurements (i.e., with a stable inhibitor concentration). It is not a trivial process, requiring a balance of NO production by a

146


suitable slow-release NO donor and NO removal via oxygen or an added scavenger. The oxygen concentration-dependence of NO removal, both directly (Lewis and Deen, 1994) and via cells and cell membranes (Shiva et al., 2001), further complicates the measurement. In contrast to the dynamic measurement, each experimental run creates only a single data point; multiple measurements are required to generate an IC50 value. However, a priori one would expect this method to generate the ‘‘true’’ IC50.

4.5. Comparison of dynamic and steady state IC50 NO measurements The dynamic method will only approximate the steady state method if the enzyme samples multiple steady states during the relief of inhibition. Given the rather slow NO dissociation rate from some terminal oxidases, this is not necessarily true. To our knowledge there has been no direct test of dynamic and steady state methods. The inhibition constants derived from both methods are broadly similar, but as already pointed out, a meaningful comparison would have to be done under identical conditions because the NO inhibition constants depend on so many variables. We have developed a simple dynamic model of NO inhibition of an oxidase, competitive with oxygen (Cooper and Giulivi, 2007). We then calculated the true steady state NO inhibition constants analytically and compared them to those generated from the model, following a bolus addition of nitric oxide. Figure 8.6 shows that provided the NOoff rate is at least 0.1 s1, dynamic and steady state methods produce identical results. However, at slower NOoff rates the dynamic method may measure artificially low IC50 values, especially at higher oxygen concentrations.

4.6. Measurement of NOoff rates and estimation of NOon rates Ligand dissociation rates can be measured as the return of respiratory activity after addition of a reagent that scavenges the inhibitory ligand. For accuracy, this type of experiment is best performed using a high-resolution respirometer. Traces from an experiment of this type, using the Oxygraph 2K instrument, are shown in Fig. 8.7. Respiration of E. coli membranes is initiated by the addition of NADH and then an aliquot of the fast-release NO donor Proli NONOate is added to inhibit respiration. Immediately after addition of NO donor, an increase in the respiration rate is observed; this is because of the reaction of nitric oxide with oxygen (Eq. (8.1)). Once respiration is fully inhibited, an excess of the NO scavenger 2-(4-carboxyphenyl)-4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide (cPTIO) is added, causing a rapid and complete depletion of nitric oxide and simultaneous recovery of the

147


Oxygen consumption

300

Competitive inhibition

k3 [O2]

250

k1 E

k4 [NO]

k2 ES

ER

ERNO k−4

Oxygen-dependent NO removal

IC50 (NO) nM

200 NO + O2

k5 [O2] [NO]

NO−2

NO off rate 0.1 s−1

150

NO off rate 0.02 s−1

100

50

0 0

50

100 [O2] mM

150

200

Figure 8.6 Use of a dynamic model to compare different methods of measuring NO IC50. A dynamic model of nitric oxide inhibition of mitochondrial respiration was used (inset) to add a bolus of NO (0.5^2 mM) at different oxygen concentrations to the enzyme in turnover. Modeled time traces and derived plots of the type shown in Fig. 8.1 were then created.These were used to create a graph of IC50 versus [O2] for two different NOoff rates (note that the NOon rate was also changed so that the KD remained unaltered). Modeled data (points) were then compared to analytically derived values (straight line) created via solving the appropriate steady state rate equation (via King^ Altman methods). Values used in the model were k1 ¼ 103 s1; k2 ¼ 325 s1; k3 ¼ 400 mM1 s1; k4 ¼ 100 mM1 s1; k-4 ¼ 0.02 s1; k5 ¼ 0.0001. The enzyme concentration was 3 nM. For the faster NOoff rate: k4 ¼500 mM1 s1; k-4 ¼ 0.1 s1. Note that the mechanism and values for removal of NO via oxygen were chosen solely to create traces that approximated original data. In reality, NO removal occurs via multiple processes, including a direct reaction that is second order in [NO](see Eq. (8.1)).

respiration rate. Data acquired during the recovery of respiration are then corrected for electrode response time, as described in Appendix A. The corrected trace, together with an exponential fit, is shown in Fig. 8.7B. The mean functional NO dissociation rate of 0.163 s1 obtained by this method is in close agreement with that of 0.133 s1 reported by Borisov et al. (2007) for the same oxidase, and the relatively small difference can be accounted for by the difference in experimental temperature.

148


A 9

B

300

NO donor Substrate

0.20 200 [O2] mM

[NO] mM

100

3

0

0

500 Time (s)

1000

Respiration rate (mM O2 s−1)

NO scavenger

6

0.15

0.10

0.05

0.00

0

0 20 40 Time after NO removal (s)

Figure 8.7 Measurement of NO dissociation rate. (A) Uncorrected traces of oxygen (black line) and nitric oxide (grey triangles) concentrations during an experiment in which NO is scavenged so that the functional NO dissociation rate can be measured. Respiration is initiated by the addition of substrate (NADH), and then an aliquot of the fast NO-releasing donor Proli NONOate is added to fully inhibit respiration; finally the NO scavenger cPTIO is added and inhibition is relieved.The rate of recovery of respiration after scavenging NO is shown (B), after correction for electrode response time and changes in oxygen background, together with exponential fit for the functional NO dissociation rate.The NO dissociation rate (0.163 s1) for this enzyme is approaching the limit of resolution for this technique and therefore a number of replicates are necessary. Experiments were performed using combined NO/high resolution O2 polarography (see Appendix B). For other details, see legend to Fig. 8.1.

In these experiments we used cPTIO as a nitric oxide scavenger. However, cPTIO needs to be used with caution as it can react directly with some reductants, such as ascorbate. In these circumstances, oxyhemoglobin can be used as an alternative NO scavenger, although it should be used above its p50 (O2) to avoid release of oxygen that interferes with the oxygen respiration trace, making it difficult to measure the recovery rate. Oxymyoglobin is not a suitable alternative as it has a much lower p50 than oxyhemoglobin ( Wittenberg, 1974). Once the NO dissociation rate and KD have been measured, the NO association rate can be estimated from the relationship.

kon ¼

koff KD

ð8:5Þ


149

5. Oxygen Kinetics In order to analyze optimally the nitric oxide inhibition kinetics, it is necessary to have a measurement of the oxygen KM under identical conditions to those of the NO inhibition kinetics. As this measurement is made in the absence of NO, the earlier critique of O2 measurement systems that are NO sensitive is not relevant. As a result, we have the full panoply of systems to choose from. A full analysis and critique of the literature surrounding the measurement of the oxygen KM of terminal oxidases, both isolated and in cells, are beyond the scope of this chapter. Gnaiger and colleagues provided a useful comparison of the different methods available in 1995; the methods available, and their associated pitfalls, have not changed noticeably since then. Perhaps the key difference is between open and closed systems (i.e., whether a gas phase is present in the study). Removal of all effects of the gas phase requires careful system design to reduce back diffusion and oxygen stores; even the choice of an oxygen-rich Teflon-coated stirrer bar can make a big difference to measurements at low oxygen. Although these effects are not present in open systems (Brookes et al., 2003; Cole et al., 1982; Petersen et al., 1976), oxygen transfer between gaseous and aqueous phases can give rise to problems with unstirred boundary layers and oxygen gradients between the sensor and the sample. For this reason we favor a sealed system with no gas phase. In this case, similar values are obtained whether using high-resolution polarographic or optical techniques (Gnaiger et al., 1995). There is a major caveat in using polarography, however. It should be noted that a standard Clark-type electrode is unable to measure an oxygen KM in the low micromolar range (Schindler, 1967). This is because of a low intrinsic sensitivity at low [O2], resulting in few data points in the critical region. To obtain more data points, the oxygen consumption rate must be decreased into a regime where oxygen back diffusion becomes a major artifact. We have seen many supposedly sealed oxygen electrode systems where even an enzyme with a very low KM cannot decrease the [O2] to 200-fold inhibition), but carbon monoxide (CO) and nitric oxide (NO) do not (less than threefold inhibition) (Dunham et al., 2003; Gilles-Gonzalez et al., 1994; Sousa et al., 2007a). The unliganded form of ferric BjFixL is fully active, and cyanide inhibits this species analogously to the O2 inhibition of the ferrous form (Gilles-Gonzalez et al., 2006). The electronic resemblance between the cyanomet species (FeIIICN-) and the substantively polarized O2-bound state (FeIIO2FeIIIO2-), together with crystal structures of liganded forms of the heme-binding domain, suggest that regulatory switching of the kinase is triggered by changed polar interactions of the heme with residues of the heme pocket (Gong et al., 1998; Hao et al., 2002; Olson and Phillips, 1997). As a rule, microbial O2 sensors serve to initiate substantive lifestyle changes. For example, hypoxia is a key determinant of the rhizobial switch from a vegetative to a nonreplicative symbiotic state (David et al., 1988; Ditta et al., 1987; Sciotti et al., 2003; Soupene et al., 1995; Virts et al., 1988). In low O2, rhizobial FixL–FixJ systems induce a cascade of gene (nif, fix) expression that produces the nitrogen-fixation enzymes and their accessory proteins,

Heme-Ligand Sensing Histidine-Protein Kinases

175

the key regulators of denitrification, and one or more high-affinity alternative terminal oxidases for respiration in low O2. The latter function, which enables respiration during hypoxia, is also reported for homologs of FixL and FixJ in microorganisms that do not fix nitrogen (Crosson et al., 2005). In addition to FixLs, several classes of histidine-protein kinases employ alternative heme-binding domains to sense gaseous ligands (GillesGonzalez and Gonzalez, 2005). For example, the Mycobacterium tuberculosis DevS and DosT hypoxia sensors couple their kinase to a heme-binding GAF domain instead of the heme-binding PAS domain found in FixLs (Ioanoviciu et al., 2007; Sardiwal et al., 2005; Sousa et al., 2007b). Like FixL, these M. tuberculosis sensors phosphorylate a response-regulating transcription factor (DevR), and the resulting broad changes in gene expression trigger a state of nonreplicative persistence: in this case a latent infection of a human host (Roberts et al., 2004; Saini et al., 2004a,b). Clearly, the readily accessible ligand-binding and enzymatic parameters of heme-based O2 sensors make them ideal subjects for studies of signal transduction, and their involvement in relevant lifestyle changes of bacteria lends additional significance to their study.

2. Assays 2.1. General considerations 2.1.1. Autophosphorylation If a histidine-protein kinase is supplied with its ATP but not its protein substrate, it slowly (10–40 min) converts itself to a phosphorylated species (Hess et al., 1991; Stock et al., 1989). For example, the addition of ATP to deoxy-FixL yields a phospho-FixL species (Gilles-Gonzalez and Gonzalez, 1993; Tuckerman et al., 2001). Autophosphorylation FixL2 þ 2 ATP ⇆ P-FixL2 þ 2 ADP The so-called autophosphorylation is really a trans-phosphorylation between the subunits of these requisitely dimeric kinases (Ninfa et al., 1993). For sensors such as FixL, this reaction will reliably report qualitative information on

Ligands that regulate the kinase Preferred divalent cation for the enzyme Contamination of the enzyme preparation with a phosphatase

176

Marie-Alda Gilles-Gonzalez et al.

However, this reaction cannot give a quantitative measure of the enzymatic activity or its regulation because it is not a valid, rate-limiting, half-reaction. Contrary to the ping-pong bi–bi mechanism often presumed for many histidine-protein kinases, FixL does not process its nucleotide and protein substrates independently (Tuckerman et al., 2001, 2002). FixJ clearly enhances the initial FixL phosphorylation. For example, phosphorylation of RmFixL is accelerated about eightfold by the inclusion of an unphosphorylatable FixJ (the D54N RmFixJ variant) in the reaction (Sousa et al., 2005). More importantly, even in a large excess of ATP the reaction of FixL with only ATP stops when only about 20% of FixL is phosphorylated. In contrast, formation of the phosphorylated FixL intermediate proceeds essentially to completion if the true intermediate is trapped by including the unphosphorylatable FixJ substrate in the reaction (Sousa et al., 2005). Finally, when FixJ is added to phospho-FixL ‘‘preformed’’ by autophosphorylation, very little phosphoryl transfer occurs; instead, most of the phosphorylated protein is hydrolyzed to free phosphate. These observations imply that the ‘‘phospho-FixL’’ produced by these two methods are not kinetically equivalent (Tuckerman et al., 2001). This may also be the case for many other sensory kinases presumed to carry out their phosphoryl transfers sequentially and reported to show highly inefficient phosphoryl transfer to their protein substrate under those conditions. 2.1.2. Turnover Since FixL-catalyzed phosphoryl transfers clearly take place with FixL, FixJ, and ATP present, the turnover rate kcat provides the best measure of the FixL reaction kinetics and the effects of regulatory ligands on those kinetics (Sousa et al., 2007a; Tuckerman et al., 2002).

Turnover: 2 FixJ þ 2 ATP

FixL2 ⇆ P-FixJ2 þ 2 ADP

This rate represents the number of molecules of a specific substrate that one molecule of a specified form of an enzyme will phosphorylate per minute while it is saturated with all of its substrates. Consequently, an accurate kcat measurement requires that the kinase be kept at levels sufficiently high to ensure preservation of its dimeric state and yet sufficiently low to guarantee that the level of enzyme–substrate complex will not change during the measurement, i.e., the time for the enzyme to reach steady state and complete at least 10 turnovers. For the FixL–FixJ system, this means about 1 mM FixL, >20 mM FixJ, and 1.0 mM ATP. For kcat determinations, it is also essential to collect complete time courses so that the portion the time course may be found where product accumulates at a


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constant rate (i.e., when the rate of phosphorylation of the protein substrate matches the rate of replenishment of the phosphorylated enzyme intermediate). For inhibited states of the enzyme (e.g., FixL partially saturated with O2), there can be a lag while the enzyme intermediates build up. For highly active states (e.g., deoxy-FixL), there can be an early deceleration of the reaction rate due to rapid depletion of substrate. An informative indicator of the efficacy of a heme ligand is its inhibition factor, defined as the ratio of turnover rates of the unliganded and the fully liganded forms for the same oxidation state. For lowaffinity native and mutant sensors, saturation with O2 may not be practical. In these cases, great caution should be used in estimating the activity of a hypothetical fully oxygenated sensor, as the activity of partially saturated mixtures is not necessarily the sum of the activities of the liganded and unliganded species weighted by their relative abundance (Sousa et al., 2007a). In addition to the aforementioned considerations about steady state, determinations of inhibition factor require verifying that the heme status (saturation, oxidation state) remains unchanged throughout the measurements of reaction kinetics.

2.2. Protein purifications 2.2.1. Strategy for purifying heme-containing histidine-protein kinases Sensory kinases such as FixL and DosT are easily monitored from their intense red color. They readily yield to traditional methods of protein fractionation, with an anion-exchange step giving the most significant purification because of their unusually low isoelectric points (pI 5–6) (Scopes, 1994). Moderate overexpression of the corresponding genes at about 5% of total cell proteins from an inducible Escherichia coli promoter (e.g., tac) usually gives the highest yield of soluble heme protein. While it is possible to obtain much higher expression with some vectors, the resulting protein is usually of low quality, with weak enzymatic activity and a tendency to aggregate. It is far easier to grow more cells than to try to recover activity from misfolded or aggregated protein. Typically, a 4-liter culture of an E. coli lacIq strain (e.g., TG1) harboring the gene on a plasmid is grown overnight in a fermentor (Bioflow 3000) at 37 , 200–500 rpm, and 20% of atmospheric O2. When the culture reaches an OD600 of about 0.5, expression of the heme protein is induced with isopropyl-b-D-thiogalactoside (1 mM ). When growth of the cells begins to slow (cell density about 30 g/liter), they are cooled to 4 , harvested, and lysed by sonication in 2 volumes of lysis buffer [20 mM Tris-HCl, pH 8.0, 100 mM NaCl, 3 mM KCl, 1 mM EDTA, 10 mM b-mercaptoethanol, 0.04 mg/ml lysozyme, 0.17 mg/ml phenylmethylsulfonyl fluoride (added at room temperature from a 40-mg/ml solution in acetone)]. The lysate is cleared by centrifugation at 70,000 rpm (Ti 70 rotor, 30 min, 4 ).

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The clear-red lysate, kept at about 4 , is slowly brought to 1.2 M ammonium sulfate with stirring and dropwise addition of 4.0 M ammonium sulfate, 2 mM EDTA, pH 7.5, and centrifuged at 12,000 rpm (SS43 rotor, 4 ). Assuming the pellet density to be about 1 g/ml, the red precipitate is diluted threefold with 50 mM Tris-HCl, 50 mM NaCl, 5% (v/v) glycerol, and 10 mM b-mercaptoethanol, pH 7.5, and desalted on a size-exclusion column (Sephadex G-25, GE Healthcare) preequilibrated with 50 mM Tris-HCl, 50 mM NaCl, 5% (v/v) glycerol, and 10 mM b-mercaptoethanol, pH 7.5, at 4 . Subsequent tracking of the heme protein is done automatically from its 415-nm absorption (Bio-Rad QuadTec UV/vis detector). The protein mixture is chromatographed on an anion-exchange column (DEAE-Sephacel, Amersham), with thorough washing in 100 mM NaCl, and elution from a gradient of 100–300 mM NaCl in 50 mM Tris-HCl, 5% (v/v) glycerol, and 10 mM b-mercaptoethanol, pH 7.5, at 4 . The heme protein-containing fractions are further purified (to about 95% purity) by gel filtration (Superdex S-200, GE Healthcare) on a column preequilibrated with 50 mM Tris-HCl, 50 mM NaCl, and 5% (v/v) glycerol, pH 8.0, at 4 . Depending on the stability to oxidation of the purified protein, it will be a mixture of FeIII and FeIIO2 states, or entirely in the FeIII state (Gonzalez et al., 1998). Concentrate the protein to about 100 mM in a filtration unit (Amicon, 10-kDa membrane cutoff ) and store in aliquots of about 0.5 ml each at –70 . 2.2.2. Quality control Heme content The heme content of the purified proteins may be quantified by a pyridine hemochromogen assay, with hemin as the standard (Appleby, 1980). When bacterial heme-containing histidine-protein kinases are overproduced in E. coli, they are typically recovered with their full complement of heme (one heme per monomer). For comparison to the heme content, the protein concentration may be measured by the BCA protein assay (Pierce Biotechnology Inc.) with bovine serum albumin as the standard. Preparation of deoxy protein Incubate the purified protein (100 mM) for about 15 min with an anaerobic solution of 10 mM dithiothreitol in 50 mM Tris-HCl, 50 mM KCl, 5.0% (v/v) ethylene glycol, and 1 mM MgCl2, pH 8.0, inside of an anaerobic chamber (Coy Laboratory Products, Inc.). This procedure works well for heme-based sensors of relatively low redox potential (e.g., FixL, EcDos, DosT) and converts them fully to the deoxy state. If this approach does not yield the deoxy state within 15 min, reduce the protein with 1 equivalent of the stronger reducing agent sodium dithionite inside the anaerobic chamber and immediately remove this chemical and its by-products by gel filtration on a Sephadex-G25 column (about 3 ml) equilibrated with the same buffer as described earlier. If the protein is to be used in the deoxy state or mixed with CO, keep it in the aforementioned buffer. If the protein is to be mixed with O2, dilute


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the solution to less than 1 mM dithiothreitol; if it is to be mixed with NO, remove all of the reducing agent. Verify the quality of the preparation by recording the 350- to 700-nm absorption spectrum. Exploiting autophosphorylation to verify phosphatase contamination Transfer the deoxy-protein solution to a sealable cuvette and measure its 350- to 700-nm absorption. Assuming the 434-nm extinction (e434) of the deoxy state to be about 130 mM-1cm-1, adjust the protein concentration to 4–5 mM in 50 mM Tris-HCl, 50 mM KCl, 5.0% (v/v) ethylene glycol, and 1 mM MgCl2, pH 8.0, and equilibrate it at 23 . Separately prepare a series of Eppendorf tubes labeled for each time point and each containing 3.3 ml of ‘‘stop buffer’’ [10 mM EDTA, 2% (w/v) sodium dodecyl sulfate, 0.40 M Tris-HCl, 50% (v/v) glycerol, and 2% (v/v) b-mercaptoethanol, pH 6.8]. Begin the reactions by introducing ATP to a final concentration of 1.0 mM (unlabeled ATP from Sigma and [g-32P]ATP from Amersham Pharmacia Biotech, specific activity 0.21 Ci/mmol). At timed intervals (e.g., 1.5, 3.0, 6.0, 12, 24, and 48 min), withdraw a reaction aliquot from the cuvette (10 ml) and mix it rapidly with the stop buffer (3.3 ml) from the appropriate tube. The stopped reactions may now be transferred to air. Electrophorese 10 ml from each time point on an 11% (w/v) SDS polyacrylamide gel (Fig. 10.1A) (Laemmli, 1970). Spot 1 ml from the later time points (e.g., 12, 24, 48 min) onto a polyethyleneimine-cellulose thin-layer chromatographic (TLC) plate, about 2 cm from the bottom and 1 cm from the next time point; include a control on the plate containing only the radiolabeled ATP (Fig. 10.1B). After air drying the plate, develop it in a sealed TLC container with a 1-cm layer of 0.75 M NaH2PO4, pH 3.5. Quantify the levels of phosphorylated protein in the dried gels and of low-molecular weight species on the TLC plates with a phosphorimager (Bio-Rad Personal Molecular Imager FX). As standards for the quantification, spot 1-ml 10-fold dilutions of the stock ATP solution onto a strip of cellulose for development alongside the gel and TLC. If the protein is free of phosphatase, the TLC will show no significant generation of free phosphate (Pi) in 45 min, and the autoradiograph will show a continuous increase or a leveling of the autophosphorylation over the same period (Fig. 10.1). If the protein is contaminated with a phosphatase, free phosphate will be obvious from the TLC, and the level of protein phosphorylation will decline after reaching an apparent peak. For such preparations, further purification is advised. Verification of kinase activity The absence of contaminating phosphatase activity is necessary but not sufficient to ensure that the entire enzyme in a preparation is active. Neither is the initial rate of autophosphorylation a reliable indicator of enzyme quality. A healthy kinase preparation should yield about 20% phosphorylated kinase at equilibrium. If a modified protein

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A 0.5

1

1.5

3

6

12

24 48 min

DevS

B Pi

ATP

3 12 48 min ATP

DevS

Figure 10.1 Quality of the M. tuberculosis O2 sensor DevS is verified readily by autophosphorylation. The increasing autophosphorylation of deoxy-DevS even after 24 min from the autoradiograph (A) and the insignificant production of free phosphate even in 48 min from the polyethyleneimine TLC plates (B) show this preparation to be free of phosphatase contamination. From Sousa et al. (2007b).

substrate unable to accept phosphate is used, phosphorylation of the kinase should proceed nearly to completion. Any enzyme that does not meet these standards should be discarded if one wishes to have measurements that are both accurate and reproducible from batch to batch. 2.2.3. Strategy for purifying response-regulator substrates These proteins are best isolated by affinity purification. We have found that an epitope-tagged 6-His (Invitrogen) introduced at the N-terminal end of the gene by recombinant DNA methods works well for purifying these response regulators. The protein is purified as recommended by the manufacturer of the tag and affinity column (nickel-charged affinity resin) and assayed based on Western blotting and antibody recognition of the tag


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(Invitrogen). For quality control, examine phosphorylation of the protein in an extended turnover assay, with time points ranging from 1 to 30 min (see later). Quantitative phosphorylation of response regulators (i,e., FixJ) entails a phosphorylation in one subunit of a dimer (Da Re et al., 1999). Acceptable preparations of kinase and protein substrate should result in about 50% phosphorylation of the protein substrate at equilibrium (Sousa et al., 2007a).

2.3. Measuring the Kd for binding of ligand 2.3.1. Direct titration with O2 Preparation of O2-saturated buffer Prepare a solution of 1.3 mM O2 in 50 mM Tris-HCl, 50 mM KCl, and 5% (v/v) ethylene glycol at pH 8.0 by bubbling pure O2 for about 30 min through this buffer at room temperature (23 ) in a septum-sealed glass vial with a needle and an escape. Move the sealed vial to an anaerobic chamber. Transfer aliquots of the O2 solution from the vial to anaerobic buffer or the deoxy protein, as necessary, with a gas-tight Hamilton syringe. Basis spectra and exploratory titration Before starting a ligand titration, it is essential to record absorption spectra of the protein at 0 and 100% of saturation with the ligand. These spectra will serve as basis spectra for data analysis and will make clear the boundaries for titration. Prepare deoxy protein (4 mM ) as described earlier; place the protein solution in a septumsealed cuvette and record its 350- to 700-nm absorption spectrum. Add a concentrated aliquot of the deoxy heme protein (final concentration 4 mM ) to a septum-sealed solution of O2-saturated buffer (1.3 mM O2) with a gas-tight syringe and record the 350- to 700-nm absorption. Record both spectra in an Excel spreadsheet. An optimum linear combination of these spectra may be used to fit the spectrum resulting from each experimental titration (acceptable overall fit should be >0.98) and generate a percentage of saturation. If the spectrum cannot be closely fit to a linear combination of liganded and unliganded spectra, this means that some process other than ligand association is occurring and that some other species is contributing to spectra collected during titration. Most commonly, this means that the heme iron is oxidizing during titration, and the error spectrum is that of the ferric state. The reason that whole spectra should always be fitted, rather than a few wavelengths, is to detect the generation of extra species during the titration. One should not attempt to correct for oxidized protein or any third species; instead, find a way to control the interfering process and repeat the titration. Heme-based sensor variants with Kd values for O2 ranging from millimolar to submicromolar may be encountered; usually, there is no previous information on the expected affinity for O2. Therefore, a quick exploratory titration is recommended. Add five O2 concentrations, ranging

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from about 1 to 600 mM O2, sequentially to the protein within the same cuvette, minimizing the headspace as much as possible, and record the absorption spectrum resulting from each O2 concentration. Estimate the point of half saturation and design an experiment for exploring the most informative range of O2 concentrations (O2 required to give 10–100% saturation). Full titration A full titration experiment might require about 10 O2 concentrations to cover the range for 10–100% saturation and should be repeated at least twice (Fig. 10.2). To minimize escape of gases or concurrent reactions (e.g., oxidation), O2 titration points should be prepared individually and their absorption recorded immediately. For example, prepare a solution of a known O2 concentration (e.g., 2.0 mM O2); combine it with deoxy protein (1–4 mM final) in a 1-cm path length cuvette, taking care to minimize the headspace, and immediately seal the cuvette. Record A Fraction Oxy-DosT

1.00 0.75 Kd = 26 mM n = 1.0

0.50 0.25 0.00

0

100

200 300 O2 (mM)

400

500

Fraction Oxy-DevS

B 1.00 0.75 Kd = 3 mM n = 1.0

0.50 0.25 0.00 0

25

50 O2 (mM)

75 150 250

Figure 10.2 The Kd for binding of O2 can be directly determined by titration with ligand. (A) Ferrous M. tuberculosis DosT (2 mM) was titrated with 1^1200 mM O2 at pH 8.0 at 25. Saturation data were fit to a nonlinear Hill equation from which the Kd and Hill coefficient n were determined. (B) The Kd for binding of O2 to the higher-affinity M. tuberculosis DevS protein was determined similarly by titrating the deoxy form (2 mM) with 0.8^256 mM O2 under the same conditions as in A. From Sousa et al. (2007b).


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the 350- to 700-nm absorption of this mixture. Repeat this procedure for all the O2 concentrations covered by titration. If the protein is prone to oxidation, the most challenging aspect of the titration is to make certain that the spectrum for each concentration is recorded after thorough mixing but before any oxidation. Reducing agents to control oxidation should be avoided, especially at low O2 concentrations, as these will consume O2. If it is necessary to use reducing agents, one must verify that the O2 saturation does not change for that specific O2 concentration during the time needed to collect the spectrum. Saturation can be determined by comparing actual spectra to linear combinations of basis deoxy and oxy spectra. Treated data are plotted and fitted to a nonlinear Hill plot equation and to a quadratic single-binding equation using one’s favorite curve-fitting software (e.g., Microsoft Excel or GraphPad Prism). An alternative to titration in a sealed cuvette is to use a system equilibrated with a large headspace of gas (e.g., a tonometer) at a controlled gas concentration. This is how most of the early titrations of hemoglobin and myoglobin were done. This approach requires designing and fabricating custom glassware, but it may be easier than the sealed approach if one has fairly large volumes of protein available. 2.3.2. Direct titration with CO Determination of the Kd value for binding CO can be done by a procedure similar to the aforementioned determination of O2 affinity for most sensors because their Kd values for binding of CO typically fall in the 0.5 to 10 mM range. The CO-saturated buffer (1.0 mM) is prepared in a fume hood by bubbling pure CO for about 30 min through an anaerobic solution of 50 mM Tris-HCl, 50 mM KCl, and 5% (v/v) ethylene glycol at pH 8.0 and 25 in a gas-washing bottle or any septum-sealed glass vial with a needle and an escape. For basis spectra, record the 350- to 700-nm absorption of the deoxy heme protein (1 mM) and the identical concentration of ferrous protein in a 1.0 mM solution of CO. For titrations, transfer aliquots of the CO solution from a septum-sealed vial to a sealable cuvette with the deoxy protein via a gastight Hamilton syringe. 2.3.3. Competition titration with NO The much higher NO affinity of heme proteins forbids their direct titration with NO. Instead, carbon monoxy protein is prepared at a known CO concentration and then titrated with NO, which displaces the CO (Fig. 10.3). The Kd value for binding of NO to the heme protein can be calculated from the ‘‘apparent’’ Kd in the presence of CO and the precisely known Kd value for binding of CO. Data analysis is the same as that of an ordinary titration, except that the basis spectrum for the unliganded state is replaced by the spectrum of carbon monoxy protein. To prepare the solution of NO-saturated heme protein, first prepare anaerobic buffer [50 mM

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Fraction NO-DosT

A 1.00 0.75 0.50

Kd (NO) ~ 5 nM

0.25 0.00 0

2 4 6 8 NO (mM), in 240 mM CO

10

Fraction NO-DevS

B 1.00 0.75 0.50 Kd (NO) ~ 10 nM 0.25 0.00

0

6 12 18 24 NO (mM), in 10 mM CO

30

Figure 10.3 The Kd for binding of NO can be determined by competition against CO. (A) Ferrous M. tuberculosis DosT in 240 mM CO (Kd(CO) ¼ 0.94 mM, n ¼ 1.0) was titrated competitively with 0.50^9.0 mM NO at pH 8.0 at 25. (B) Ferrous M. tuberculosis DevS in 10 mM CO (Kd(CO) ¼ 0.036 mM, n ¼ 1.0) was titrated competitively with 0.50^28 mM NO under the same conditions as in A. From Sousa et al. (2007b).

Tris-HCl, 50 mM KCl, and 5% (v/v) ethylene glycol at pH 8.0 and 23 ] inside of an anaerobic chamber in a vessel fitted with a three-way valve. Maximize the surface area of the solution and do not fill more than one quarter of the vessel with liquid. Seal the vessel, transfer it to a fume hood, and connect it to nitrogen and NO tanks via the three-way valve. Briefly flow nitrogen through the line and reservoir and then switch to the NO tank and flow a gentle stream of NO through the reservoir for 30 min. Transfer the NOsaturated buffer (2.0 mM NO, 1 ml) from the reservoir to the anaerobic chamber in a gas-tight Hamilton syringe. Prepare a solution of 1 mM ferrous protein in 20–40 mM NO; the heme protein should be fully saturated with NO. The very popular NO-generating reagents are not suitable for quantitative analytical work because they do not allow concentrations of NO to be precisely controlled and determined and they generate reactive species other than NO. The importance of handling NO and its solutions properly cannot be overemphasized. The pure gas is extremely reactive and rapidly destroys most metals and plastics. Nitric oxide vessels and tanks should be kept in a fume

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hood. Only polytetrafluoroethylene, silicone rubber, stainless steel, and glass should be used in apparatus that contacts NO. All traces of O2 and other substances that will react with NO must be scrupulously excluded from all solutions. Otherwise, the very remarkable phenomena that you will certainly observe will be only indirectly because of NO. A serious leak will result in nitric acid generation, which can be easily verified by checking the sample pH. One may also generate highly oxidizing species such as peroxynitrite. A sure sign that there is a leak somewhere in the NO-handling apparatus, or that something in the buffer reacts with NO, is that the heme protein absorption spectrum irreversibly changes or the protein denatures. One should also be aware that the pure NO in the supply tank, even in the absence of contaminants, may disproportionate into N2O5 and N2 over time. In the event that a novel heme protein displays highly unusual behavior toward NO, one should always test a protein control such as myoglobin, whose reactions with NO are thoroughly known, before announcing one’s discovery in a press release.

2.4. Determination of turnover rates, kcat 2.4.1. General procedure For heme-controlled histidine-protein kinases such as FixL, assays of kcat measure the rates at which a sensor of known heme status phosphorylates its protein substrate (Fig. 10.4). Such studies are essential for quantifying the

Phospho-FixJ (pmol)

100 80 60 40

1.0 2.5 5.0 10 20 30 min

20 0 0

10

20 Time (min)

30

40

Figure 10.4 The best measure of kinase activity for B. japonicum FixL is its rate of turnover (kcat) of FixJ to phospho-FixJ. This reaction was catalyzed by deoxy-BjFixL (1 mM) for 25 mM BjFixJ and 1 mM ATP at pH 8.0 at 23. The kcat is 1.5 min-1 (gray line). Quantitative phosphorylation of FixJ occurred at equilibrium, verifying the excellent quality of the enzyme and substrates, with one-half of the BjFixJ (200 pmol) being phosphorylated within 30 min. From Sousa et al. (2007a).

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influence of the heme on the kinase. For example, a typical experiment might compare phosphorylation of a response regulator (e.g., FixJ or DevR) by the deoxy, oxy, or carbon monoxy states of its kinase partner (e.g., FixL or DevS). For the turnover of FixJ to phospho-FixJ, reaction mixtures typically contain 1 mM FixL and 25 mM FixJ in 50 mM Tris-HCl, 50 mM KCl, 50 mM MnCl2, 1 mM MgCl2, and 5% (v/v) ethylene glycol, pH 8.0. Reactions are started by introducing the ATP (i.e., unlabeled ATP from Sigma and [g-32P]ATP of specific activity 0.42 Ci/mmol from Amersham Pharmacia Biotech) (see Fig. 10.4). They are stopped at 1.0, 2.5, 5.0, 10, 20, and 30 min by mixing 10-ml aliquots of the reaction mixtures with 3.3 ml of ‘‘stop buffer’’ [10 mM EDTA, 4% (w/v) sodium dodecyl sulfate, 0.50 M Tris-HCl, 0.20 M NaCl, 50% (v/v) glycerol, and 2% (v/v) b-mercaptoethanol, pH 6.8]. The products are electrophoresed on 15% (w/v) polyacrylamide gels (Laemmli, 1970). The levels of phosphorylated FixJ protein in the dried gels are quantified with a phosphorimager (Bio-Rad Personal Molecular Imager FX). A good preparation of kinase, in the on state, should quantitatively phosphorylate its protein substrate at equilibrium (see Fig. 10.4). Interpretation of kinetic data such as these critically relies on the assumption that the heme state remains the same throughout every reaction time course. It is therefore essential to verify 350- to 700-nm absorption spectra before and after every time course. It is also important to sample a sufficient number of time points so that each turnover rate may be computed at steady state, i.e., when the rate of FixJ phosphorylation matches the rate of replenishment of the phospho-FixL intermediate. One must additionally ensure that the concentration of all substrates is constant or else well above Km throughout the period of the reaction time course on which rate measurements are based, as extraction of fundamental rate constants from the reaction time course will otherwise be prohibitively complex and error prone. 2.4.2. Sensors with partially saturated heme Prepare the liganded protein as described earlier. Calculate saturation based on linear regression using whole spectra. Again, verify that the heme state remains unchanged throughout every time course, make sure that the enzyme is saturated with its substrates during the entire reaction, and collect enough data points to ensure coverage of the turnover rates at steady state.

2.5. Novel heme-containing histidine-protein kinases The vast majority of proteins predicted to be heme-containing histidine-protein kinases, or even identified as FixLs from their genetic context or possession of heme, have not been examined for their enzymatic activity and regulation of this activity by heme ligands (Freitas et al., 2003; Gilles-Gonzalez and Gonzalez, 2005; Iyer et al., 2003; Sardiwal et al., 2005;


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Taylor and Zhulin, 1999). We anticipate that many of these sensors will soon be studied for signal transduction as the complete enzymes. For initial study of a novel heme-containing kinase, additional considerations should be divalent-cation and heme-state requirements for activity. Although most histidine-protein kinases work well with MgII, others display strong preferences for specific divalent cations (Hess et al., 1991). For example, S. meliloti FixL performs best in assay mixtures that include MnII (50 mM) (Gilles-Gonzalez and Gonzalez, 1993). Likewise, although all hemecontaining kinases examined so far work best in the deoxy state, it is quite possible that this represents the ‘‘off ’’ state of some sensors, and inclusion of a ligand in the assays is necessary to manifest the ‘‘on’’ state.

ACKNOWLEDGMENTS This work was supported by NSF Grant 620531, the National Research Initiative of the USDA Cooperative State Research, Education and Extension Service, Grant 2002–35318– 14039, and Welch Foundation Grant I-1575.

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Gilles-Gonzalez, M. A. (2001). Oxygen signal transduction. IUBMB Life 51, 165–173. Gilles-Gonzalez, M. A., Caceres, A. I., Sousa, E. H., Tomchick, D. R., Brautigam, C., Gonzalez, C., and Machius, M. (2006). A proximal arginine R206 participates in switching of the Bradyrhizobium japonicum FixL oxygen sensor. J. Mol. Biol. 360, 80–89. Gilles-Gonzalez, M. A., Ditta, G. S., and Helinski, D. R. (1991). A haemoprotein with kinase activity encoded by the oxygen sensor of Rhizobium meliloti. Nature 350, 170–172. Gilles-Gonzalez, M. A., and Gonzalez, G. (1993). Regulation of the kinase activity of heme protein FixL from the two-component system FixL/FixJ of Rhizobium meliloti. J. Biol. Chem. 268, 16293–16297. Gilles-Gonzalez, M. A., and Gonzalez, G. (2005). Heme-based sensors: Defining characteristics, recent developments, and regulatory hypotheses. J. Inorg. Biochem. 99, 1–22. Gilles-Gonzalez, M. A., Gonzalez, G., Perutz, M. F., Kiger, L., Marden, M. C., and Poyart, C. (1994). Heme-based sensors, exemplified by the kinase FixL, are a new class of heme protein with distinctive ligand binding and autoxidation. Biochemistry 33, 8067–8073. Gong, W., Hao, B., Mansy, S. S., Gonzalez, G., Gilles-Gonzalez, M. A., and Chan, M. K. (1998). Structure of a biological oxygen sensor: A new mechanism for heme-driven signal transduction. Proc. Natl. Acad. Sci. USA 95, 15177–15182. Gonzalez, G., Gilles-Gonzalez, M. A., Rybak-Akimova, E. V., Buchalova, M., and Busch, D. H. (1998). Mechanisms of autoxidation of the oxygen sensor FixL and Aplysia myoglobin: Implications for oxygen-binding heme proteins. Biochemistry 37, 10188–10194. Hao, B., Isaza, C., Arndt, J., Soltis, M., and Chan, M. K. (2002). Structure-based mechanism of O2 sensing and ligand discrimination by the FixL heme domain of Bradyrhizobium japonicum. Biochemistry 41, 12952–12958. Hess, J. F., Bourret, R. B., and Simon, M. I. (1991). Phosphorylation assays for proteins of the two-component regulatory system controlling chemotaxis in Escherichia coli. Methods Enzymol. 200, 188–204. Ioanoviciu, A., Yukl, E. T., Moenne-Loccoz, P., and Montellano, P. R. (2007). DevS, a heme-containing two-component oxygen sensor of Mycobacterium tuberculosis. Biochemistry 46, 4250–4260. Iyer, L. M., Anantharaman, V., and Aravind, L. (2003). Ancient conserved domains shared by animal soluble guanylyl cyclases and bacterial signaling proteins. BMC Genomics 4, 5–12. Laemmli, U. K. (1970). Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685. Lois, A. F., Weinstein, M., Ditta, G. S., and Helinski, D. R. (1993). Autophosphorylation and phosphatase activities of the oxygen-sensing protein FixL of Rhizobium meliloti are coordinately regulated by oxygen. J. Biol. Chem. 268, 4370–4375. Monson, E. K., Weinstein, M., Ditta, G. S., and Helinski, D. R. (1992). The FixL protein of Rhizobium meliloti can be separated into a heme-binding oxygen-sensing domain and a functional C-terminal kinase domain. Proc. Natl. Acad. Sci. USA 89, 4280–4284. Mukai, M., Nakamura, K., Nakamura, H., Iizuka, T., and Shiro, Y. (2000). Roles of Ile209 and Ile210 on the heme pocket structure and regulation of histidine kinase activity of oxygen sensor FixL from Rhizobium meliloti. Biochemistry 39, 13810–13816. Ninfa, E. G., Atkinson, M. R., Kamberov, E. S., and Ninfa, A. J. (1993). Mechanism of autophosphorylation of Escherichia coli nitrogen regulator II (NRII or NtrB): Trans-phosphorylation between subunits. J. Bacteriol. 175, 7024–7032. Olson, J. S., and Phillips, G. N. (1997). Myoglobin discriminates between O2, NO, and CO by electrostatic interactions with the bound ligand. J. Biol. Inorg. Chem. 2, 544–552.


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Reyrat, J. M., David, M., Blonski, C., Boistard, P., and Batut, J. (1993). Oxygen-regulated in vitro transcription of Rhizobium meliloti nifA and fixK genes. J. Bacteriol. 175, 6867–6872. Roberts, D. M., Liao, R. P., Wisedchaisri, G., Hol, W. G., and Sherman, D. R. (2004). Two sensor kinases contribute to the hypoxic response of Mycobacterium tuberculosis. J. Biol. Chem. 279, 23082–23087. Saini, D. K., Malhotra, V., Dey, D., Pant, N., Das, T. K., and Tyagi, J. S. (2004a). DevRDevS is a bona fide two-component system of Mycobacterium tuberculosis that is hypoxiaresponsive in the absence of the DNA-binding domain of DevR. Microbiology 150, 865–875. Saini, D. K., Malhotra, V., and Tyagi, J. S. (2004b). Cross talk between DevS sensor kinase homologue, Rv2027c, and DevR response regulator of Mycobacterium tuberculosis. FEBS Lett. 565, 75–80. Sardiwal, S., Kendall, S. L., Movahedzadeh, F., Rison, S. C., Stoker, N. G., and Djordjevic, S. (2005). A GAF domain in the hypoxia/NO-inducible Mycobacterium tuberculosis DosS protein binds haem. J. Mol. Biol. 353, 929–936. Sciotti, M. A., Chanfon, A., Hennecke, H., and Fischer, H. M. (2003). Disparate oxygen responsiveness of two regulatory cascades that control expression of symbiotic genes in Bradyrhizobium japonicum. J. Bacteriol. 185, 5639–5642. Scopes, R. K. (1994). ‘‘Protein Purification Principles and Practice,’’ 3rd ed. SpringerVerlag, New York. Soupene, E., Foussard, M., Boistard, P., Truchet, G., and Batut, J. (1995). Oxygen as a key developmental regulator of Rhizobium meliloti N2 fixation gene expression within the alfalfa root nodule. Proc. Natl. Acad. Sci. USA 92, 3759–3763. Sousa, E. H. S., Gonzalez, G., and Gilles-Gonzalez, M.-A. (2005). Oxygen blocks reaction of the FixL/FixJ complex with ATP but does not influence binding of FixJ or ATP to FixL. Biochemistry 44, 15359–15365. Sousa, E. H. S., Tuckerman, J. R., Gonzalo, G., and Gilles-Gonzalez, M.-A. (2007a). A memory of oxygen binding explains the dose response of the heme-based sensor FixL. Biochemistry 46, 6249–6257. Sousa, E. H. S., Tuckerman, J. R., Gonzalo, G., and Gilles-Gonzalez, M.-A. (2007b). DosT and DevS are oxygen-switched kinases in Mycobacterium tuberculosis. Protein Sci. 16, 1708–1719. Stock, J. B., Ninfa, A. J., and Stock, A. M. (1989). Protein phosphorylation and regulation of adaptive responses in bacteria. Microbiol. Rev. 53, 450–490. Taylor, B. L., and Zhulin, I. B. (1999). PAS domains: Internal sensors of oxygen, redox potential, and light. Microbiol. Mol. Biol. Rev. 63, 479–506. Tuckerman, J. R., Gonzalez, G., Dioum, E. M., and Gilles-Gonzalez, M. A. (2002). Ligand and oxidation-state specific regulation of the heme-based oxygen sensor FixL from Sinorhizobium meliloti. Biochemistry 41, 6170–6177. Tuckerman, J. R., Gonzalez, G., and Gilles-Gonzalez, M. A. (2001). Complexation precedes phosphorylation for two-component regulatory system FixL/FixJ of Sinorhizobium meliloti. J. Mol. Biol. 308, 449–455. Virts, E. L., Stanfield, S. W., Helinski, D. R., and Ditta, G. S. (1988). Common regulatory elements control symbiotic and microaerobic induction of nifA in Rhizobium meliloti. Proc. Natl. Acad. Sci. USA 85, 3062–3065.


C H A P T E R

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Reactions of Nitric Oxide and Oxygen with the Regulator of Fumarate and Nitrate Reduction, a Global Transcriptional Regulator, during Anaerobic Growth of Escherichia coli Jason C. Crack,* Nick E. Le Brun,* Andrew J. Thomson,* Jeffrey Green,† and Adrian J. Jervis† Contents 1. Introduction 2. Production of 4Fe-FNR Protein 2.1. Purification of native 4Fe-FNR 2.2. Purification of reconstituted 4Fe-FNR 2.3. Cleaning and concentration of 4Fe-FNR 3. Determination of Iron and Acid-Labile Sulfide Content of FNR 3.1. Determination of iron content 3.2. Determination of sulfide content 4. UV-Visible Absorbance Spectra of FNR 5. Cluster Reaction with Nitric Oxide and Oxygen 5.1. Reaction of 4Fe-FNR with nitric oxide 5.2. Reaction of 4Fe-FNR with oxygen 6. Purification of 2Fe-FNR 7. Detection of Other Reaction Products 7.1. Detection of sulfide 7.2. Detection of iron 7.3. Detection of superoxide and peroxide 8. Conclusions References

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Centre for Metalloprotein Spectroscopy and Biology, School of Chemical Sciences and Pharmacy, University of East Anglia, Norwich, United Kingdom Department of Molecular Biology and Biotechnology, University of Sheffield, Sheffield, United Kingdom


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Abstract The Escherichia coli fumarate and nitrate reductase (FNR) regulator protein is an important transcriptional regulator that controls the expression of a large regulon of more than 100 genes in response to changes in oxygen availability. FNR is active when it acquires a [4Fe-4S]2þ cluster under anaerobic conditions. The presence of the [4Fe-4S]2þ cluster promotes protein dimerization and sitespecific DNA binding, facilitating activation or repression of target promoters. Oxygen is sensed by the controlled disassembly of the [4Fe-4S]2þ cluster, ultimately resulting in inactive, monomeric, apo-FNR. The FNR [4Fe-4S]2þ cluster is also sensitive to nitric oxide, such that under anaerobic conditions the protein is inactivated by nitrosylation of the iron-sulfur cluster, yielding a mixture of monomeric and dimeric dinitrosyl-iron cysteine species. This chapter describes some of the methods used to produce active [4Fe-4S] FNR protein and investigates the reaction of the [4Fe-4S]2þ cluster with nitric oxide and oxygen in vitro.

1. Introduction Fumarate and nitrate reductase (FNR) regulator proteins are members of the cAMP receptor protein (CRP)/FNR superfamily of transcription regulators (Korner et al., 2003). All members of the family are structurally related to the founder member (Schultz et al., 1991). Thus, CRP/FNR family members share a basic two-domain structure consisting of an N-terminal sensory domain and a C-terminal DNA-binding domain. The versatility of this framework is illustrated by the range of metabolic and environmental signals perceived by different members of the family and by the variety of physiological responses that they control (Green et al., 2001). The best-characterized FNR protein is that of Escherichia coli. In E. coli FNR acts as a direct oxygen sensor and is the primary transcriptional regulator of the switch between aerobic and anaerobic growth (Green et al., 2001; Guest, 1992, 1995; Sawers, 1999). Thus, under anaerobic conditions FNR is in its active state and is able to bind specific palindromic sequences of DNA (Eiglmeier et al., 1989; Green et al., 1996b; Lazazzera et al., 1993, 1996). Once bound to DNA, FNR activates target gene expression by recruiting RNA polymerase (RNAP) or represses transcription by preventing the formation of productive RNAP:DNA complexes (Barnard et al., 2003; Wing et al., 1995). Generally, FNR activates genes encoding products involved in anaerobic metabolism, such as the nar operon (nitrate reductase), the dms operon (dimethyl sulfoxide reductase), and the frd operon (fumarate reductase), and represses genes encoding products involved in aerobic

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metabolism, such as the sdh operon (succinate dehydrogenase) and ndh (NADH dehydrogenase II). FNR is activated under anaerobic conditions by the acquisition of one [4Fe-4S]2þ cluster per monomer (designated 4Fe-FNR). Each [4Fe-4S]2þ cluster is ligated by four cysteine residues (Cys20, Cys23, Cys29, and Cys122), and the presence of the cluster promotes FNR dimerization, increasing its capacity to bind specifically to DNA (Green et al., 1996a; Khoroshilova et al., 1997; Kiley and Beinert, 1999; Lazazzera et al., 1993; 1996; Melville and Gunsalus, 1990; Sharrocks et al., 1990). When exposed to oxygen the [4Fe-4S]2þ cluster is oxidized to a [2Fe-2S]2þ cluster, via a transient [3Fe-4S]1þ form, resulting in the formation of 2Fe-FNR (Crack et al., 2004, 2007; Jordan et al., 1997; Khoroshilova et al., 1995; Lazazzera et al., 1996; Popescu et al., 1998; Sutton et al., 2004a). If aerobic conditions persist, the [2Fe-2S]2þ cluster is further degraded, yielding apo-FNR (Achebach et al., 2005; Green et al., 1991; Sutton et al., 2004b; Unden and Schirowski, 1997). In contrast to 4Fe-FNR, apo-FNR and 2Fe-FNR are monomeric and are unable to bind efficiently to DNA and are therefore inactive (Green et al., 1991; Lazazzera et al., 1996; Sutton et al., 2004a,b). If anaerobic conditions return, 4Fe-FNR is generated either by recycling apo-FNR or 2Fe-FNR or by de novo synthesis of 4Fe-FNR (Dibden and Green, 2005; Khoroshilova et al., 1997; Mettert and Kiley, 2005). In addition to its reaction with oxygen, the FNR [4Fe-4S]2þ cluster is also sensitive to nitric oxide (NO). Upon exposure to NO the [4Fe-4S]2þ cluster becomes nitrosylated, forming a combination of monomeric and dimeric dinitrosyl-iron-cysteine (DNIC) complexes, again abolishing its ability to bind DNA (Cruz-Ramos et al., 2002). Among the genes repressed by FNR under anaerobic conditions is hmp, which encodes the flavohemoglobin that is one of the major mechanisms for detoxifying NO in E. coli (Poole, 2005; Poole et al., 1996). When cultures of E. coli are exposed to NO, FNR repression of hmp is relieved, suggesting that the reaction between FNR and NO is physiologically significant (Cruz-Ramos et al., 2002). This suggestion is supported by transcript profiling experiments that reveal that the abundances of many FNR-activated genes are lower, and many FNR-repressed genes are greater, when E. coli is exposed to NO (Justino et al., 2005; Pullan et al., 2007). Thus, 4Fe-FNR is a sensor of both oxygen and NO; consequently, purification and manipulation in vitro must be carried out under strictly anoxic conditions. This chapter provides information on how to isolate 4Fe-FNR and study its reaction with oxygen and NO. It is hoped that some of the techniques will be transferable, such that they are of value not only for those interested in FNR proteins, but also for those working with proteins possessing labile iron-sulfur clusters.

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2. Production of 4Fe-FNR Protein Two current methods are used to obtain functional, anaerobic 4Fe-FNR. The first is to overproduce and purify native 4Fe-FNR under anaerobic conditions (Crack et al., 2006; Sutton et al., 2003). The second is to produce apo-FNR under aerobic conditions and then reconstitute the purified protein with the [4Fe-4S]2þ cluster under anaerobic conditions in vitro (Crack et al., 2004; Green et al., 1996a). There has been some debate centered around whether reconstituted 4Fe-FNR behaves in the same way as 4Fe-FNR that has been isolated under anaerobic conditions, although evidence indicates that there is essentially no difference in their reaction with oxygen (Crack et al., 2006, 2007). Regardless of the method used to produce anaerobic 4Fe-FNR, the protein is very oxygen sensitive, and hence certain stages of production, purification, and storage must be carried out under strictly anaerobic conditions. To achieve this, anaerobic cabinets (Belle Technology/Don Whitley) typically operating at 2.0 ppm oxygen are required. To help maintain anaerobic conditions in the cabinets, all buffers are sparged with oxygen-free nitrogen gas for a minimum of 2 h before introduction into the anaerobic cabinet. In addition, plastic items are equilibrated in the anaerobic cabinet for a minimum of 24 h before being used. Furthermore, it is beneficial if the anaerobic cabinets are also equipped with a protein purification system, a fridge-freezer for anaerobic sample storage (Belle Technology), and a liquid nitrogen access port.

2.1. Purification of native 4Fe-FNR The fnr gene was polymerase chain reaction amplified using oligonucleotide primers that introduced a unique upstream NcoI site and included the native downstream BamHI site. The amplified DNA was then ligated into the commercially available vector pET11d (Novagen) to create pGS1859 (Crack et al., 2006). This expression plasmid has a copy of fnr under the control of the inducible T7 promoter. Native FNR is produced from cultures of E. coli BL21lDE3 containing pGS1859 as described by Sutton and Kiley (2003) with the following modifications. Cultures that have been induced for FNR production are subsequently incubated on ice for 5 to 10 min and supplemented with 200 mM ferric ammonium citrate, 50 mM L-methionine, and 0.001% (v/v) antifoam 204 (Sigma) prior to sparging with oxygen-free nitrogen gas at 4 overnight. Unless otherwise stated, all subsequent work and manipulations are performed under strictly anaerobic conditions. The sparged cultures are transferred to O-ring-sealed centrifuge tubes (Beckman), and bacteria are collected by centrifugation outside the anaerobic cabinet at 6500 g for 10 min at 4 .


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From a total culture volume of 5 liters the bacterial cell pellets are resuspended in 105 ml of buffer A [10 mM potassium phosphate, 100 mM KCl, 10% (v/v) glycerol, pH 6.8]. Then 12 ml of CelLytic lysis buffer (CelLytic B 10; Sigma), lysozyme (21.3 mg ml-1), DNase I (0.5 mg ml-1), phenylmethylsulfonyl fluoride (PMSF, 0.1 mM), and benzamidine (1 mM ) are added (total volume 120 ml). The suspension is mixed gently and incubated at room temperature for up to 20 min. The cell lysate is then transferred to O-ring-sealed centrifuge tubes (Nalgene) and centrifuged outside the anaerobic cabinet at 40,000 g for 45 min at 2 . The resulting cell-free extract is made 2% (v/v) with buffer B [10 mM potassium phosphate, 800 mM KCl, 10% (v/v) glycerol, pH 6.8] and is applied to a 10-ml HiTrap SP FF cation-exchange column (GE Healthcare) equilibrated previously with buffer A. After application of the extract, the column is washed with 15% (v/v) buffer B (in buffer A), and bound proteins are eluted using a linear gradient (60 ml) of 15 to 100% (v/v) buffer B. Fractions containing FNR are pooled, diluted threefold with buffer C [10 mM potassium phosphate, 10% (v/v) glycerol, pH 6.8], and loaded onto a 2-ml HiTrap heparin column (GE Healthcare) equilibrated previously with 5% (v/v) buffer B. The column is washed with 10% (v/v) buffer B, and bound protein is eluted using a linear gradient (50 ml) from 10 to 100% (v/v) buffer B. Fractions containing FNR are pooled, concentrated if necessary (see later), and stored in glass vials in an anaerobic fridge-freezer until needed.

2.2. Purification of reconstituted 4Fe-FNR In this method, FNR is produced aerobically with an N-terminal GST tag and then purified under anaerobic conditions by affinity purification using a glutathione-Sepharose column (GE Healthcare). On column cleavage of the GST-FNR fusion protein results in apo-FNR, which contains no cluster. To obtain 4Fe-FNR, apo-FNR is reconstituted with a [4Fe-4S]2þ cluster by in vitro incubation with L-cysteine and NifS [a cysteine desulfurase purified as described by Zheng and co-workers (1993)] as a source of sulfur and ammonium ferrous sulfate as a source of iron. To create the GST-FNR fusion protein, the same fnr-containing DNA fragment used for native FNR production (see earlier discussion) is ligated onto vector pGEX-KG (Guan and Dixon, 1991) to create plasmid pGS572 (Green et al., 1996b). E. coli BL21lDE3 pGS572 cultures are grown in 2.5-liter conical flasks containing 500 ml L-broth (Sambrook and Russell, 2001) medium supplemented with ampicillin (100 mg liter-1) at 37 with shaking at 250 rpm. When cultures reach A600 0.6, the production of GST-FNR is induced by the addition of 1 mM isopropyl-b-thiogalactopyranoside and incubation is continued for 3–4 h. The cultures are then transferred to O-ring-sealed centrifuge tubes (Beckman), and bacteria are

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pelleted at 18,000 g for 10 min. The supernatant is discarded, and the cell pellet is either used immediately or stored at –80 until needed. Cell pellets are resuspended in 35 ml of buffer D (25 mM HEPES, 2.5 mM CaCl2, 100 mM NaCl, 100 mM NaNO3, pH 7.5) per liter of culture, and 1 mM benzamidine and 0.1 mM PMSF are added before cell lysis by passage through a French pressure cell three times at 20,000 psi or by sonication in the presence of lysozyme (21.3 mg ml-1). Unbroken cells are removed by centrifugation at 40,000 g for 20 min, and the resulting supernatant is then filtered through a 0.45-mm pore syringe filter. To isolate FNR from extracts containing GST-FNR, all steps are carried out in an anaerobic cabinet. Aliquots of the filtered supernatant from 1 liter of culture are applied to three columns containing 3 ml glutathioneSepharose (GE Healthcare) preequilibrated with 60 ml buffer D. Once the extract has passed through the column, the matrix is washed with 60 ml of buffer D. Once washed, the column is capped and 10 units of thrombin (Sigma) in 0.5 ml buffer D is added and mixed into the matrix bed. The column is then incubated at an ambient temperature (21 ) overnight to allow release of FNR from the bound GST-FNR fusion, by specific cleavage of the linker sequence joining GST to FNR. The anaerobic apo-FNR protein is eluted by the addition of 4 ml buffer D in 1-ml aliquots, which are collected as 1-ml fractions. Fractions are then pooled, giving a total of 12 ml eluent from 1 liter of culture. A 200-ml sample of the pooled fractions is taken for protein assays (Bio-Rad) to determine its concentration (usually close to 80 mM) using a correction factor of 0.83 (Green et al., 1996a). Purity is usually assessed by SDS-PAGE analysis. To reconstitute the iron-sulfur cluster into apo-FNR solutions, FeS-1 (50 mM L-cysteine, 125 mM dithiothreitol, 25 mM HEPES, 2.5 mM CaCl2, 100 mM NaCl, 100 mM NaNO3, pH 7.5) and FeS-2 [20 mM (NH4)2Fe(SO4)2, 25 mM HEPES, 2.5 mM CaCl2, 100 mM NaCl, 100 mM NaNO3, pH 7.5] are freshly prepared under anaerobic conditions using anaerobic buffer D to dissolve the solutes. Reconstitution of the ironsulfur cluster is initiated by the addition of an aliquot of FeS-1 (1 mM L-cysteine, 2.5 mM dithiothreitol, final concentrations), an appropriate amount of FeS-2, such that there is a 7 M excess of Fe2þ ions per FNR monomer, and an aliquot of NifS (225 nM final concentration). Small-scale (¼3 ml) reconstitutions are carried out in anaerobic cuvettes sealed with a screw cap containing a septa made from rubber and coated with a layer of Teflon (Hellma UK, Ltd.). Once sealed, the cuvettes may be removed from the anaerobic cabinet and transferred to a spectrophotometer fitted with a thermostatic cell holder to follow the progress of the reconstitution reaction. Larger reconstitutions are carried out inside the anaerobic cabinet using a small conical flask fitted with a water jacket. The reaction is sealed using a rubber stopper.


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Reconstitutions, via either method, are stirred (using a magnetic follower) throughout and typically are complete after 4.5 h at 37 . Reconstituted FNR protein is then cleaned and concentrated using a 1-ml HiTrap heparin column (GE Healthcare) as described later.

2.3. Cleaning and concentration of 4Fe-FNR After reconstitution, 4Fe-FNR is separated from low molecular weight components using a 1-ml HiTrap heparin column (GE Healthcare). This step can also be used to concentrate dilute preparations of 4Fe-FNR from either purification method. The 4Fe-FNR binds to the columns at low KCl concentrations (100 mM ), but is eluted at higher concentrations (>400 mM KCl). All stages are carried out under strictly anaerobic conditions. The heparin column is equilibrated with 10 ml of the appropriate buffer (i.e., buffer A for native preparations and buffer D for reconstitutions, see earlier discussion). The pooled FNR fractions are then passed through a 0.22-mm filter onto the column and washed with 10–20 ml of the equilibration buffer to remove low molecular weight species. The column is then inverted, and FNR is eluted from the end of the column to which it was applied, using buffer (typically buffer A or D) containing 500 mM KCl. The eluent is collected in microcentrifuge tubes, four drops at a time to obtain the highest concentration. At low concentrations 4Fe-FNR has a straw brown color but is green/black at high concentrations. The most concentrated fractions are pooled together. The concentration of 4Fe-FNR present in the final preparation is determined by assaying for protein (described earlier), iron, acid-labile sulfide (see later), and spectrophotometrically assuming e405 nm of 16,200 M-1 cm-1 per [4Fe-4S]2þ cluster (Crack et al., 2006).

3. Determination of Iron and Acid-Labile Sulfide Content of FNR 3.1. Determination of iron content The iron content of FNR is determined as follows: 0.1 ml of 21.7% HNO3 is added to the same volume of protein and incubated at 95 for 30 min. Cooled samples are centrifuged to remove any precipitate, treated with 0.6 ml of 7.5% (w/v) ammonium acetate, 0.1 ml of 12.5% (w/v) ascorbic acid, and 0.1 ml of 10 mM ferene (5,50 (3-(2-pyridyl)-1,2,4-triazine5,6-diyl)-bis-2-furansulfonate), mixed, and incubated at room temperature for 30 min. The concentration of the ferene-iron complex is calculated from measurements of its absorbance at 593 nm. Iron concentrations are

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determined by reference to a calibration curve generated from Fe(III) solutions in the range of 0–200 mM, prepared from SpectrosoL standard iron solution (BDH), and treated as described earlier.

3.2. Determination of sulfide content Acid-labile sulfide is determined, via the formation of methylene blue, according to the method of Beinert (1983). Briefly, 0.2 ml of appropriately diluted sample is treated with 0.6 ml of freshly prepared 1% (w/v) zinc acetate and 50 ml of 12% (w/v) NaOH. The sample tube is capped, inverted, and incubated at room temperature for 15 min. After incubation the sample is centrifuged at 3000 rpm for 60 s, treated with 0.15 ml 0.1% (w/v) N,N-dimethyl-p-phenylenediamine dihydrochloride dissolved in 5 M HCl and 0.15 ml 10 mM FeCl3 dissolved in 1 M HCl. The sample is then vortexed for 30 s and incubated at room temperature for a further 30 min. The concentration of the acid-labile sulfide is calculated from the formation of methylene blue measured through its absorbance at 670 nm. Sulfide concentrations are determined by reference to a calibration curve generated from anaerobic Na2S solutions containing 10 mM NaOH in the range of 0–300 mM, according to the method of Beinert (1983), and treated as described earlier.

4. UV-Visible Absorbance Spectra of FNR The state of FNR can be followed using UV-visible spectroscopy as the [4Fe-4S]2þ cluster has characteristic spectral features. Apo-FNR typically exhibits only protein-associated absorbance at 280 nm, whereas 4Fe-FNR shows absorbance maxima at 320 nm (e ¼ 20,075 350 M-1 cm-1) and 405 nm (e ¼ 16,200 135 M-1 cm-1) with a characteristic broad shoulder around 420 nm. Examples of apo-FNR and of [4Fe-4S] FNR spectra are shown in Fig. 11.1.

5. Cluster Reaction with Nitric Oxide and Oxygen FNR acts as both a NO and an oxygen sensor via its [4Fe-4S]2þ cluster (see references given earlier). Much research has been performed to investigate the reaction of the cluster with molecular oxygen, but less on its reaction with NO. The two main types of experiment aim to determine the rates and stoichiometries of the reactions.

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35,000 1.4 30,000 Absorbance

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Figure 11.1 UV-visible spectrum of isolated 4Fe-FNR. (Inset) Changes in the UV-visible spectrum of apo-FNRduring reconstitution of 4Fe-FNR.

When 4Fe-FNR is reacted with either NO or oxygen the electronic spectrum changes, most notably at 420 nm, because of alteration of the [4Fe4S]2þ cluster. To follow these changes, reactions are generally carried out in anaerobic cuvettes sealed with a screw cap containing a septum made from rubber and coated with a layer of Teflon (as used in the reconstitution reaction). The seal allows the injection of buffer containing known concentrations of the appropriate reactant (NO or oxygen) after which the electronic spectrum of the protein can be obtained, or samples can be removed for electron paramagnetic resonance (EPR) spectroscopy. Reactions of FNR with NO or oxygen are fast, typically being complete within 5 min at 20 .

5.1. Reaction of 4Fe-FNR with nitric oxide Generally the fast NO-releasing compound proline NONOate is used as a source of NO, although NO solutions can be used (Cruz-Ramos et al., 2002). Each NONOate molecule releases 1.9 molecules of NO, with a half-life of 13 s in the working buffer at 25 (Cruz-Ramos et al., 2002). An anoxic 4.17 mM stock solution of proline NONOate is made just before use in 25 mM Tris, pH 10.0, 100 mM NaCl and stored at –1 until needed. Typically 1 ml of 4Fe-FNR with a cluster concentration of 30 mM is transferred to a cuvette in an anaerobic cabinet. Then fixed concentrations of proline NONOate are introduced into the cuvette by injection through

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the septum, and the solution is mixed. The reaction of NO with 4Fe-FNR is very rapid and is essentially dependent on the release of NO from proline NONOate at 25 . Reaction with NO (Fig. 11.2A) results in the appearance A

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Figure 11.2 Spectral changes upon reaction of 4Fe-FNR with NO. (A) UV-visible spectra of anaerobic 4Fe-FNR (27 mM ) before (thin line) and after (thick line) treatment with NO (140 mM ). (B) EPR spectra at 77 K of anaerobic 4Fe-FNR treated with NO as indicated. The microwave power was 2.000 mW and the frequency was 9.669 GHz. Modulation amplitude, frequency, and receiver gain were 10 gauss, 100 kHz, and 1 105. For both experiments the buffer used was 25 mM HEPES, 2.5 mM CaCl2, 100 mM NaCl, and 100 mM NaNO3, pH 7.5.


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of a shoulder at 360 nm in the UV-visible spectrum together with loss of the broad 420-nm shoulder. An isosbestic point is observed at 480 nm (CruzRamos et al., 2002). The final spectrum corresponds to a mix of monomeric and dimeric DNIC species in an approximate 20:80 ratio, giving a relatively intense yellow color to the solution. Because the product of reaction of 4Fe-FNR with NO is stable for several hours, the stoichiometry of the reaction can be determined by adding limiting amounts of NONOate, allowing it to react, and then recording spectra. By plotting the dA360 against NO:[4Fe-4S]2þ it was found that the reaction of NO and FNR is complete at a ratio of 3:1 (Cruz-Ramos et al., 2002). Determining the rate of reaction when using proline NONOate is difficult, as the NO is not immediately available for reaction when introduced. To avoid this, aqueous solutions of dissolved NO gas could be used. However, some protein precipitation has been observed, resulting in baseline changes, while using aqueous NO solutions. Another useful tool for investigating the reaction of 4Fe-FNR with NO is EPR spectroscopy. The EPR system used consists of an X-band Bruker EMX spectrometer equipped with an ESR-900 helium flow cryostat and a TE-102 microwave cavity (Oxford Instruments). The spin intensities of paramagnetic samples can be quantified by the integration of EPR spectra using 1 mM Cu(II), 10 mM EDTA as the standard. Neither apo-FNR nor 4Fe-FNR exhibits an EPR signal; however, samples of 4Fe-FNR reacted with NO, as described earlier ([NO]:[4Fe-4S]2þ of 5.0), that have been frozen rapidly in EPR tubes to 77 K, display a signal with an axial g tensor centered on g 2.03, reminiscent of EPR signals observed for monomeric cysteine DNIC species (see Fig. 11.2B) (Cruz-Ramos et al., 2002).

5.2. Reaction of 4Fe-FNR with oxygen Similar experiments to those described earlier can be carried out to investigate the FNR cluster reaction with oxygen following the addition of air-saturated buffer (buffer A or D, see earlier discussion) to anaerobic samples of FNR. The concentration of dissolved oxygen present in the aerobic buffers at the reaction temperature is determined chemically (Vogel, 1989). Briefly, aliquots (100 ml) of the buffer are equilibrated to the reaction temperature in a water bath and then sealed. Dissolved oxygen is fixed by injecting Mn(II)SO4 H2O (2.1 M, 1 ml) below the surface of the sample, followed by 1 ml of a solution of NaOH (40%), NaI (20%), and NaN3 (0.5%). The headspace of the volumetric flask is then purged briefly with oxygen-free nitrogen and the contents are mixed by inversion. After incubation for 15 min, 2 ml of phosphoric acid (85%) is added and the liberated iodine is titrated with a solution of Na2S2O3 5H2O (49.9 mM )

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and a starch indicator (1%). Air-saturated buffers typically contain 220 mM oxygen at 20 . Figure 11.3A shows electronic spectra for 4Fe-FNR before and after oxidation with oxygen. The reaction with oxygen changes the electronic spectrum. The 420-nm shoulder is replaced with a broad shoulder at 430 nm, the absorbance maxima at 310 nm (e ¼ 12,089 M-1 cm-1) and 420 nm (e ¼ 8690 M-1 cm-1) decrease in intensity, and there is an increase in absorbance between 500 and 600 nm (Crack et al., 2004; Jordan et al., 1997; Khoroshilova et al., 1997). This new spectrum indicates the presence of 2Fe-FNR and is characterized by a red/brown color. Stoichiometric experiments carried out as described earlier (and plotting O2:[4Fe-4S]2þ against dA420) show the reaction of 4Fe-FNR with oxygen to be 87% complete at an O2:[4Fe-4S]2þ of 2.0, with the addition of further oxygen only having a minimal effect (Crack et al., 2004). To investigate the rate of reaction of oxygen with 4Fe-FNR, typically a total volume of 2 ml is used. Aerobic and anaerobic buffers are mixed in the cuvette to give a known oxygen concentration, and the cuvette is transferred to a spectrophotometer. The reaction is started by the addition of an aliquot of 4Fe-FNR to the desired concentration. Changes in absorbance spectra during the reactions are monitored at a single wavelength, typically 420 nm, with data points being recorded every 0.5 s. Hence, the conversion of 4Fe-FNR to 2Fe-FNR can be followed. The concentration of 4Fe-FNR for these experiments is variable depending on the ratio of O2: [4Fe-4S]2þ required, but typically it is in the range of 5–30 mM cluster. To calculate the kinetics of the reaction, the decrease in absorbance at 420 nm is plotted. As mentioned earlier, 4Fe-FNR, 2Fe-FNR, and apo-FNR are devoid of an EPR signal. However, when 4Fe-FNR is reacted with stoichiometric amounts of oxygen and frozen rapidly in EPR tubes to 77 K, samples display a signal centered at g 2.024, which behaves in a manner characteristic of [3Fe-4S]1þ clusters (Crack et al., 2004, 2007). Time-resolved appearance of the [3Fe-4S]1þ cluster is observed by reacting 4Fe-FNR with excess oxygen (e.g., a O2:[4Fe-4S]2þ of 10) and rapidly freezing aliquots in EPR tubes throughout the time course (see Fig. 11.3A). Initially, the [3Fe-4S]1þ signal grows in intensity, before decreasing; this behavior is typical of an intermediate, transient species (Crack et al., 2004, 2007). Another useful tool in following the reaction of 4Fe-FNR with oxygen is circular dichroism (CD) spectroscopy. 4Fe-FNR samples are prepared as described previously and reacted with oxygenated buffer, and spectral changes are recorded using Jasco J-810 spectropolarimeter scanning at 200 nm min-1. CD spectroscopy of 4Fe-FNR reveals weak, but reproducible, bands in the 280- to 800-nm region with three positive features (lmax 330, 380, and 420 nm) that, upon oxygen titration, are replaced by a broader spectrum with two positive features (lmax 320 and 440 nm) (see Fig. 11.3C).

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A

35,000 30,000

e M −1 cm−1

25,000 20,000 15,000 10,000 5000 0 260

360


760

860

g = 2.024 g = 2.031 g = 2.019

B 0s 16 s

22 s 45 s 95 s 125 s

3330

3380

3430

3480

3530

Field (gauss) C6 5

4Fe-FNR 4.4 mM O2 6.5 mM O2 10.7 mM O2 21.3 mM O2

4 3 2 1 0 280

330

380

430

480

530

580

630

680

−1 −2

Wavelength (nm)

Figure 11.3 Spectral changes upon reaction of 4Fe-FNRwith oxygen. (A) UV-visible spectra of anaerobic 4Fe-FNR (110 mM ) before treatment with oxygen (thick line). Isolated 2Fe-FNR (80 mM ) generated after treatment of 4Fe-FNRwith oxygen (thin line). (B) EPR spectra of anaerobic 4Fe-FNR (20 mM ) at different times during treatment with oxygen (220 mM ). (C) CD spectra of anaerobic 4Fe-FNR (30.8 mM ) titrated with oxygen (22 mM final). In all cases the buffer used was 25 mM HEPES, 2.5 mM CaCl2,100 mM NaCl, and 100 mM NaNO3, pH 7.5.

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6. Purification of 2Fe-FNR Once produced, 2Fe-FNR is stable for several hours even in the presence of oxygen and can be purified after exposure of 4Fe-FNR to oxygen, as described previously. To obtain sufficient yields, typically a 1-ml solution of 4Fe-FNR in the region of 120 mM [4Fe-4S]2þ is transferred to a glass vial and removed from the anaerobic cabinet. The lid of the vial is removed and 500 ml aerobic buffer is added. The solution is exposed to the air while being agitated gently for a maximum of 90 s. The lid is then replaced and the vial is transferred back to the anaerobic cabinet. The 2Fe-FNR is isolated from cluster breakdown products using a preequilibrated Sephadex G25 PD10 column (GE Healthcare) according to the manufacturer’s instructions, collecting the colored fractions in microcentrifuge tubes.

7. Detection of Other Reaction Products Techniques have been developed or modified to allow the detection of potential products of the reaction of 4Fe-FNR with oxygen, including ferrous ions, sulfide, hydrogen peroxide, and superoxide. Precise identification of these products is important because the nature of the reaction between [4Fe-4S]2þ clusters and oxygen (metal or sulfide based) can be determined by such analyses.

7.1. Detection of sulfide If the oxidation of 4Fe-FNR is sulfide based, then sulfur, rather than sulfide, would most likely be the product of the reaction with oxygen. To test for the production of sulfide during the reaction of 4Fe-FNR with oxygen, a modified version of the protocol reported by Nashef and co-workers (1977) was developed (Crack et al., 2006). Briefly, aliquots of 4Fe-FNR in anaerobic cuvettes (final concentration 5–10 mM) are treated with 200 mM 5,50 dithiobis-(2-nitrobenzoic acid) (DTNB) under anaerobic conditions at room temperature. After 2 min the absorbance is measured at 412 nm. This measurement allows calculation of the amount of reactive thiol in the sample. Oxygen-saturated buffer is then injected to a final oxygen concentration of 40 mM and left to react for 12 min at which point the reaction is complete. The change in absorbance at 412 nm is measured, allowing calculation of the amount of sulfide released in the conversion of 4Fe-FNR to 2Fe-FNR, after correcting the value for the change at 412 nm because of cluster


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conversion (obtained by reacting 4Fe-FNR with oxygen in the absence of DTNB). The reactions are calibrated using a standard sodium sulfide solution, prepared as described by Beinert (1983). The molar extinction coefficients used are: e412 nm ¼ 14,151 M-1 cm-1 in phosphate buffer (buffer A) and e412 nm ¼ 12,344 M-1 cm-1 in HEPES buffer (buffer D).

7.2. Detection of iron The oxidation state of the iron atoms released during the 4Fe-FNR to 2FeFNR conversion is of considerable current interest. Ferene, an Fe(II)specific chelator, has been used to detect iron release from 4Fe-FNR (Crack et al., 2007; Sutton et al., 2004a). It is important to establish that the iron-sulfur cluster under consideration is stable in the presence of ferene. This is the case for FNR in the absence of oxygen. In the presence of Fe(II), the [Fe(II)(ferene)3]4þ complex is formed, which can be quantified using its intense absorbance at 593 nm (e593 nm for [Fe(II)(ferene)3]4þ in phosphatebased buffers is 39,600 M-1 cm-1; e593 nm for [Fe(II)(ferene)3]4þ in HEPESnitrate-based buffers is 32,243 M-1 cm-1). The amount and oxidation state of iron released upon 4Fe-FNR cluster oxidation may be determined by introducing 4Fe-FNR (2 mM final concentration) into phosphate buffer (50 mM potassium phosphate, 400 mM KCl, 10% glycerol, pH 6.8) containing different amounts of oxygen and ferene (100 mM ). Under these conditions, 1Fe(II) is detected for each [4Fe-4S]2þ cluster oxidized (Crack et al., 2007). The total amount of iron released can then be determined by repeating the reaction in the presence of the reductant, sodium ascorbate (4 mM ). This converts any Fe(III) released to Fe(II), which is again detected as a ferene complex. For the oxidation of FNR by oxygen, these experiments suggest that 1Fe(II) and 1Fe(III) are released in the 4Fe-FNR to 2Fe-FNR conversion (Crack et al., 2007). A similar mixture of Fe(II) and Fe(III) has been observed using Mo¨ssbauer spectroscopy in vivo and in vitro (Khoroshilova et al., 1997; Popescu et al., 1998).

7.3. Detection of superoxide and peroxide Two methods have been developed to detect the oxygen species generated during the reaction of 4Fe-FNR with oxygen. To detect superoxide, the method described by McCord and Fridovich (1968) was modified. This method is based on the spectrophotometric detection of cytochrome c reduction by superoxide. Thus, 4Fe-FNR (10 mM ) is injected into a cuvette containing cytochrome c (74 mM, Sigma) in aerobic buffer A (220 mM oxygen, at 21 ). After mixing, the reaction is incubated at room

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temperature. By monitoring changes in absorbance at 550 nm the reduction of cytochrome c is monitored (De550 nm ¼ 21,000 M-1 cm-1). To show that the reduction of cytochrome c is caused by superoxide, control reactions containing superoxide dismutase (16 units) are required (Crack et al., 2007). Superoxide can dismutate spontaneously to form hydrogen peroxide, which will reoxidize reduced cytochrome c. Therefore, reactions containing catalase (234 units) are also informative. For FNR oxidation, the presence of catalase increased the amount of superoxide detected, indicating that some hydrogen peroxide is generated during the oxidation of 4FeFNR (Crack et al., 2007). Indeed, hydrogen peroxide can be detected in a coupled reaction using the fluorescent probe Amplex Red (Molecular Probes). A solution of Amplex Red (400 mM ) containing 2 units of horseradish peroxidase is prepared in FNR purification buffer. One milliliter of 4Fe-FNR (60 mM ) is added to 1 ml of Amplex Red solution in a standard fluorescence cuvette under anaerobic conditions. The sealed cuvette is transferred to a Perkin Elmer LS55 spectrofluorimeter and titrated with oxygen by injection of aliquots of aerobic purification buffer. Fluorescence spectra are recorded using an excitation wavelength of 545 nm (5-nm slit width) and an emission slit width of 2 nm. Hydrogen peroxide is detected by an emission maximum at 587 nm and can be quantified by calibrating the reactions by the addition of an aliquot of a standard hydrogen peroxide solution into the reaction mixture at the end of titration (Crack et al., 2004).

8. Conclusions Using the methods described in this chapter, the reactions of FNR with oxygen and NO have been investigated. The application of these techniques has provided new insight into FNR [4Fe-4S] cluster chemistry, suggesting the following reactions with NO [Eq. (11.1)] and oxygen [Eq. (11.2)]:

½4Fe 4S2þ þ 4NO ! 2FeðNOÞ2 þ 2Fe3þ þ 4S2 ½4Fe 4S2þ þ O2 ! ½2Fe 2S2þ þ Fe2þ þ Fe3þ þ 2S2 þ O 2

ð11:1Þ ð11:2Þ

It should be appreciated that quantitative experiments to investigate the reactions of iron-sulfur clusters with oxidants are technically demanding and chemically complex, and despite the progress that has been made there are many outstanding questions, which will require the development of new methodologies before a deeper understanding of these processes is achieved.


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C H A P T E R

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Genome-Wide Identification of Binding Sites for the Nitric Oxide-Sensitive Transcriptional Regulator NsrR Sam Efromovich,* David Grainger,† Diane Bodenmiller,‡ and Stephen Spiro§ Contents 1. Introduction 2. Strain Construction 3. Reference and Control Samples 4. Culture Conditions 5. Immunoprecipitation of DNA Targets Associated with NsrR 6. DNA Labeling, Microarray Hybridization, and Processing 7. Visualization and Analysis of DNA Microarray Data 8. A New Statistical Methodology for Treatment of ChIP-on-Chip Data 9. Conclusions Acknowledgments References

212 214 216 217 218 219 220 222 231 231 231

Abstract NsrR is a nitric oxide-sensitive regulator of transcription. In Escherichia coli, NsrR is a repressor of the hmp gene encoding the flavohemoglobin that detoxifies nitric oxide. Three other transcription units ( ytfE, ygbA, and hcp-hcr) are known to be subject to regulation by NsrR. This chapter describes experimental and statistical protocols used to identify NsrR-binding sites in the E. coli chromosome using chromatin immunoprecipitation and microarray analysis. The methods are applicable, with suitable modifications, to any regulatory protein and any organism.

* Department of Mathematical Sciences, University of Texas at Dallas, Richardson, Texas School of Biosciences, University of Birmingham, Edgbaston, Birmingham, United Kingdom School of Biology, Georgia Institute of Technology, Atlanta, Georgia } Department of Molecular and Cell Biology, University of Texas at Dallas, Richardson, Texas { {


#


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Sam Efromovich et al.

1. Introduction Escherichia coli uses nitrate and nitrite as terminal electron acceptors for anaerobic respiration, reducing nitrate to nitrite, and nitrite to ammonia. There is good evidence to indicate that low concentrations of nitric oxide (NO) are made as a by-product of this respiratory metabolism (Corker and Poole, 2003; Ji and Hollocher, 1988; Van Doorslaer et al., 2003). This endogenous formation of NO provides a physiological rationale for the expression of enzymes that reduce or oxidize NO to less toxic species, and regulatory proteins that sense NO and mediate transcriptional responses to NO exposure. In E. coli, three enzymes have established roles in detoxifying NO: the flavohemoglobin (Hmp) oxidizes NO to nitrate (in the presence of oxygen) and reduces NO to nitrous oxide (Poole and Hughes, 2000); the flavorubredoxin (FlRd) reduces NO to nitrous oxide (Gardner et al., 2002; Gomes et al., 2002); and the respiratory nitrite reductase (Nrf ) reduces NO to ammonia (Poock et al., 2002). The same enzymes may allow pathogenic strains of E. coli and related enteric bacteria to detoxify the NO made by host cells (Bang et al., 2006; Sebbane et al., 2006; Stevanin et al., 2002). A number of E. coli regulatory proteins have been reported to be sensitive to NO and to control gene expression in response to NO, including SoxR, OxyR, FNR, Fur, and MetR (Spiro, 2007). However, the most important regulators (in terms of providing physiological regulation of expression of genes encoding enzymes that detoxify NO) are, probably, NsrR and NorR (Spiro, 2007). NorR activates expression of the genes encoding the flavorubredoxin and its reductase in response to NO (D’Autreáux et al., 2005; Gardner et al., 2003). NsrR is a recently described repressor protein, which is sensitive to NO (or possibly another N compound) and controls expression of, among others, genes encoding Hmp and Nrf (Bodenmiller and Spiro, 2006; Spiro, 2007). This chapter describes methods used to identify NsrR-binding sites in the E. coli genome. A recent study made use of comparative genomics to reconstruct the transcriptional networks that respond to nitric oxide (and other nitrogen oxides and oxyanions) in a wide range of prokaryotic species (Rodionov et al., 2005). In the case of E. coli, this study predicted that the product of the nsrR (formerly yjeB) gene is a transcriptional regulator of four transcription units: hmp, ytfE, ygbA, and hcp-hcr (Rodionov et al., 2005). Of these, the products of the hmp and ytfE genes have established roles in protecting against NO ( Justino et al., 2005, 2006; Poole, 2005). It has been confirmed experimentally that NsrR is a repressor of hmp, ytfE, and ygbA and that repression is relieved in the presence of physiological or chemical sources of NO (Bodenmiller and Spiro, 2006). We have subsequently also confirmed

The NsrR Regulon of Escherichia coli

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(Filenko et al., 2007) that NsrR is a repressor of the hcp-hcr genes (encoding the hybrid cluster protein and its redox partner) and the nrf genes encoding the respiratory periplasmic nitrite reductase, which also has NO reductase activity (Poock et al., 2002). Thus, it seems that NsrR mediates widespread changes in the E. coli transcriptome in response to NO. It is of considerable interest to identify the full extent of the NsrR regulon, in other words, to identify all genes subject to regulation by NsrR. To this end, we have employed chromatin immunoprecipitation with microarray analysis (ChIPon-chip) to identify binding sites for NsrR in the E. coli genome. This chapter describes the techniques involved, which are applicable, with some modifications, to any regulatory protein (given a suitable antibody) and to any organism for which a high-density microarray is available. Chromatin immunoprecipitation is an in vivo technique used to monitor interactions between DNA-binding proteins and their targets. Proteins are cross-linked to DNA in vivo (typically by treatment with formaldehyde); after cell disruption the cross-linked chromatin is sheared randomly by sonication or enzymatic digestion. Protein–DNA complexes are selectively immunoprecipitated with a suitable antibody, the cross-linking is reversed, and the precipitated DNA is purified. The polymerase chain reaction (PCR) can then be used with specific primers to test whether candidate target sequences are enriched in the precipitated DNA population. In ChIP-on-chip, the precipitated DNA is (in some protocols) amplified and is suitably labeled for hybridization to a microarray. Since the precipitated DNA is typically in the range of 500–1000 bp, a conventional array with one probe per gene can be used. In this case the outcome is that probes within 500–1000 bp of the protein-binding site give a positive signal, allowing the experimenter to associate the site with one or two genes with a reasonable level of confidence (Molle et al., 2003). A modification of this approach involves the use of high-density microarrays, with short gaps between probes, which correspond to both coding and noncoding regions. This improves the resolution of data and allows signals to be associated unambiguously with a single transcription unit or with two divergently oriented transcription units. General considerations for the design and interpretation of ChIP-on-chip experiments have been reviewed elsewhere (Buck and Lieb, 2004). Chromatin immunoprecipitation and ChIP-on-chip methods were developed in eukaryotic systems, but have more recently been applied successfully in prokaryotes, for example, Bacillus subtilis (Molle et al., 2003; Rokop et al., 2004), Salmonella enterica (Shin and Groisman, 2005), and E. coli (Grainger et al., 2004, 2005). ChIP-on-chip has several advantages and disadvantages compared to other methods used to define regulons, for example, the use of microarrays to compare the transcriptomes of wild-type and mutant strains. Genes that

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are subject to indirect regulation by NsrR will show NsrR-dependent responses to NO in a microarray experiment, but be negative in a ChIPon-chip experiment. Thus, the latter approach identifies candidate genes whose transcription is directly regulated by NsrR. This is both a strength and a weakness of the approach, depending on the investigator’s priorities. ChIP-on-chip requires no prior knowledge about other regulatory mechanisms that may impinge upon genes controlled by NsrR. For example, if expression of an NsrR-regulated gene is also dependent on another factor, then that gene may not be identified in a transcriptomics experiment, unless the appropriate growth conditions are used. ChIP-on-chip does identify binding sites associated with genes expressed at a low level, and where the regulatory protein may have little effect (Grainger et al., 2006). This is a potential disadvantage of the approach if binding at such sites has no biological significance. A major limitation of ChIP-on-chip is that some binding sites known from other approaches are not identified, in other words there is a significant false-negative rate (Grainger et al., 2006). Importantly, ChIP-on-chip only identifies factor-binding sites, demonstrating regulation of transcription occurring at those sites requires confirmation by an independent technique. Further, where binding sites are located in noncoding regions between divergently transcribed genes, ChIP-on-chip data cannot distinguish which gene (if either) is subject to regulation. An overview of the procedure is illustrated in Fig. 12.1 and a detailed protocol is presented here.

2. Strain Construction A requirement for ChIP-on-chip is the availability of an antibody for the selective precipitation of cross-linked protein–DNA complexes. In the case of NsrR, specific anti-NsrR antibodies are not yet available. Instead, we chose to epitope tag NsrR by introduction of the 22 amino acid 3 Flag tag at the C terminus of the protein. Introduction of this construct onto the chromosome avoids any artifacts associated with the overexpression that may result from the use of plasmid-borne constructs. The epitope tag is introduced by a PCR-based method with plasmid pSUB11 as the template (Uzzau et al., 2001). The tagged construct is then introduced onto the chromosome of E. coli strain BW25113 using the lRed recombinase method (Datsenko and Wanner, 2000). At each stage, DNA and strain constructions are confirmed by PCR and/or sequencing. This approach results in the introduction of a kanamycin resistance cassette in the chromosome downstream of the tagged gene. The cassette can be removed by recombination (Datsenko and Wanner, 2000), although this was not done

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Figure 12.1 Schematic of the ChIP-on-chip protocol. Two anaerobic cultures are grown of a strain expressing NsrR with a C-terminal epitope tag. Because growth in the presence of nitrate causes derepression of NsrR targets (Bodenmiller and Spiro, 2006), the occupancy of NsrR binding sites is lower under these conditions. Step1: nucleoprotein is cross-linked by treatment with formaldehyde. Step 2: chromatin is extracted and randomly sheared by sonication to produce fragments in the 500- to 1000-bp range. Step 3: NsrR:DNA complexes are selectively immunoprecipitated with an antibody raised against the epitope tag. Step 4: cross-linking is reversed and DNA is purified and fluorescently labeled using the Klenow fragment of DNA polymerase and random hexanucleotide primers. Step 5: labeled DNA samples are hybridized to a high-density microarray.

for the experiments described later. The cassette also provides a selectable marker allowing the epitope-tagged gene to be transferred to other strains by phage P1 transduction. It is important to confirm that the epitope tag

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does not interfere with the normal activity of the regulatory protein. To do this, we transfer the construct into strain JOEY19, which contains a ytfElacZ reporter fusion (Bodenmiller and Spiro, 2006), and confirm that the epitope-tagged NsrR exerts NO-sensitive repression of the ytfE promoter in a manner indistinguishable from the wild-type protein. The tagged nsrR gene is also transferred into the wild-type strain MG1655 by transduction to generate strain JOEY135, which is used for the ChIP-on-chip experiments. The nsrR gene is cotranscribed with rnr, encoding ribonuclease R (Cairrao et al., 2003). The epitope-tagging procedure described earlier results in the introduction of a kanamycin resistance cassette between nsrR and rnr, which may perturb the level of expression of rnr by polarity. While this would be a concern for a transcriptomics experiment, we believe that it is not an issue for the ChIP-on-chip experiment described here because (a) an altered activity of RNase R is unlikely to have any consequence for the ability of NsrR to bind to its target sequences and (b) the test and control strains used in the protocol described later are genetically identical; they both contain the same construct used to epitope tag NsrR.

3. Reference and Control Samples For ChIP-on-chip experiments, we employed a dual-labeling protocol (Grainger et al., 2005), in which the immunoprecipitated DNA (later referred to as the experimental sample) is labeled with Cy5 (or Cy3) and a reference DNA is labeled with Cy3 (or Cy5). The two samples are then hybridized to the array simultaneously, and the fluorescence ratio at each probe is measured. Thus, each ChIP-on-chip experiment requires a hybridization reference DNA. Ideally, this should be the same for each experiment to allow comparisons to be made between experiments (Buck and Lieb, 2004). For example, a sample of genomic DNA taken after the sonication step and prior to immunoprecipitation can be used as the reference (Molle et al., 2003). The control sample is designed to detect nonbiological sources of variation. Possible sources of the control DNA are (1) DNA from a ‘‘dummy’’ immunoprecipitation reaction in which no antibody is present, (2) DNA immunoprecipitated from a control strain in which the regulatory protein is not epitope tagged, and (3) DNA immunoprecipitated from the epitope-tagged strain grown under conditions where the regulatory protein does not bind to DNA. In practice, criteria for an ideal experiment may not be met for technical reasons. For example, we have found that chromatin immunoprecipitation from strain MG1655 (in which NsrR is not epitope tagged) with the commercially available anti-Flag monoclonal antibody yielded insufficient DNA to use as a control in the ChIP-on-chip protocol.


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Therefore, we prepared the control DNA from a culture of strain JOEY135 grown under conditions leading to derepression of NsrR targets, in other words, conditions where NsrR-binding sites are not fully occupied. This DNA was used as the reference in array hybridizations in which the experimental DNA came from a culture grown in the absence of nitrate. It has been argued that the control sample should not be used as the hybridization reference, as the ‘‘perfect’’ control experiment would not contain any DNA (Buck and Lieb, 2004). However, the approach described here has the advantage of increasing the information content of a single experiment in that where sites are identified we can be certain that binding at those sites is regulated (in this case by nitrate). Indeed, it is typical in published experiments for a control DNA to be used as the hybridization reference (Grainger et al., 2006), which is the approach recommended in protocols provided by Oxford Gene Technology (OGT)(www.ogt.co.uk).

4. Culture Conditions It is likely that the effector for NsrR is NO, either supplied exogenously or made endogenously as a by-product of the respiratory metabolism of nitrate and nitrite (Bodenmiller and Spiro, 2006; Spiro, 2007). Thus, preparation of the control DNA requires growth of a culture in the presence of NO. Experimentally, NO can be provided as a bolus of aqueous solution, or in the gas phase, but this makes the concentration difficult to control over time, as cultures of E. coli consume NO rapidly, and NO is susceptible to autooxidation, especially in the presence of oxygen. Compounds that release NO with known kinetics are also useful experimental tools. For the experiments described here, we instead chose to use a physiological source of NO, as it is known that E. coli makes NO endogenously as a by-product of the respiration of nitrate and nitrite (Corker and Poole, 2003; Van Doorslaer et al., 2003) and the NsrR regulon is derepressed in anaerobic cultures respiring nitrate or nitrite (Bodenmiller and Spiro, 2006). Because nitrite is toxic, for studies of gene expression we typically add it to cultures at a low concentration (5 mM ) shortly before samples are taken. Nitrate is not toxic and can therefore be present in cultures throughout growth in excess at high millimolar concentrations. For the purposes of this experiment, it is not necessary to know the concentration of NO made during nitrate and nitrite respiration. We assume that NO is made continuously during the logarithmic phase of growth in the presence of excess nitrate and know from studies of gene expression that both NorR and NsrR are sufficiently sensitive to respond to the NO made under these conditions. For the ChIP-on-chip experiment, the control culture of JOEY135 is grown anaerobically (in filled bottles) in 200 ml of L-broth supplemented with 0.5% (w/v) glucose and 50 mM potassium nitrate.

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The experimental culture is grown under identical conditions, except that nitrate is omitted from the medium. In late logarithmic phase (OD650 nm 0.75–1.0), formaldehyde is added to cultures (final concentration 1%), which are then incubated for another 20 min. To quench the cross-linking, glycine is added to a final concentration of 500 mM, and cultures are incubated for a further 5 min.

5. Immunoprecipitation of DNA Targets Associated with NsrR The following protocols for chromatin immunoprecipitation and DNA labeling and hybridization are based on published methods (Grainger et al., 2005) with some minor modifications (e.g., volumes are scaled up to account for the lower yield of anaerobic cultures). 1. Harvest 200 ml culture by centrifugation and wash twice in 10 ml of Tris-buffered saline (TBS), pH 7.5. After the second wash, resuspend cells in 0.5 ml of lysis buffer (10 mM Tris-HCl, pH 8.0, 20% sucrose, 50 mM NaCl, 10 mM EDTA, 10 mg/ml lysozyme) and incubate for 30 min at 37 . A stock of lysis buffer without lysozyme can be prepared in advance and supplemented with lysozyme immediately prior to use. 2. After treatment with lysozome, cell lysis is completed by the addition of 2 ml of immunoprecipitation buffer (50 mM HEPES-KOH, pH 7.5, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100, 0.1% sodium deoxycholate, 0.1% SDS). To minimize protein degradation, phenylmethylsulfonyl fluoride is added to a final concentration of 1 mM and the sample is chilled on ice in preparation for sonication. 3. Sonication is used to fragment randomly the nucleoprotein content of the sample, with the goal being to obtain DNA fragments in the 500 to 1000 bp range. The sonication protocol depends on the type of sonicator and the amount of biomass used to generate the sample and must be determined empirically by electrophoresis of samples taken at various intervals. For 200 ml late log phase anaerobic cultures prepared as described earlier, we found that eight 15 s treatments, with 30 s intervals for chilling, with a Branson S150D sonicator on the 7.5 power setting was sufficient to generate DNA fragments 500–1000 bp in length. 4. Remove cell debris from the sample by centrifugation. 5. Wash 25 ml of Ultralink protein A/G beads (Pierce) three times with TBS prior to use. After the final wash, remove as much of the liquid phase as possible. To immunoprecipitate cross-linked NsrR-3FlagDNA complexes, add an 800 ml aliquot of the sonicated chromatin to the washed protein A/G beads and then 5 ml of monoclonal anti-Flag antibody (Sigma). Mix overnight on a rotating wheel at 4 .


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6. After immunoprecipitation, the protein A/G beads can be collected and washed in Spin-X columns supplied by Corning Life Sciences (Grainger et al., 2005). We have found a convenient and effective alternative to be harvesting and washing by centrifugation in a plastic microcentrifuge tube, and resuspension of the pellet in the wash buffers. In this way, the beads are washed twice in 1 ml immunoprecipitation buffer, once in 1 ml of immunoprecipitation buffer supplemented with 500 mM NaCl, once with 1 ml wash buffer (10 mM Tris-HCl, pH 8.0, 250 mM LiCl, 1 mM EDTA, 0.5% Nonidet P-40, 0.5% Na deoxycholate), and once with 1 ml Tris-EDTA, pH 7.5. 7. Transfer the beads in TE to a 2 ml screw cap tube, centrifuge, and resuspend in 50 ml of elution buffer (50 mM Tris-HCl, pH 7.5, 10 mM EDTA, 1% SDS) and incubate the sample at 65 for 10 min and then at 42 for 2 h, without agitation. 8. Add 0.8 mg pronase (made up in water or TBS) per milliliter of sample and incubate overnight at 65 . Pronase is added in sufficient water to dilute the sample twofold, so this step is done in 0.5 elution buffer. Centrifuge the sample and transfer the supernatant to a clean tube. 9. Purify the DNA fragments using a Qiagen PCR purification kit according to the manufacturer’s protocol. Elute the DNA from the Qiagen column in 30 ml of water and determine the DNA concentration (a Nanodrop is useful for this measurement). A single immunoprecipitation usually yields about 100 ng of DNA.

6. DNA Labeling, Microarray Hybridization, and Processing The high-density E. coli MG1655 microarrays fabricated and supplied by Oxford Gene Technology have been described previously (Grainger et al., 2005). Briefly, the arrays comprise 21,321 60-base-long oligonucleotides, with an average spacing between them of 160 bp. The probes are not in genome order on the array, which provides a control against position effects. For our analysis, we labeled DNA immunoprecipitated from each culture with Cy5 and Cy3 and compared the two samples by simultaneous hybridization to the array. Note that there is no amplification of the precipitated DNA in this protocol, which avoids possible artifacts because of amplification bias. 1. Before beginning, DNA samples should be in a volume of 20 ml and at a minimum concentration of 5 mg/ml. Mix the DNA with 20 ml of 2.5 random primer (Bioprime kit, Invitrogen) and 0.25 ml of sterile water and denature by heating to 94 for 3 min.

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2. After denaturation, add 5 ml of dNTP mix (2 mM dATP, 2 mM dGTP, 2 mM dTTP, 0.5 mM dCTP), 3.75 ml of Cy5- or Cy3-labeled dCTP (1 mM; GE Healthcare), and 1 ml of the Klenow fragment of DNA polymerase (Bioprime kit), mix gently, and incubate at 37 for 2 h. 3. Add a further 0.5 ml of DNA polymerase to each tube and incubate at 37 for a further 2 h. 4. Purify DNA fragments from the labeling reaction using QIAquick PCR purification columns (Qiagen), according to the manufacturer’s instructions. The labeled DNA should be eluted from the column using 50 ml of Qiagen elution buffer. 5. The labeled DNA samples can now be mixed and hybridized to the array according to the manufacturer’s instructions (freely available at www. ogt.co.uk). Oxford Gene Technology arrays are manufactured on an Agilent platform, and hybridizations are best performed in an Agilent SureHyb apparatus.

7. Visualization and Analysis of DNA Microarray Data After scanning the microarray, a list of Cy5 and Cy3 signal intensities is generated. The fluorescence intensity ratio is then calculated for each probe on the array and plotted against the genomic coordinate of the probe, generating a genome-wide DNA-binding profile for NsrR. This analysis can be done in a Microsoft Excel spreadsheet, and binding sites are localized approximately by comparison of peak heights with a suitable E. coli database. Alternatively, the profile can be scrutinized using a ‘‘genome browser,’’ in which the fluorescence ratios are superimposed on the genetic map of E. coli. Oxford Gene Technology’s ChIP Browser software (www.ogt.co.uk) facilitates this type of analysis. Our ChIP-on-chip data for NsrR binding to the ytfE promoter from three repeat experiments are shown as an example in Fig. 12.2A. In typical ChIP-on-chip data, several genetically adjacent probes register a fluorescence ratio that is significantly above the background (see Fig. 12.2A). This feature in data is usually referred to as a ‘‘peak’’ (although, strictly speaking, the peak is the single probe with the highest fluorescence ratio). In what follows, we use ‘‘peak’’ in the conventional manner to refer to a series of probes with an above-background fluorescence ratio. The peak should be centered over the genome location of the protein-binding site, and its width and shape depend on the size of DNA fragments after sonication and labeling (Buck et al., 2005). In many cases, these peaks can simply be identified by visual inspection of data. If required, a cutoff (which is necessarily arbitrary) in the fluorescence ratio can be

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Figure 12.2 Strong and weak signals in raw ChIP-on-chip data. (A) ChIP-on-chip data for a 2-kb window around the NsrR-regulated ytfE promoter (Bodenmiller and Spiro, 2006). The fluorescence ratios are from immunoprecipitated cross-linked DNA extracted from cells grown in the absence and presence of nitrate. Raw data from three experiments are shown. Each individual dataset was centered on a ratio of 1 (by subtraction of the mean fluorescence ratio for that dataset þ 1); this is a necessary prerequisite to subsequent analysis with ChIPOTle (Buck et al., 2005). The probe giving the highest fluorescence ratio is at coordinates 4429599^4429659; the ytfE start codon is at 4429561, and the predicted NsrR-binding site (Bodenmiller and Spiro, 2006) is centered on 4429598.5. The exact coincidence of the peak maximum with the NsrR-binding site in this particular case is a consequence of the fortuitous fact that the probe on the array overlaps the binding site. More typically, peaks in raw data may be up to 100 bp away

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applied, and peaks assigned if two or more adjacent probes exceed the cutoff. This can be a laborious process in large data sets with many protein-binding sites, and some data sets are more challenging, especially if there are peaks that represent real binding sites, but do not clearly rise above the background noise. This issue can be illustrated by our ChIP-onchip data for the ygbA promoter (see Fig. 12.2B). This promoter is known to be regulated by NsrR and contains an NsrR-binding site (Bodenmiller and Spiro, 2006), but gives a weak signal in data that may, or may not, be discarded if judged by purely subjective criteria (see Fig. 12.2B). The identification of peaks can be addressed computationally, for example, by using chromatin immunoprecipitation on tiled arrays (ChIPOTle), a freely available and easily applied Microsoft Excel macro (Buck et al., 2005). ChIPOTle analyzes ChIP-on-chip data using a sliding window and reports the locations of statistically significant peaks. The user sets parameters for the window and step sizes and for the P value cutoff. This approach allows rapid identification of all significant peaks in a large data set and assigns a quantitative confidence measure (a P value) to each peak (Buck et al., 2005). This valuable approach allows rapid identification of peaks in large data sets. However, some real binding sites may fail to exceed the statistical cutoff and therefore fail to be reported. Relaxation of search criteria may help resolve this problem, but will also increase the rate of false positives. In the case of ygbA, ChIPOTle finds a significant peak (P ¼ 0.0001) in two of the three data sets. The ygbA promoter would not be reported as positive if an arbitrary cutoff were employed of two or more adjacent probes showing greater than twofold enrichment in at least two data sets (see Fig. 12.2B).

8. A New Statistical Methodology for Treatment of ChIP-on-Chip Data In ChIP-on-chip experiments, each probe on the array measures the abundance of a population of DNA fragments that differ in length as a consequence of sonication and labeling by random priming. As a result, from the binding site. Note that the coordinates assigned to probes by Oxford GeneTechnology refer to a non-current annotation of the E. coli genome (http://genolist.pasteur. fr/Colibri/). A gray line is plotted at the two-fold enrichment level. (B) ChIP-on-chip data for a 2-kb window around the NsrR-regulated ygbA promoter (Bodenmiller and Spiro, 2006). Data are averaged from the same three experiments, shown in a, and were manipulated in the same way.The peak in the noncoding region between ygbA and mutS is statistically significant (P ¼ 0.0001) in two datasets, when data are analyzed with ChIPOTle (Buck et al., 2005). Note that this result would not be reported as a positive if, as the cutoff, two or more adjacent probes are required to show greater than twofold enrichment in the immunoprecipitated population in two of the three experiments.


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several probes close to the protein-binding site will give a relatively large positive signal. Depending on the number and location of probes near a binding site, and the distribution of labeled fragment lengths, a specific profile of intensities of signals for genetically adjacent probes is created. In general (and always for equally spaced probes) there should also be a maximum enrichment for the probe closest to the binding site. A ‘‘peak’’ in data is therefore characterized by a relatively large signal for several adjacent probes (see Fig. 12.2A). The second feature of ChIP-on-chip data is that the measurements are a mixture of two distributions, with the larger one being a background noise and the smaller being of larger fluorescence ratios. These larger ratios cannot be explained by background distribution and thus are positive outliers from background distribution. In a traditional transcriptomics experiment, the distribution of signals is two tailed and mostly symmetric; both low- and high-intensity signals are potentially of interest. In ChIP-on-chip data, only the larger signals are of interest because they correspond to the population of genomic fragments specifically enriched by the ChIP. Because of such an asymmetric distribution of signals with an extremely heavy right tail, it is often recommended to analyze a logarithmically transformed signal. Typically a log with base 2 is used and applied to the ratio of experimental to reference signals (Buck et al., 2005). The main statistical issue frequently considered in ChIP-on-chip literature is how to find peaks in (log-transformed) fluorescence ratios. To this end, several different methods and software have been published. In the aforementioned ChIPOTle, moving windows are used to smooth data and then to find peaks as local points of maximum (Buck et al., 2005). Then, an algorithm calculates P values for proposed peaks based on the standard normal error function; a conservative Bonferroni correction may complement the procedure. Another approach is to use a percentile rank analysis (Bieda et al., 2006). For a more precise estimation of binding-site locations, a ‘‘triangle’’ peak-finding model has been proposed (Kim et al., 2005). Given the 100 bp resolution of the array, the authors (with some assumptions) derived a model that states that the log ratio should decrease linearly with distance from the true binding site. Then model fitting is done by traditional linear regression. This method of peak finding is interesting because it seeks to estimate the binding-site locations underlying each peak. However, the method has been criticized on the basis that profiles of log ratios near a peak varied greatly in their waveform, amplitude, and size, thus making the rigid triangle profile not feasible (Bieda et al., 2006). Instead, an approach for peak finding that makes minimal assumptions about the shape and amplitude of the log-ratios profile is recommended (Bieda et al., 2006). To this end, the authors suggested the use of a set of percentile thresholds (specifically the 95th and 98th percentiles), and then analysis of a peak is conducted via a

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pattern of indicator functions that a log ratio exceeds the threshold (Bieda et al., 2006). The underlying probability theory of such an approach is based on the theory of random runs. Here, we propose a novel approach to the estimation of binding site locations from ChIP-on-chip data, and the following considerations underlie the proposed methodology. An important feature of OGT microarrays is the relatively large (on average 160 bp) and highly irregular distances between the probes. The probe that gives the highest fluorescence ratio may thus be 100 bp (or, in some cases, considerably more) distant from the protein binding site. We have developed a statistical approach to estimate the locations of protein binding sites from ChIP-on-chip data obtained from OGT (and similar) microarrays, where probes are relatively sparse and irregularly spaced. To locate a binding site, a probabilistic model for prediction of the shape of fluorescence ratio data between experimental and reference signals for a particular configuration of neighboring probes is developed. This model assumes (i) a particular probability distribution of DNA fragment lengths after sonication and labeling by random priming; (ii) that for a given immunoprecipitated DNA fragment, the probability of a binding site occurring anywhere within that fragment is equal and independent of the fragment length; (iii) DNA fragments will hybridize to probes if the two overlap for at least half the length of the probe (30 bp); (iv) a given DNA fragment has equal chances of hybridizing to those probes that it overlaps; (v) the probability that a DNA fragment will hybridize to a particular probe is the same for reference and experimental samples, but may differ from probe to probe; and (vi) each hybridized DNA fragment adds a fluorescence signal intensity that is proportional to its length, and the coefficient of proportionality is the same for all fragments and probes. The model predicts a profile of fluorescence intensity ratios for probes surrounding a conjectured binding site. The conjectured site is then moved along the chromosome in the region of a large positive fluorescence signal, and for each conjectured site the statistical performance of the model is evaluated via empirical standard deviations of residuals (differences between observed signals and those predicted by the model). Because the standard deviation of residuals depends on the location of the binding site, the estimate of the true site location is that which minimizes the standard deviation of residuals. If several experiments are conducted, then binding sites are calculated for each experiment and then reported together with corresponding standard deviations; the reasoning behind this approach is explained later. The method of statistical visualization and analysis we have developed consists of three major steps. Step 1 is a preliminary step, which allows one to visualize the experimental and reference signals and their ratio for each experiment (in our case 3).

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The procedure is illustrated for a 65.5 kb window around the hcp gene in Fig. 12.3 The hcp promoter contains an NsrR binding site (Rodionov et al., 2005), and we have confirmed that NsrR is a repressor of hcp transcription (Filenko et al., 2007). A pronounced peak is visible in denoised fluorescence data for the reference sample (see Fig. 12.3B). All other potential peaks in this region result from noise and thus the procedure shrinks them to zero. The procedure used is based on data-driven multiwavelet Efromovich block-shrinkage filtering (Efromovich, 2001; Efromovich et al., 2004). Raw and denoised data for the experimental sample are shown in Figs. 12.3C and 12.3D. The denoised experimental signal exhibits two peaks: one corresponds to the one observed in the denoised reference signal (which is therefore a candidate for being technical artifact) and the second peak localizes to the hcp promoter region. In denoised fluorescence ratio data (see Fig. 12.3F ) the only prominent signal results from NsrR binding to

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Figure 12.3 Observed and denoised signals for the first of three ChIP-on-chip experiments. A 65.5-kb region of the E. coli chromosome is shown. Raw (A, C, and E) and denoised (B, D, and F) fluorescence data are shown for reference (A and B) and experimental samples (C and D), and for the ratios (E and F ).The peak at 9.14 105 bp in fluorescence ratio data is in the promoter region of the hcp gene.

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a site in the vicinity of the hcp promoter. The two weaker signals in the fluorescence ratios should be explored with extra statistical precaution, and by visual inspection, because they are candidates for false positives, or may represent true binding sites. The developed method of a multiwavelet signal denoising allows one to visualize peaks in the ratio signal and to identify possible false positives. Thus, in the first step of the process, it is useful to examine reference and experimental fluorescence data separately. Step 2 is devoted to another preliminary analysis of data, which allows one to obtain a preliminary estimate of the possible number of binding sites, as well as to check the assumption that the observed background ratios (because of pure noise or technical artifacts) have a normal distribution. This is done by visualization of the histogram of z-scored ratios (Fig. 12.4). Obtaining z-scored ratios for a particular experiment is based on the following procedure. First, a sample median is calculated. Then a robust standard deviation estimator is calculated as 1.48 times the median of absolute deviations from the median in a procedure called mad (Efromovich, 1999). Finally, a classical z scoring is performed. Histograms in the left panels of Fig. 12.4 show z-scored ratios that do not exceed a threshold T ¼ 4.89. This threshold is chosen to satisfy the relation P(max(Z1,. . .,Zn) >T)

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Methods in Enzymology 492 - Biothermodynamics, Part 4

Methods in Enzymology 455 - Biothermodynamics, Part 1

Vitamins & Coenzymes, Part L, Volume 282 (Methods in Enzymology)

Methods in Enzymology Vol 327: Applications of Chimeric Genes and Hybrid Proteins, Part B: Cell Biology and Physiology

Globins and Other Nitric Oxide-Reactive Proteins, Part B (Methods in Enzymology, Volume 437)